Bryophyte Ecology (ver Jul 2020) [2]

Table of contents :
Bryophyte Ecology About the Author
Bryophyte Ecology About the Book
About the Book
The Format
Acknowledgments
Bryophyte Ecology Glossary
Literature Cited
Bryophyte Ecology Table of Contents
Chapter 1 - The Fauna A Place to Call Home
Chapter 2 - Protozoa
2-1Protozoa Diversity
2-2Protozoa Ciliophora & Heliozoa
2-3Rhizopod Diversity
2-4Rhizopod Ecology
2-5Peatland Rhizopods
2-6Protozoa Ecology
Chapter 3 - Slime Molds
3-1Slime Molds - Biology and Diversity
3-2Slime Molds - Bryophyte Associations
3-3Slime Molds - Ecology and Habitats - Bark and Logs
3-4Slime Molds - Ecology and Habitats - Lesser Habitats
Chapter 4 - Invertebrates
4-1Invertebrates Introduction
4-2Invertebrates Sponges Gastrotrichs Flatworms
4-3Invertebrates Nematodes
4-4Invertebrates Annelids
4-5Invertebrates Rotifers
4-6Invertebrates Rotifer Taxa Bdelloidea
4-7aInvertebrates Rotifer Taxa Monogonata 1
4-7bInvertebrates Rotifer Taxa Monogonata 2
4-7cInvertebrates Rotifer Taxa Monogonata 3
4-8Invertebrates Mollusks copy
Chapter 5 - Tardigrades
5-1Tardigrades Survival
5-2Tardigrades Reproduction & Food
5-3 Tardigrade Habitats
5-4Tardigrades Species Relationships
5-5Tardigrade Densities and Richness
5-6Tardigrade Ecology
Chapter 6 - Onychophora
Chapter 7 - Arthropods Spiders
7-1Arthropods Habitat Relations
7-2Arthropods Spider Biology
7-3Arthropods Spider Habitats
7-4Arthropods Peatland Spiders
7-5Arthropods spiders in Danish and tundra peatlands
7-6Table 2 Species List
Chapter 8 - Arthropods Harvestmen and Pseudoscorpions
Chapter 9 - Arthropods Mites
9-1Arthropods Mites
9-2Arthropods Mite Habitats
Chapter 10 - Arthropods Crustacea
10-1Arthropods Crustacea - Copepoda and Cladocera
10-2Arthropods Crustacea - Ostracods and Amphipods
10-3Arthropods Crustacea - Isopoda, Mysida, and Decapoda
Chapter 11 - Aquatic Insects
11-1Aquatic Insects Biology
11-2Aquatic Insects - Bryopyte Roles as Insect Habitats
11-3Aquatic Insects Bryopyte Habitat and Fauna
11-4Hemimetabolous Insects - Collembola and Ephemeroptera
11-5Hemimetabolous Insects - Odonata
11-6Hemimetabolous Insects - Plecoptera
11-7Hemimetabolous Insects - Hemiptera
11-8Holometabolous Insects - Neuroptera, Megaloptera
11-9Holometabolous Insects - Coleoptera Adephaga
11-10Holometabolous Insects - Coleoptera Polyphaga
11-11Holometabolous Insects - Trichoptera, Annulipalpia
11-12Holometabolous Insects - Trichoptera, Integristipula & Spicipalpia
11-13aHolometabola - Diptera, Suborder Nematocera
11-13bHolometabola - Diptera, Suborder Nematocera
11-14Holometabola - Diptera, Suborder Brachycera
Chapter 12 - Terrestrial Insects
12-1Terrestrial Insects - Habitats and Biology
12-2 Terrestrial Insects - Collembola
12-3Terrestrial Insects Hemimetabola - Odonata
12-4Terrestrial Insects Hemimetabola -Orthopteroidea
12-5 Terrestrial Insects Hemimetabola - Notoptera and Psocoptera
12-6 Terrestrial Insects Hemimetabola - Hemiptera
12-7 Terrestrial Insects Hemimetabola - Hemiptera2 & Thysanoptera
12-8 Terrestrial Insects Holometabola - Megaloptera & Neuroptera
12-9a Terrestrial Insects Holometabola - Coleoptera Biology and Ecology
12-9b Terrestrial Insects Holometabola - Coleoptera Families b
12-9c Terrestrial Insects Holometabola - Coleoptera Families c
12-10 Terrestrial Insects Holometabola - Hymenoptera
12-11 Terrestrial Insects Holometabola - Trichoptera
12-12 Terrestrial Insects Holometabola - Lepidoptera Biology
12-13 Terrestrial Insects Holometabola - Lepidoptera 1
12-14 Terrestrial Insects Holometabola - Lepidoptera 2
12-15 Terrestrial Insects Holometabola - Lepidoptera 3
12-16 Terrestrial Insects Holometabola - Mecoptera
12-17 Terrestrial Insects Holometabola - Diptera Overview
12-18 Terrestrial Insects Holometabola - Diptera Nematocera
12-19 Terrestrial Insects Holometabola - Diptera Nematocera 2
12-20 Terrestrial Insects Holometabola - Diptera Brachycera
Chapter 13 - Fish
Chapter 14 - Amphibians
14-1Amphibians Anuran Adaptations
14-2Anuran Conservation Issues
14-3Ground-dwelling Anurans
14-4Anurans Waterfalls, Treefrogs, and Mossy Habitats
14-5Anurans Central and South American Mossy Habitats
14-6Salamanders and Adaptations
14-7Hynobiidae, Ambystomatidae, and Plethodontidae
14-8Salamander Mossy Habitats
14-9Amphibians Bryophyte-dwelling Salamander Checklist
Chapter 15 - Reptiles
Chapter 16 - Birds
16-1 Birds and Bryophyte Intersections
16-2 Birds and Bryophytic Food Sources
16-3Bird Nests
16-4Bird Nests - Non-Passeriformes 1
16-5Bird Nests - Non-Passeriformes 2
16-6Bird Nests - Passeriformes 1
16-7Bird Nests - Passeriformes 2
Chapter 17 - Rodents
17-1Rodents - Muroidea - Muridae
17-2Rodents - Muroidea - non-Muridae
17-3Rodents - non-Muroidea
Chapter 18 - Large Mammals
18-1Large Mammals - Ruminants - Cervidae
18-2Large Mammals - Ruminants - non-Cervidae
18-3Large Mammals - Non-Ruminants

Citation preview

About the Author e Janice Glime is Professor Emerita in the Department of Biological Sciences at Michigan Technological University, Houghton, Michigan, USA. She has a Bachelor of Science degree in elementary education from Frostburg State University, Maryland, USA (1962), a Master of Science in botany from West Virginia University (1964), and a Doctor of Philosophy in botany from Michigan State University (1968). She specialized in teaching freshmen in general biology and botany, and has taught ecology, evolution, systems ecology, and bryology. She is past President of the International Association of Bryologists (IAB) and is the manager of Bryonet-L, the IAB email discussion group on bryophytes. She has published over 100 papers, mostly on b1yophyte ecology, is author of the book The Elfin World of Mosses and Live1worts of Isle Roya{e and the Upper Peninsula of Michigan, co-author with Dinesh Saxena of Uses of Bryophytes, and editor of Methods in Bryofogy. Her primary research interests are on aquatic bryophytes and on the interactions of bryophytes and on the interactions of bryophytes with other organisms.

About the Book For more than ten years I have been working on a book on bryophyte ecology and was joined by Heinjo During, who has been very helpful in critiquing multiple versions of the chapters. But as the book progressed, the field of bryophyte ecology progressed faster. No chapter ever seemed to stay finished, hence the decision to publish online. Furthermore, rather than being a textbook, it is evolving into an encyclopedia that would be at least three volumes. Having reached the age when I could retire whenever I wanted to, I no longer needed be so concerned with the publish or perish paradigm. In keeping with the sharing nature of bryologists, and the need to educate the non-bryologists about the nature and role of bryophytes in the ecosystem, it seemed my personal goals could best be accomplished by publishing online. This has several advantages for me. I can choose the format I want, I can include lots of color images, and I can post chapters or parts of chapters as I complete them and update later if I find it important. Throughout the book I have posed questions. I have even attempt to offer hypotheses for many of these. It is my hope that these questions and hypotheses will inspire students of all ages to attempt to answer these. Some are simple and could even be done by elementary school children. Others are suitable for undergraduate projects. And some will take lifelong work or a large team of researchers around the world. Have fun with them!

The Format The decision to publish Bryophyte Ecology as an ebook occurred after I had a publisher, and I am sure I have not thought of all the complexities of publishing as I complete things, rather than in the order of the planned organization. But I wanted to reach a worldwide audience that included not only professional bryologists, but beginners, non-bryologist ecologists, teachers, naturalists, anyone who wanted to know something about bryophytes. Many of these people would never be willing or able to pay the cost of such a book in print copy. And the cost of the numerous color plates would be prohibitive. Some chapters have been easier for me to do and some will simply need help from others. The "book" will actually be multiple volumes, with the first being physiological ecology, but including an introduction to the broad classification of phyla and classes, morphology, structures, and life cycles. Communities, habitats, roles, interactions, and methods, among others, are in various stages of completion. Large chapters and those with many images difficult to download, so chapters are broken into smaller segments that I shall call subchapters. Sections, chapters, and subchapters will not always be posted in order, so each begins new pagination. Where possible, I will try to number sections of a chapter continuously. New chapters will be added as they are ready but may not cover all planned topics at the onset. Bryologists are encouraged to send me text or images for consideration, or to volunteer to write a chapter. I am considering making this like an online journal with reviewers, but that needs more planning and is likely to make style and nomenclature inconsistent. Your thoughts on the idea would be appreciated.

Acknowledgments The contributors to this book are far too numerous to mention all of them by name. To my graduate students and students of bryology, I owe a debt of gratitude for their enthusiasm for this project and for helping me to write for a somewhat less than professional and experienced audience by critically reviewing early chapters. To the members of Bryonet, I thank you not only for your wonderful contributions through Bryonet, but for the promptness with which I receive help for my many requests for images, information, ideas, and publications, reminding me over and over what a wonderful group of people comprise bryology. From Heinjo During I received numerous helpful suggestions and encouragement to keep going. As my co-author he obtained a contract with Cambridge, which we later abandoned. In the end, he modestly withdrew from authorship, claiming to have made no contribution, but his contributions in reading my chapters have been invaluable. Irene Bisang, as my co-author on the Sexual Strategies chapter update, kept me organized, and I still feel her presence and advice as I work on other chapters. To many persons I owe an immense debt of gratitude for permission to use their images. Without this wide array of choices, the book would have been of incredibly dull appearance on the web, and much less instructive. But most of all, I owe the beauty of the book to Michael Lüth, who gave me blanket permission to use as many of his wonderful images as I wished. They have provided more than half the bryophyte images used and put my own early photographic efforts to shame. Finally, I acknowledge the support of the Botanical Society of America, the International Association of Bryologists, and most of all, the Department of Biological Sciences of Michigan Technological University for sponsorship of the web version of the book. To my department chair, John Adler, I appreciate his cooperation and support in publishing this as an online book instead of a printed one. To Emil (Tiger) Groth, I owe the web layout and all the web activity needed to place the book there in an accessible and searchable form. And to Annelise Doll and the E. R. Lauren Library staff, we all owe gratitude for selecting this book for Digital Commons and for doing her best to meet all my requests, assuring that this book will be preserved for posterity.

Glime, J. M. and Chavoutier, L. 2017. Glossary. In: Glime, J. M. Bryophyte Ecology. Ebook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 16 July 2020 and available at .

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GLOSSARY JANICE GLIME AND LEICA CHAVOUTIER

1n: having only one set of chromosomes 2n: having two sets of chromosomes

2,4-D: 2,4-dichlorophenoxyacetic acid; herbicide that mimics IAA 6-methoxybenzoxazolinone (6-MBOA): glycoside derivative; insect antifeedant; can stimulate reproductive activity in some small mammals that eat them by providing growth substances >>: much greater ♀: sign meaning female, i.e. bearing archegonia ♂: symbol meaning male

A α-amylase: enzyme that hydrolyses alpha bonds of large, alphalinked polysaccharides, such as starch and glycogen, yielding glucose and maltose A horizon: dark-colored soil layer with organic content and minerals intermixed ABA: abscisic acid; plant hormone (growth regulator) associated with water stress, drought hardening, growth inhibition, stomatal closing, and seed dormancy in some plants; known from mosses abandoned land: land having previous human use abaxial: referring to lower surface of leaf; facing away from stem of plant

Abbreviations aff.: related to agg.: aggregate, designating group of species which are difficult to distinguish from one another auct.: Latin abbreviation for auctor, meaning author c.: Latin circa, meaning around, about cf.: Latin confer, compare with cfr. (c. fr.): Latin cum fructibus, meaning with sporophytes cm: centimeter det.: Latin determinavit, meaning determined by e.g.: Latin exempli gratia, meaning for example fo.: Latin forma, meaning form ibid.: Latin ibidem, meaning in the same book i.e.: Latin id est meaning that is IPL: inner peristomial layer leg.: Latin legit, meaning collected by µm: micrometer; micron; length unit = 1/1 000 mm. n: chromosome number (haploid). op. cit.: Latin opus citatum, meaning mentioned, cited above OPL: meaning outer peristomial layer PPL: meaning primary peristomial layer s.d.: Latin sine die, meaning without date sensu: Latin sensu, meaning in the sense (of) s.l.: Latin sensu lato, meaning in broad sense s.n.: Latin sine numero, meaning without number

s.s.: Latin sensu stricto, meaning strict sense sp.: species spp.: more than one species ssp.: subspecies var.: variety

abiosis: absence or lack of life; nonviable state abiotic: referring to non-living and including dust and other particles gained from atmosphere, organic leachates from bryophytes (and host trees for epiphytes), decaying bryophyte parts, and remains of dead inhabitants; usually includes substrate abortive: having development that is incomplete, abnormal, stopped before maturity abscisic acid: ABA; plant hormone (growth regulator) abscission: process where plant organs are shed; e.g. deciduous leaves in autumn absent: missing abundance: numerical representation of species; measure of amount of given species in sample local abundance is relative representation of species in particular ecosystem, usually measured as number of individuals found per sample relative species abundance is calculated by dividing number of species from one group by total number of species from all groups acaulescent: provided with very short stem ACC: acetyl-CoA carboxylase; ethylene precursor; biotindependent enzyme that catalyzes irreversible carboxylation of acetyl-CoA to produce malonyl-CoA through its two catalytic activities, biotin carboxylase (BC) and carboxyltransferase (CT) accession number: number assigned to specimen when it is entered into herbarium record accessory pigment: pigment that captures light energy and passes it to chlorophyll a accidentally foliicolous: accidentally, not normally, growing on leaves acclimation: gradual and reversible adjustment of organism to environmental fluctuations; e.g. adjustment to winter cold or summer heat; compare to adaptation, which is persistent genetic change that provides organism with better ability to survive its environmental conditions accrescent: continuing to grow after reproduction accumulation enrichment factor: amount of metal in plants divided by its stream water concentration -aceae: suffix denoting family in Plant Kingdom acellular: not divided into multiple cells

G-2

Glossary

acetylcholine: chemical formed by choline and acetyl group; neurotransmitter in nervous system used to transmit nerve impulses achlorophyllous: lacking chlorophyll acicole: growing on or among needles of conifers acid: substance with pH less than 7.0 acid flush: concentrated pollutants released rapidly during snow melt acid precipitation: precipitation having pH less than 5.4 acidicline: preferring weakly acidic substratum acidophile: plant growing best on acidic substrate acidophilous: growing on acidic substrates acrocarp: moss species that produces sporophyte at apex of stem or main branch acrocarpous: gametophyte producing sporophyte at apex of stem or main branch; generally upright mosses with terminal sporangia, usually unbranched or sparsely branched acrogynous: in many leafy liverworts, sporophyte growing at top of stem (from apical cell), e.g. Mesoptychia collaris [ant. anacrogynous] acropetal: referring to movement of substance from base to apex; of growth, outward toward shoot (or root) apex [ant. basipetal] acrotelm: living layer of peat actinomorphic: having radial symmetry, like spokes of wheel activation conditions: conditions of sufficient moisture and light for germination acuminate tip: prolonged tip adaptation: genetic change, arrived at through process of natural selection, which enables organism to compete more effectively under given set of conditions (L. adaptare = to fit in); compare to acclimation, gradual and reversible adjustment of organism to environmental fluctuations adaxial: on side toward axis (stem) of plant, such as upper surface of leaf [ant. abaxial] adenine: nitrogenous base; one member of base pair adeninethymine in DNA adherent: strongly attached to substratum e.g. Frullania dilatata adhesion tube: in Collembola, attachment to abdomen that may be used for cleansing body and as means of transferring drops of water from surface of body to mouth where they are then absorbed; previously thought to provide adherence adhesive organ: structure by which some nematodes adhere to bryophytes adhesive peg: structure of fungus that contacts rotifer or other entrapped organism, stimulating fungus to release glue from its trap adnate: said of two fused structures, e.g. peristome and epiphragm of Atrichum undulatum adsorption: fixation of elements on surface adventitious: growing on atypical place, e.g. adventitious rhizoids on costa in Conardia compacta adventitious root: root that arises from stem or other non-root axis point, as seen in corn adventive: introduced aerenchyma: in some thallose liverworts, loose parenchyma, with empty spaces between groups of cells aerobiology: study of airborne microorganisms, pollen, spores, and seeds, especially as agents of infection; form of passive transport aerohaline: subject to influence of salty sea spray

aerohygrophyte: plant growing in habitats having high air humidity aerophyte: plant growing on aerial parts of another aestivation: state of animal dormancy, similar to hibernation, but taking place in summer rather than winter Afro-alpine: high mountains of Ethiopia and tropical East Africa, which represent biological 'sky islands' with high level of endemism Afromontane: subregions of Afrotropical realm, one of Earth's eight biogeographic realms, covering plant and animal species found in mountains of Africa and southern Arabian Peninsula aggregate: clustered together; group of species which are difficult to distinguish from one another aggressive mimicry: form of mimicry in which predators, parasites, or parasitoids share similar signals, using harmless model, allowing them to avoid being correctly identified by their prey or host; e.g. playing dead Agral 600: horticultural wetting agent agroforest: land use management forest in which trees or shrubs are grown around or among crops or pastureland air chamber: in some thallose liverworts, specialized aircontaining cavity air layering: method of propagating plant by girdling or cutting part way into stem or branch and packing area with moist medium, as Sphagnum moss, stimulating root formation so that stem or branch can be removed and grown as independent plant air pore: in some thallose liverworts, opening of air-chamber alanine: non-polar amino acid that is relatively insoluble in water; defense compound that enables plants to withstand various stresses such as hypoxia, waterlogging, and drought alar cell: cell at basal angle of moss leaf, usually different in size and shape from other leaf cells -ales: suffix applied to order of plants or algae (e.g. Dicranales, Orthotrichales) alginate: viscous gum; general term for salts of alginic acid, especially sodium but also calcium or barium ions; composed of guluronic and mannuronic acids alkaline: rich in bases, having pH of more than 7 alkalinity: capacity of water to resist changes in pH that would make water more acidic; equivalent sum of bases that are titratable with strong acid alkaloid: basic organic compound containing nitrogen; toxic allele: particular form of gene allelopathic: having ability to inhibit growth of another organism through secondary metabolite allelopathy: condition in which one organism makes environment chemically unsuitable to another through secondary metabolism; type of chemical warfare in plants allochthonous: originating from elsewhere allopatric: said of two species which have separate (nonoverlapping) areas of distribution allopolyploidy: type of polyploidy (multiple sets of chromosomes) in which chromosome complement consists of more than two copies, with chromosomes derived from different species, producing hybrid species alluvium: deposit of clay, silt, sand, and gravel left by flowing water in river valley or delta, usually as fertile soil alpestrine: subalpine; growing to tree line alpha amylase: enzyme that hydrolyses alpha bonds of large, alpha-linked polysaccharides, such as starch and

Glossary

glycogen, yielding glucose and maltose; substance that helps some rotifers identify plant substrate alpha diversity: mean species diversity in sites or habitats at local scale, i.e. local species diversity alpine: habitat above treeline of mountain alternation of generations: alternating cycle of sporophyte (2n) and gametophyte (1n) generations altimontane: montane grasslands, shrublands, and woodlands alveola (pl. alveolae): more or less polygonal surface depression alveolate: with depressions on surface Amass: leaf mass per area Amax: maximum assimilation ambush predator: sit and wait predator, often having camouflage amensalism: interaction in which one species is harmed by other while other is neither harmed nor benefitted ametabiotic: describes metabolic state of life entered by organism in response to adverse environmental conditions such as desiccation, freezing, or oxygen deficiency; all measurable metabolic processes stop, preventing reproduction, development, and repair; cryptobiotic ametabolic state: state of life entered by organism in response to adverse environmental conditions such as desiccation, freezing, or oxygen deficiency; cryptobiotic state in which all measurable metabolic processes stop, preventing reproduction, development, and repair; including tardigrades, free-living nematodes, and rotifers; having no obvious metamorphosis amictic: non-sexual, as in some rotifers, with asexual reproduction recurring until conditions are favorable amidon: macromolecule composed of glucose constituents; starch; (L. amylum = complex carbohydrate) amoeboflagellate: in some slime molds, diploid cell stage that includes flagellated cells and amoeboid cells that develop directly into plasmodium amorphous: without definite form amphibious: capable of living in or out of water amphigastrium (pl. amphigastria): underleaves of leafy liverworts; few mosses where upper or lower leaves are differentiated from lateral leaves and smaller, as in Racopilum amphispory: spore size frequencies and mean spore size frequencies grouped around 2 mean sizes in varying ratios; small spore fraction is aborted amphithecium (pl. amphithecia): outer layer of embryonic capsule that gives rise to capsule tissues amphitropical: distributed on both sides of tropics amplexus: mating stage in which male amphibian grasps female with his front legs prior to depositing sperm on her eggs amyloid: waxy translucent substance of various complex proteins in combination with polysaccharides and staining blue with iodine (like starch) deposited in tissues in different disease processes and tissue degeneration; builds up inside tissue in amorphous way amyloplast: colorless plastid that forms starch granules in plants; statolith; might play role in gravitropism anabiosis: temporary state of suspended animation or greatly reduced metabolism anacrogynous: designating sporophyte growing in lateral position on stem, branch or thallus (e.g., thallose liverworts like Pellia endiviifolia)

G-3

anadromous: referring to fish living in ocean and migrating up freshwater streams to spawn anaerobic: without oxygen anagenesis: species formation without branching of evolutionary line of descent anagenetic speciation: speciation on islands through gradual change from founder population analogous: said of structures not having common phylogenetic origin but having similar function anastomosis: condition of union of one structure with another, usually crisscrossing; interconnecting; may be applied to irregularly divided peristome teeth (e.g. endothecium of Anthelia juratzkana) or river with islands and meanders anchor ice: submerged ice anchored to bottom of stream or other water body ancophile: plant living in canyon forests ancophilous: living in canyon forests Andreaeobryopsida: class of mosses in Bryophyta Andreaeopsida: class of mosses in Bryophyta androcyte: cell that will give rise to antherozoid androecial branch: specialized branch bearing antheridia and bracts androecium (pl. androecia): male inflorescence; antheridia and surrounding bracts androgametophyte: male gametophyte androgynogametophyte: autoicous or synoicous gametophyte androgynous: male and female organs in same inflorescence, monoicous anemochorous: wind-dispersed anemochory: dispersal by wind, such as spore, gemma, or other propagule angle of incidence: angle formed between direction of light and vertical (difference from straight on), so low sun has higher angle of incidence, thus small leaf angle (approaching vertical) creates effect of large angle of incidence anhydrobiosis: dormant state; strategy of surviving dehydrated state or extreme temperature conditions; reviviscence anion: negatively charged ion anisogamy: size, shape, or behavioral differences in gametes anisophyllous: having two types of leaves on same stem; stem leaves and branch leaves morphologically different, as in Sphagnum [ant. isophyllous] anisosporous: having bimodal distribution of spore sizes with smaller spores generally producing males anisospory: condition having bimodal distribution in spore size; genetically determined condition of two spore sizes anisotropic dispersal: directional dispersal annotinous: with yearly growths annual: plant that germinates, reproduces, and dies all within one year [ant. perennial]; see Mägdefrau life forms annual shuttle: species that requires small disturbances that last 1-2 years; survive severe stress periods annular: ring-shaped annulus: zone of differentiated cells between capsule urn and operculum, facilitating opening of capsule anoxybiosis: biological response triggered by lack of oxygen in which organism takes in water and becomes turgid and immobile, possibly form of cryptobiosis; used by tardigrades to survive unfavorable conditions

G-4

Glossary

antagonistic: interaction in which one species benefits at expense of another anterior: dorsal, abaxial [ant. posterior] anterior whiplash flagellum: thin whiplike structure on front end of cell (L. flagellum = whip) antheraxanthin: bright yellow accessory pigment found in many organisms that perform photosynthesis; xanthophyll cycle pigment, oil-soluble alcohol within xanthophyll subgroup of carotenoids; in pathway to making ABA antheridiophore: specialized antheridium-bearing branch antheridium (pl. antheridia): male gametangium found in all sexual plants except seed plants; sperm container, multicellular globose to broadly cylindric stalked structure producing sperm antherozoid: spermatozoid, male gamete Anthocerotophyta: phylum of hornworts, characterized by thallose gametophyte with hornlike sporophyte having continued growth at its base anthocyanin: water-soluble blue, purple, or red flavonoid pigment found in cell vacuole of plants, especially flowers and autumn leaves; in bryophytes, usually based on 3desoxyanthocyanidins located in cell wall anthracine: coal black anthropochorous: dispersal of propagules associated with human activities anthropogenic: relative to ecosystem, resulting from action of humans antical: relative to surface of thallus, upper side [ant. postical] antifeedant: compound that discourages herbivory antifreeze protein (AFP): protein that prevents freezing antrorse: forward, upward, toward tip, e.g. antrorse teeth in Dichodontium pellucidum [ant. retrorse] aperturate: with opening aperture: opening, hole, orifice apex: tip; end farthest from point of attachment or from base of organ (L. apex = point) aphyllous: without leaves apical: at tip or apex apical cell: single meristematic cell at apex of shoot, thallus, or other organ that divides repeatedly apical dominance: phenomenon whereby main, central stem of plant is dominant over other side branches, typically by supressing their growth apiculate: with short and abrupt point apiculus (pl. apiculi): short point, e.g. leaf tip of Entodon concinnus apogamous: condition of producing sporophyte without union of gametes apogamy: asexual multiplication, without fusion of gametes [syn. apomixis] apomixis: asexual multiplication, without fusion of gametes [syn. apogamy] apophysis: strongly differentiated sterile neck at base of capsule, e.g. Splachnum rubrum [syn. hypophysis] apoplast: capillary spaces in cell wall apoplastic: outside cell membrane, such as cell walls and dead cells; used to describe water transport between cells aposematic mimicry: resemblance to organisms with behavior or morphology serving to warn or repel

aposematism: warning coloration; advertising by animal to potential predators that it is not worth attacking or eating; may indicate poisonous or bad taste or carnivorous attack aposporous: producing gametophyte from sporophyte tissue without meiosis apparency: hypothesis predicts that apparent plants (i.e., most easily found in vegetation) would be most commonly eaten by herbivores, including humans; grouping of plants, including bryophytes, that are most conspicuous photosynthetic food items available apparent plants: conspicuous plants, easily found by herbivores apparent quantum yield: measure of how many molecules of certain substance such as H2O2, dissolved inorganic carbon, etc. can be produced per photon absorbed by, for example, colored dissolved organic matter appressed: referring to leaves lying closely or flat against stem or plant to substrate [Frullania dilatata] aquatic: pertaining to water habitat arabinoglucan: new polysaccharide from mosses, made of glucose and arabinose; has potential medicinal value arabinose: monosaccharide sugar containing five carbon atoms, and including aldehyde (CHO) functional group arable land: land used for or suitable for growing crops arachidonic acid: polyunsaturated, essential fatty acid that makes membranes more pliable in cold arachnoid: covered with fine and tangled hairs, e.g. Marchantia polymorpha ssp. montivagans archegoniophore arboreal: living in trees arbuscular hypha (pl. hyphae): mycorrhizal filament characterized by formation of unique structures, arbuscules, and vesicles by fungi of phylum Glomeromycota arbuscule: finely branched organ produced by endomycorrhizal fungi inside host cells; interface at which fungus and plant exchange phosphorus and photosynthates archegoniophore: specialized archegonia-bearing branch archegonium (pl. archegonia): multicellular egg-containing structure that later houses embryo; female gametangium; flask-shaped structure consisting of stalk, venter, and neck present in Bryophyta and all tracheophytes except flowers archesporium: layer of cells which give rise to spores Arctic: present in areas around North pole arctic-alpine: distribution in arctomontane: distribution in Arctic region and montane areas in lower latitudes; climatic type of Arctic and high elevations area: region of distribution arenicolous: growing on sand areola (pl. areolae): small, angular or polygonal surface area differentiated on thallus and overlying chamber, forming pattern or network, as in Conocephalum areolate: divided into chambers areolation: cellular network of leaf or thallus argillicolous: growing on clay soils arginine: highest nitrogen to carbon ratio among 21 proteinogenic amino acids; amino acid with basic group, alkaline in solution; water soluble; major storage and transport form for organic nitrogen in plants arid: having little or no rain arista: awn; hair point, e.g. leaf tip of Syntrichia caninervis aristate: ending in awn, e.g. Syntrichia ruralis leaves

Glossary

arthrodontous: having lateral walls of peristome teeth eroded with uneven thickenings (arthro = jointed; don = tooth), e.g. peristome of Orthotrichum cupulatum ascending: pointing obliquely upward, away from substrate Ascomycota: phylum of fungi commonly known as sac fungi because spores are produced in sacs called asci aseptic: free of disease-causing microorganisms asexual: referring to reproduction without union of gametes, such as gemmae in Marchantia asl: above sea level aspartate: amino acid with higher molecular weight and protonated -NH+3 aspect: compass direction slope faces astomous: without stomata (capsule); capsule that doesn't open ATP: adenosine triphosphate; energy-storing compound atratous: turning black Aufwuchs: German word for small organisms living firmly attached to substratum, but not penetrating it; see also periphyton auricle: earlike lobe, sometimes at base of moss leaf or liverwort underleaf; in Blasia houses Cyanobacterial partner auroxanthin: diepoxy carotenoid pigment known in Fontinalis austral: of Southern Hemisphere author(s): name(s) of bryologist(s) (sometimes abbreviated) who contributed to taxonomic description and nomenclature of taxon autoclave: oven-like equipment capable of high temperatures for heat sterilization autogamy: within one gametophytic self-fertilization autohydrolysis: hydrolysis (molecule of water ruptures one or more chemical bonds) of peptide or enzyme catalyzed by itself autoicous: having male and female reproductive organs in separate clusters (different branches) on same plant autolysis: release of enzymes when cells die, causing cells to break down quickly; common in many insects autopolyploidy: all chromosomes derived from same species, frequently same individual; in bryophytes, having more than 1 set of homologous chromosomes in gametophyte autotomy: self-amputation; behaviour whereby animal sheds or discards one or more of its own appendages, usually as selfdefense mechanism to elude predator's grasp or to distract predator and thereby allow escape autotropism: tendency of plant organs to grow in straight line when not influenced by external stimuli auxin: plant growth-regulating hormone, usually referring to hormone indoleacetic acid (IAA); influences cellular elongation, among other things avoidance strategy: adaptations that permit organism to alter factor so that it is no longer significantly damaging, such as minimizing hydrodynamic forces by adaptive life form awn: hair-point, e.g. leaf tip of Cirriphyllum piliferum axenic: pure (sterile) culture, without other organisms axial strand: column formed of elongated cells and located in center of some stems or thalli; central strand in mosses axil: angle formed where leaf joins stem axillary: forming in axis between stem and leaf axis: main stem axopod: sticky pseudopod on some Protozoa

G-5

B B horizon: dark soil layer of accumulated transported silicate, clay, minerals, iron, and organic matter, having blocky structure Baas-Becking hypothesis: everything is everywhere, but, the environment selects; applied to small organisms and propagules such as spores bacterivore: consumes primarily bacteria Baermann funnel: apparatus for extracting turbellarians (as well as nematodes, copepods, and tardigrades) from bryophytes; cheese cloth, muslin, or tissue paper is placed in funnel to hold sample, usually supported by piece of screening; water is run through sample with rubber tubing clamped at end of funnel; sample sits overnight or longer, then water is released from funnel and collected; first few drops will have concentration of nematodes, which are heavier than water Baker's law: loss of dispersal power and bias toward selfcompatibility after immigration to islands ballooning: phenomenon in which spider ascends to something taller, like fence, points its spinnerets upward, then secretes thread, then jumps or is blown with thread serving as anchor bana: low Amazon caatinga tall bana: type of low caatinga with trees over 10 m tall low bana: type of low caatinga with maximum tree height typically less than 5 m open bana: in central low caatinga where trees are even shorter and very widely spaced bank: land along side body of water scientific unit of measurement of pressure; 1 bar  1 atmosphere of pressure (0.986923 tam)  14.503 psi = 750 mm Hg = 99.992 kPa barbate: with tufts of long hairs, beard-like bark: outermost layer of stems and roots of woody plants; surrounding wood of tree or shrub basal cells: group of cells located at base, in proximal part of leaf basal membrane: short cylinder at base of peristome (single peristome) or at base of endostome (double peristome) supporting segments and cilia basic: alkaline, containing base, having pH higher than 7 Basidiomycota: phylum of fungi; fungi composed of hyphae and reproducing sexually by formation of specialized clubshaped end cells called basidia that normally bear external meiospores (usually four) basionym: original name on which current taxon name is based basipetal: referring to movement of substance from apex to base; tissue or organ developing or maturing from apex toward base [ant. acropetal] basiphile: preferring basic habitats (limestone, sandstone, chalk, dolomite, etc.) [ant. acidophile] Batesian mimicry: mimicry in which one organism resembles toxic or otherwise dangerous organism, but is not dangerous itself beaded stream: pools connected by narrow channels behavioral drift: occurring at particular time of day or night; may result from crowding, competition, need for food, predation, making new case, or attempting to reach land at emergence time beneficial acclimation hypothesis (BAH): hypothesis that predicts animals will have their best performance at temperature to which they are acclimated benthic: living on bottom of body of water bar:

G-6

Glossary

Bergmann's rule: within broadly distributed taxonomic clade, populations and species of larger size are found in colder environments, while populations and species of smaller size are found in warmer regions; usually applied to endotherms Berlese funnel: apparatus using light and/or temperature gradient that separates mobile organisms such as arthropods and annelids from litter or bryophytes in funnel; organisms collected in preservative (usually alcohol) below funnel beta diversity: ratio between regional and local species diversity bet hedger: organism that uses combination of two or more strategies, thus never having optimal adaptations to extremes but being prepared to lesser degree for most circumstances; plant that seems to have both good sexual reproduction and means of vegetative reproduction, e.g. bryophyte that produces frequent capsules but also produces gemmae, as in Tetraphis pellucida and Marchantia polymorpha bicostate: with two nerves bicuspidate: with two points, e.g. leaves of Cephalozia lunulifolia Bidder's organ: structure on male toads that can become ovary under right conditions bidentate: with two teeth (different from double teeth) biennial: cycle of two season’s duration (generally less than two years) bifarious: on two opposite rows, distichous biflagellate: having two flagella; functions in cell motility bilobate: divided into two lobes or segments, e.g. Lophocolea bidentata binding site: site for attachment, usually referring to ions; can occur on cell walls, soil particles, glass containers, etc. binocular: having two eyepieces binomial: expression used to designate species; formed of two Latin terms: generic and specific term; by convention this binomial is written in italics because it is foreign word bioassay: use of living organism for assessing effects of biologically active substances biocoenosis: association of different organisms living together in habitat; biotic community (or biocenosis) along with its physical environment (or biotope) biomass: quantitative estimate of total mass of organisms or parts being considered biotope: ensemble of physical, chemical and climatic conditions of habitat; biotope plus biocenosis form ecosystem twice pinnately branched, e.g. Thuidium bipinnate: tamariscinum bipolar: said of species found in both polar regions biramous: divided into two branches, e.g. pincers on end of crab claw or divided antenna bird cliffs: steep cliffs with numerous small shelves that serve as nesting locations for bird colonies bisexual: having both sexes on same individual; monoicous (gametophyte) or monoecious (sporophyte of tracheophytes) bistratose: having two layers of overlapping cells, as in some moss leaves bivoltine: producing two broods per season blade: portion of leaf excluding stalk (Plagiomnium) bloom: powder covering some capsules or leaves, e.g. leaves of Saelania glaucescens bog: acidic, wet area in which nutrients are received by rainfall and groundwater flow is negligible; consists mostly of decaying moss and other plant material; characterized by low nutrients

bog moss: usually meaning Sphagnum bole: main trunk of tree bonkei: tray landscape, typically made with bryophytes bonsai: dwarfed ornamental tree, often with mosses at base border: land at edge of habitat; in bryophytes, edge; margin (cells of different shape, size, or color than other cells of structure), e.g. leaf of Mnium thomsonii boreal: pertaining to north; life zone bounded on south by growth-season accumulated temperature above 6.1ºC of 5538ºC and mean daily temperature of 18ºC for six hottest weeks (L. boreas = north) boreal forest: predominantly conifer forest extending across northern North America and parts of Europe and Asia BOREAS: climate model for boreal region botryoid: like bunch of grapes, e.g. oil bodies of Calypogeia suecica boundary layer resistance: boundary layer is that layer of fluid in immediate vicinity of bounding surface; boundary layer resistance is resistance to movement of CO2, heat, and other substances through that thin layer brachycyte: short cell; seen on protonemata treated with ABA brachypterous: short-winged bract: modified leaf associated with gametangium or gemmaecup bracteole: modified underleaf associated with gametangium in liverworts branch: lateral subdivision of stem or axis Braun-Blanquet method: method uses cover-abundance scale to describe vegetation; these levels are divided into cover classes, typically using 5-7 categories:

1 2 3 4 5 6 7

10% eufoliicolous: true leaf-dwelling euhydrobiont: living in water eukaryotic: having nucleus euryoecious: able to live in variety of conditions eutrophic: relative to habitat rich with mineral nutrients and so supporting dense population [ant. oligotrophic] eutrophication: process characterized by excessive plant and algal growth due to increased availability of one or more

G-15

limiting growth factors needed for photosynthesis, such as sunlight, carbon dioxide, and nutrient fertilizers evacuolate: lacking vacuoles evanescent: relative to rib which ends just before apex of leaf, fading, disappearing evaporative cooling: process in which evaporation of water removes heat from system; can occur at plant, animal, or ecosystem level evapotranspiration: loss of water through evaporation from among plants and from plants themselves (transpiration) evenness: similarity of frequencies of different units (species) making up population or sample evergreen: condition where plant remains green and retains its leaves for full year or longer; persistent; green year-round everything is everywhere: Baas-Becking hypothesis that everything is everywhere, but, environment selects; applied to small organisms and propagules such as spores evolution: series of genetic changes (changes that are heritable) that causes organisms to change through time (L. evolutio = unrolling) evolutionary drivers: selection pressures EX: extinct (IUCN) ex: in case of validation after formation of name, e.g. Straminergon stramineum (Dicks. ex Brid.) Hedenäs ex-: prefix meaning "sans," "non" excavate: hollowed, concave exchange site: location on plant cell wall or soil particle where ions are traded, such as replacement of hydrogen from COOH by Ca+2; when charge of new ion is greater than that of one it replaces, it is shared by more than one exchange site exchanger: organism capable of replacing one ion for another, usually replacing hydrogen with cation such as Ca+2 excurrent: relative to rib, beyond apex of leaf, e.g. leaf costa of Fissidens taxifolius exine: outer layer of spore exogenous: growing or originating from outside organism, e.g. fungus can be source of IAA for protonema exogenous: generated by outside source; external origin exohydric: having water transport essentially external by surface flow; including capillary flow between leaves or though surface papillae exoskeleton: rigid external covering for body in some invertebrate animals, especially arthropods, providing both support and protection; e.g. in crayfish exosporic: condition in which first mitotic division occurs outside spore after rupture of spore wall, typical of most bryophytes exostome: outer peristome of arthrodontous capsule, e.g. outer peristome of Orthotrichum striatum exothecial: relative to exothecium, outer capsule wall exothecium: relative to capsule, outermost layer exotic: foreign; introduced from foreign country (L. exoticus = foreign) explant: portion of plant transplanted to artificial medium explerent: life strategy for non-competitive species that fills spaces between others exposed feeder: organism that feeds at exposed surface exserted: relative to capsule that far exceeds perichaetial leaves, e.g. capsules of Orthotrichum anomalum exsiccatum (pl. exsiccata): distributed and labelled reference specimen

G-16

Glossary

extant: existing today [ant. extinct] extensin: glycoprotein thought be involved in cell wall extension extern: relative to surface of leaf, dorsal face, abaxial face extirpation: local extinction extinct: no longer present on Earth [ant. extant] extinction rate: rate of disappearance of species extracellular: on outside of cell extremophile: organism with optimal growth in environmental conditions considered extreme and challenging for carbonbased life form with water as solvent to survive extrorse: turned outwards exuvia (pl. exuviae): cast-off outer skin of tardigrade or arthropod after molt

F ♀: sign meaning female, in bryophytes bearing archegonia face: side facies: general appearance (habit of species), or appearance of plant community dominated by taxon or small number of taxa Factor H: adenine derivative hormone stimulant for inhibiting caulonema growth and promoting formation of gametophore buds in bryophytes facultative: not occurring regularly; occurring optionally in response to circumstances rather than by nature; for example, terrestrial but occasionally surviving in water facultative aquatic: having some degree of tolerance to desiccation and xerophytic conditions facultative diapause: resting period that can change based on conditions facultative epiphyte: organism that lives on trees, but lives on other substrates as well falcate: sickle-shaped falcate-secund: sickle-shaped and turned towards only one side of stem falcation: condition of being curved like sickle, e.g. leaves of many Dicranum species fallow land: plowed and harrowed but left unsown for period false anisospory: condition of having small, non-viable spores found among dimorphic spores in certain species of bryophytes due to factors such as spore abortion; non-genetic condition of more than one spore size false leaf trace: in bryophytes, extension into cortex from leaf but not connected with central strand of stem; found in Mniaceae and Splachnaceae family: subdivision of order – next major classification level; ending in "aceae" fan: life form found on vertical substrate, usually where there is lots of rain; creeping, with branches in one plane and leaves usually flat; e.g. Neckeraceae, Pterobryaceae, Thamnobryum, some Plagiochila; see Mägdefrau life forms farinaceous: farinose, covered with white bloom fascicle: small tuft or cluster of fibers, leaves, branches, or flowers; in Sphagnum, clump of branches on stem fasciculate: arranged in fascicles fastigiate: with branches erect, nearly parallel and nearly same length fault: break in rocks that make up Earth's crust, rocks on each side have moved past each other feces: excrement; waste material discharged from gut

fecundity: number of offspring produced by organism during its lifetime fecundity-advantage model: need of species needs to produce large number of eggs feldmark: plant community characteristic of sites where plant growth is severely restricted by extremes of cold and exposure to wind, typical of alpine tundra and sub-Antarctic environments female: organism that produces egg femur (pl. femora): third segment of leg fen: minerotrophic peatland or moss-dominated ecosystem that gets its nutrients primarily from ground water or surface water; poor fens have low nutrient content, intermediate fens are characterized by intermediate nutrient levels, and rich fens have highest nutrient levels among these habitats; this term has been variously defined in different countries with older North American literature including poor fens as bogs fenestrate: pierced, perforated with openings like windows, e.g. peristome of Grimmia crinitoleucophaea ferredoxin: iron-sulfur protein needed for conversion of nitrogen oxides to NH4+ ferricrete: hard, erosion-resistant layer of sedimentary rock, usually conglomerate or breccia, cemented together by iron oxides ferrugineous, ferruginous: rust colored fertile: producing sex organs (antheridia, archegonia), bearing sporophytes [ant. sterile] fertilization: fusion of gametes resulting in formation of zygote; act of adding nutrients by applying fertilizer to improve plant growth ferulic acid: phenolic compound and major constituent of fruits and vegetables with strong antioxidant and antiinflammatory properties; only released after severe hydrolysis; present in shoots but absent in young capsules of Mnium hornum fibrilla (pl. fibrillae): thickened bands across hyaline cells of Sphagnum, strengthen cell walls; fibril fibrillose: with fibrils, e.g. leaf hyaline cells of Sphagnum field: area of open land, especially one planted with crops or pasture fine adjustment: knob on microscope used for fine-tuning focus; used with high magnifications; see coarse adjustment fire place: construction in which to build fire fistulated: having passageway cut from rumen to outside flank: in some thallose liverworts, zone between median groove and margin of thallus, e.g. thallus of Riccia flavonoids: group of plant pigments that absorb UV light fleshy: soft and thick floristic list: list of species present on site flagellate: possessing flagellum flagelliform: whiplike, gradually tapering from base to tip of branch flagellum (pl. flagella): slender, whip-like appendage that enables cells to move through liquids; differs from cilia in having only one or two per cell; found on most sperm; as propagule, slender branches with reduced leaves that occur in axils of upper leaves – basal portion multicellular, separating them from caducous branchlets flavonoid: group of plant pigments that absorb UV light and include anthocyanins

Glossary

flotation: separation technique requires that density of flotation liquid be greater than that of arthropods but less than that of debris or bryophytes fluorescence: emission of light by substance that has absorbed light or other electromagnetic radiation of different wavelength; due to excited electrons returning to ground state; visible or invisible radiation emitted by certain substances as result of incident radiation of shorter wavelength such as X-rays or ultraviolet light flush: area where water from underground flows out onto surface to create area of saturated ground, rather than welldefined channel; piece of boggy ground, especially where water frequently lies on surface; swampy place; pool of water in field maximum fluorescence of dark adapted material; Fm: fluorescence resulting from flashing bright light on leaf in dark fo.: abbreviation meaning "forma" fogging: technique used for killing insects that involves using fine pesticide spray which is directed by blower fog-stripping: condensing water vapor from frequent fog and mist; often primary means for bryophytes to obtain water in cloud forest foliicolous: growing on leaves [syn. epiphyllous] foliose: leaf-like, leafy foot: basal portion of most bryophyte sporophytes, embedded in gametophyte foot candle: intensity of light from one candle on square foot of surface one foot from candle foot gland: in some rotifers, gland on foot to secrete glue footpath: narrow path suitable for walking foraging: in bryophytes, use of horizontal growth that permits mosses or liverworts to take wider advantage of nutrients and light forb: non-grass herbaceous flowering plant forest: wooded habitat forest gap: opening in forest canopy, often due to fallen tree forest track: something resembling large wooded area, especially in density form: lowest level of classification (below variety), often determined by environment founder principle: small population becomes separated to new location, representing only small portion of variability of species; loss of genetic variation in new population established elsewhere by very small number of individuals from larger population; in bryophytes, includes arrival of only one sex to colonize particular location fount: spring or fountain fountain: natural spring of water fovea: spore ornamentation, depression like golf-ball foveolate: pitted FPOM: fine particulate organic matter fragmentation: breaking into fragments (pieces) frank water: obvious pools of water, as opposed to water adhering to moss frass: excrement of insect larvae; insect feces; fine powdery refuse or fragile perforated wood produced by activity of boring insects freeze avoidance: survival strategy that prevents body fluids (especially arthropods) from freezing at temperatures well below 0°C

G-17

freeze tolerance: ability of plants to withstand subzero temperatures through formation of ice crystals in xylem and intercellular space, or apoplast, of their cells freezing longevity: length of time bryophyte can remain frozen and survive fresh: fresh state; in presence of sufficient moisture freshet: flood of river from heavy rain or melted snow; rush of fresh water flowing into sea freshwater: not salt water frieze: as endive salad, e.g. thallus of Anthoceros agrestis fringe: margin lined with cilia frondose: habit that is densely branched, fern-like frost tolerance: lowest temperature at which no more than defined percent (typically 50%) suffer irreversible damage in net photosynthetic activity relative to unfrozen plants fructification: in slime molds, process of forming sporangia; analogy to vascular plants, synonymous term with sporophyte; used for bryophytes, but considered by some authors as unsuitable for bryophytes fruit inappropriate term by some authors, meaning sporophyte fugacious: fleeting fugitive: life strategy of species that lives in unpredictable environment; generally stays only 1-2 years while habitat remains suitable at site and produce small spores that permit them to be dispersed easily fulvous: reddish yellow functional grouping: species having similar roles in ecosystem fungus (pl. fungi): kingdom and common name for group of non-photosynthetic organisms; sometimes placed in kingdom Mycota; formerly classified as plants, but food reserves, cell wall components, and other biochemical differences have caused biologists to re-classify them into their own kingdom funiform: like rope furcula: forked appendage at end of abdomen in springtail, by which insect jumps furfuraceous: covered with scales furrow: groove, e.g. in thallus of Riccia sorocarpa furrowed: sulcate, grooved fuscous: dark brown and somber color fusiform: elongated, spindle-shaped; tapering at both ends Fv: variable fluorescence of dark-adapted material; difference between maximum and minimum fluorescence Fv/Fm: in photosystem II, variable vs maximum fluorescence; measure of chlorophyll fluorescence; measurement ratio that represents maximum potential quantum efficiency of Photosystem II if all capable reaction centers are open; T. arcula > T. minuta. Likewise, the Assulina-alkanovia group exhibits wet to dry as A. seminulum > A. muscorum > Hyalosphenia elegans and the Trinema lineare group appears as T. lineare var. truncatum/T. lineare > T.

Chapter 2-5: Protozoa: Peatland Rhizopods

lineare var. terricola. Interestingly, these species gradients also follow a large to small size gradient, indicating that small taxa survive better than large ones under dry conditions. It appears that having spines is a disadvantage in dry habitats. Within the genera Euglypha and Placocista, the spined forms (Figure 49) are typical of wetter habitats than are those with shorter spines or no spines. These relationships suggest that the most effective use of these rhizopods for reconstruction of the past water regime is to use the lowest possible level of identification, i.e. species and varieties. One interesting question that arises is whether these spined taxa are really different species and varieties, i.e., genetically different, or if they represent ecotypes – morphological representations of the microenvironment where they occur. For example, Laminger (1975) found that Centropyxis aculeata from greater depths lacked spines and their tests were covered with mineral particles. To test the possibility of ecological morphs, Booth (2001) examined four of the most common taxa in two Lake Superior coastal wetlands: Arcella spp., Assulina spp., Centropyxis cassis type, and the Nebela tincta-parvulacollaris group. Using 74 microsites, Booth compared testate amoeba assemblages based on percent moisture, depth to water table, pH, porosity, depth of living moss, and associated bryophyte and tracheophyte species. He used such parameters as test length and aperture diameter for amoebae from at least ten microsites. In general, there was little correlation between morphological variation and microenvironmental parameters. However, in the Nebela tincta-parvula-collaris group, the test size correlated significantly with pH (r2 = 0.68). Booth concluded that these testate rhizopods are sensitive indicators of waterlevel and pH changes.

Figure 49. Placocista spinosa, a rhizopod typical of wet habitats. Photo by Yuuji Tsukii, with permission.

Many more studies on testate amoeba ecology have been conducted in the Northern Hemisphere than elsewhere (Mitchell & Meisterfeld 2005), making their comparisons somewhat easier. In the East Carpathian peatlands of eastern Europe, species such as Amphitrema flavum (Figure 17) and Hyalosphenia papilio (Figure 12) indicate wet conditions were present (Schnitchen et al. 2003). Assulina muscorum (Figure 50), Difflugia pulex, and Nebela militaris (Figure 23) indicate that conditions were dry.

2-5-15

Figure 50. Test of Assulina muscorum. Photo by Edward Mitchell, with permission.

In Sphagnum peatlands of the Rocky Mountains, USA, surface moisture determines the distribution of fossil rhizopods (Zygmunt et al. 2003). As suggested by the ecological studies of Lamentowicz and Mitchell (2005) and others (Booth & Zygmunt 2005), Booth and Jackson (2001) could track the history of an ombrotrophic peatland in northeastern Lower Michigan, USA, through 2800 years of changes using the moisture preferences of these organisms. Such fossils as these testae of rhizopods permit us to determine past changes in water table depth (Warner 1991; Woodland 1998; Woodland et al. 1998). Booth and Zygmunt (2005) further argued that the widespread geographic nature of the rhizopod relationships makes interpretation of their community structure widely applicable. Charman and Warner (1997) used 60 samples from 14 peatlands in Newfoundland, Canada, and found 40 species that occurred in more than six samples. They used these to model the relationships between the species and the water table depth. Species with narrow tolerances provided the best indicators. These include Amphitrema stenostoma, Arcella discoides, Cryptodifflugia sacculus, Difflugia bacillifera, Nebela carinata, Nebela griseola, Nebela marginata, Quadrulella symmetrica, and Sphenoderia lenta. Charman and Warner recommend that for most accurate results modern constructs from wide regions should be used to interpret the data from peatland cores that represent palaeoecological time series. Fortunately, most of the testate amoeba taxa are cosmopolitan, permitting the studies from the Northern Hemisphere to be used in less-studied areas such as New Zealand (Charman 1997; Wilmshurst 1998). In fact, Charman (1997) modelled the hydrologic relationships of protozoa and Sphagnum in peatlands of New Zealand and suggested that "palaeohydrology could be accurately inferred from fossil faunas." Schoning et al. (2005) used peatland amoebae to reconstruct 125 years of peatland amoebae in Sweden. Unlike the cases in other areas in Europe, the changes in water table correlated primarily with changes in mean annual temperature, whereas in most other studies, precipitation was also an important factor. They caution that spatial differences must be considered in these historic interpretations and thus more study is needed on these influences.

2-5-16

Chapter 2-5: Protozoa: Peatland Rhizopods

In a Michigan, USA, study, Booth (2002) found that most of the eleven peatlands he studied had similar testate assemblages. As in most other studies, depth to water table was the best predictor of the protozoan assemblages. Nevertheless, within a given peatland, community variability was correlated with environmental heterogeneity, adding support to the suggestion of Schoning et al. (2005) regarding spatial considerations. But the testate amoebae in bog/fen habitats also had distinct differences in species between May and late summer-early autumn. Testate amoebae in the swamp community, on the other hand, had no clear difference in community structure between dates. They attributed these differences to the more constant water table and moisture conditions in the swamp. Warner et al. (2007) add further support to the importance of considering seasons, particularly for living rhizopods. In southern Ontario, Canada, the usual factors of soil water content and water table influenced the distribution of amoeboid species and these differ with seasons. But the big differences were in the open bog/fen community, whereas in the swamp community there was no clear seasonal difference between May and August or October. The historical record will not take us back forever. In their study on bogs in Ontario and Minnesota, Warner and Charman (1994) found that cores spanning the entire Holocene era only exhibited rhizopods present in the last 6500 years. They indicated that the fauna changed from the early rich fens with sedges and brown mosses. At those early stages, the protozoan communities were dominated by Cyclopyxis and Centropyxis. By 5000 BP, the habitat had become Sphagnum-dominated and the predominant protozoan taxa had shifted to Amphitrema flavum, Assulina muscorum, Heleopera sphagni, and Hyalosphenia subflava. As the habitat became drier, taxa again shifted to Nebela griseola, N. militaris, and Trigonopyxis arcula.

difference in the estimations of water table depth. However, in minerotrophic peatlands, with large numbers of this Euglyphida group, the loss of these tests leads to an underestimation of the water table depth. Data on more alkaline fens are lacking, and the community structure there is not well known. If this idiosome group is not dominant there, reconstruction may be more accurate. Swindles and Roe (2007) likewise found that under conditions of low pH, such as found in peatlands, the degree of dissolution was highly variable, but it did not seem to relate to xenosomic (using "foreign" materials) vs. idiosomic tests. Euglypha (Figure 51) is particularly susceptible, whereas Assulina muscorum (Figure 50), Amphitrema flavum (Figure 34), and Trigonopyxis arcula (Figure 52) are affected little by acidity. Payne (2007) found similar results by subjecting rhizopod tests to weak acid, nutrient enrichment, and desiccation over 28-months, and used shorter-term experiments with stronger acids in peatlands. He determined that during dry periods the record may be altered by differential preservations of the tests, as demonstrated by significant effects of long-term desiccation and short-term acid treatment at two different concentrations. This consequence could lead to overestimating water table depths.

Geographic Differences Despite a considerable number of studies indicating usefulness of these organisms, use of testate amoebae to determine past habitats can at times be misleading. Harnish examined mires in Central Europe (1927 in Paulson 1952-53) and in Lapland, North Sweden (1938 in Paulson 1952-53), and found that the communities were not similar. Rather, associations from Central Europe did not exist in raised bogs in Lapland. In fact, the Amphitrema association existed in Lapland, but in different habitats, not raised bogs, whereas in Central Europe it was confined to raised bogs. The Hyalosphenia type was also absent in the Lapland raised bogs. Problems in Using Rhizopods There are caveats in using fossilized amoeba tests to assess past communities of testate rhizopods. Not all tests are equally preserved (Mitchell et al. 2007). The Euglyphida, which includes the common Euglypha species (Figure 51), are an idiosome group that secretes its own test and its biosilica plates (Beyens & Meisterfeld 2001). This biological test decays more readily than the testae of the other groups (Mitchell et al. 2007). In Sphagnum peatlands, this differential decay seems to make little

Figure 51. SEM detail of biosilica plates of Euglypha penardi, a protozoan for which the test is especially susceptible to dissolution. Photo by Edward Mitchell, with permission.

Human Influence on Development In New Zealand, it appears development of Sphagnum bogs has been dependent on human activity such as clearing or modifying the vegetation, resulting in Sphagnum dominance (Wilmshurst 1998). In other places, clearing of a peatland means that without human intervention it is gone forever. After such loss, it is often desirable to reconstruct the peatland. Testate amoebae have been used to define the past nature of the peatland for reconstruction purposes (Charman 1997; Charman & Gilbert 1997). In a Polish peatland, a rapid shift in peat accumulation and lower pH occurred ~110-150 years ago, with a shift to

Chapter 2-5: Protozoa: Peatland Rhizopods

a Sphagnum-dominated poor fen (Lamentowicz et al. 2007). The protozoa supported this history. Researchers interpreted this to be a result of forest clearance in surrounding areas. Whereas peatlands are often destroyed by human activity, in some cases those activities make conditions more favorable to peatland development. In this case, Sphagnum peatland replaced a species-rich poor fen.

Figure 52. Trigonopyxis arcula test showing opening for pseudopod. This test is more stable than that of Euglypha. Photo by Yuuji Tsukii, with permission.

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vertical profile, whereas in the minerotrophic fen they were numerous only at the surface. As in other studies, moisture conditions were important, but peat composition and minerals also played important roles. Following restoration, species that indicated dry conditions disappeared, whereas the moisture gradient seemed to result in less defined community differences. In fact, the minerals seemed to have a greater effect.

Figure 53. Bullinularia indica. Photo by Edward Mitchell, with permission.

Laggoun-Défarge et al. (2008) found testate amoebae can be used to reflect disturbances that result from peat harvesting. Where better carbohydrate preservation was present, along with more heterogeneous peat composition, the testate amoebae exhibited a higher diversity, thus serving as a biological indicator of conditions. Use in Peatland Regeneration Regeneration of peatlands can use remains of testate amoebae to determine the species to re-introduce or to follow the progress in a less labor-intensive fashion by monitoring the amoebae. In the Jura Mountains, Switzerland, Laggoun-Défarge et al. (2008) examined a peatland that had been mined for heating fuel until World War II and found that amoeba communities changed as peatlands changed during regeneration. The Sphagnum habitat shifted from moderately acidic, wet conditions to more acidic, drier conditions. During these changes, biomass and mean size of amoebae declined while remaining higher at the undamaged site. At the same time, species richness and diversity increased while density declined. As reported by Mitchell et al. (2004), changes in the amoeba community lagged behind that of the returning Sphagnum community. Moreover, during the forty years of 1961-2001, overall amoeba richness (33) remained unchanged, but richness per sample decreased from 11.9 to 9.6 (Kishaba & Mitchell 2005). Relative abundance changed, with three species increasing significantly [Bullinularia indica (Figure 53) (+810%), Cyclopyxis eurystoma (+100%, 0 in 1961), Nebela tincta (Figure 54) (+97%)] and two species declining [Assulina muscorum (Figure 50) (-63%), Euglypha compressa (Figure 55) (93%)]. The researchers concluded the expected changes in richness were complete before the 1961-2001 study began. Jauhianinen (2002) demonstrated in an ombrotrophic bog that the testacean shells were present throughout the

Figure 54. Nebela tincta test with living amoeba. Photo by Edward Mitchell, with permission.

Figure 55. Opening of test of Euglypha compressa. Photo by Edward Mitchell, with permission.

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Chapter 2-5: Protozoa: Peatland Rhizopods

Lamentowicz et al. (2008) demonstrated that the testate amoebae record in a Baltic coast peatland in Northern Poland correlated well with the stable isotope data in the same core. The large number of testate protozoans known from peatlands, their relatively cosmopolitan distribution, and the understanding we have of the water table requirements for many of these species provide us with a useful tool for understanding the past history of many peatlands.

Summary Peatlands support an abundant bryophyte fauna, with Amphitrema, Assulina, Corythion, Difflugia, Euglypha, Heleopera, Hyalosphenia, and Nebela typically being the most common genera. Sphagnum sports more species than those found among other mosses or tracheophytes. These taxa are widespread and thus are very reliable indicators of moisture conditions in the peatlands and are less affected by water chemistry than are the tracheophytes. Diversity is lowest in the driest peatland habitats, but the number of individuals is highest. Abundance increases with depth if oxygen is not limiting. Dry habitat species are more tolerant of changes in water depth than are wet habitat species. Rich fen amoeba species differ from those of acid bogs, but Euglyphidae are prominent in all these habitats. Paraquadrula irregularis and Centropyxis discoides are restricted to fens, with Arcella discoides indicative of rich fens. Detritus forms a major portion of the protozoan diet in the peatlands. Vertical zonation presents the symbiotic taxa in the light zone at the top of the moss, with those requiring more moisture occurring at the greatest depths. Shell size, pH, moisture, light, nutrients, and available food all contribute to the distribution. Horizontal variation results from differences in bryophyte species and microtopography, resulting in differences in distance from water table and in pH. Seasonal differences reflect some of these same changes in moisture and food availability and are effective in separating niches of closely related species. CO2 enrichment may cause a reduction in testate amoebae while at the same time increasing bacterial biomass. Loss of the ozone filter and consequent increase in UV-B radiation may actually favor some testate amoebae in Sphagnum peatlands. Amoebae form more constant associations in peatlands than do the plants. And testate species, with few exceptions, are well preserved even after death. Therefore, they can serve as appropriate markers of past climates as well as indicators of predisturbance conditions, although tests of some species, especially Euglyphidae, decompose more easily than others and can skew the results. The best indicators are those with narrow tolerance ranges, especially for moisture.

Acknowledgments Edward Mitchell was particularly helpful in providing me with needed pictures. Most of the others came from

photos by Yuuji Tsukii who gave me permission to use anything of his on the Protist Information Server website. Thank you to Matthieu Mulot for suggesting a correction to one of the Protozoa names.

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testate amoebae communities on Qeqertarsuaq (Disco Island), West Greenland. Acta Protozool. 44: 253-263. Mauquoy, D. and Barber, K. 2002. Testing the sensitivity of the palaeoclimatic signal from ombrotrophic peat bogs in northern England and the Scottish Borders. Rev. Palaeobot. Palynol. 119: 219-240. Mazei, Y. A. and Bubnova, O. A. 2007. Species composition and structure of testate amoebae community in a Sphagnum bog at the initial stage of its formation. Biol. Bull. 37: 619-628. [Orig.: Izvestiya Akademii Nauk, Seriya Biologicheskaya, 2007, No. 6, pp. 738-747.]. Mazei, Y. A. and Tsyganov, A. N. 2007/2008. Species composition, spatial distribution and seasonal dynamics of testate amoebae community in a Sphagnum bog (Middle Volga region, Russia). Protistology 5: 156-206. Mazei, Y. A., Tsyganov, A. N., and and Bubnova, O. A. 2007. Structure of a community of testate amoebae in a Sphagnum dominated bog in upper Sura Flow (Middle Volga Territory). Biol. Bull. 34: 382-394. McGlone, M. S. and Wilmshurst, J. M. 1999a. A Holocene record of climate, vegetation change and peat bog development, east Otago, South Island, New Zealand. J. Quatern. Sci. 14: 39-254. McGlone, M. S. and Wilmshurst, J. M. 1999b. Multiproxy surface wetness records from replicate cores on an ombrotrophic mire: Implications for Holocene palaeoclimate records. J. Quatern. Sci. 14: 451-463. Meisterfeld, R. 1977. Die horizontale und vertikale Verteilung der Testaceen (Rhizopoda, Testacea) in Sphagnum. Arch. Hydrobiol. 79: 319-356. Meisterfeld, R. 1978. Die Struktur von Testaceenzönosen (Rhizopoda, Testacea) in Sphagnum unter besonderer Berücksichtigung ihrer Diversität. Verh. Gesell. Ökol. 7: 441-450. Meisterfeld, R. 1979a. Zur Systematik der Testaceen (Rhizopoda, Testacea) in Sphagnum. Eine REMUntersuchung. Arch. Protistenk. 121: 246-269. Meisterfeld, R. 1979b. Clusteranalytische Differenzierung der Testaceenzönosen (Rhizopoda, Testacea) in Sphagnum. Arch. Protistenk. 121: 270-307. Meisterfeld, R. and Heisterbaum, M. 1986. The decay of empty tests of testate amoebae (Rhizopoda, Protozoa). Symp. Biol. Hung. 33: 285-390. Mieczan, T. 2006. Species diversity of protozoa (Rhizopoda, Ciliata) on mosses of Sphagnum genus in restoration areas of the Poleski National Park. Acta Agrophys. 7: 453-459. Mieczan, T. 2007. Epiphytic protozoa (testate amoebae and ciliates) associated with Sphagnum in peatbogs: Relationship to chemical parameters. Polish J. Ecol. 55: 7990. Mitchell, E. A. D. 2004. Response of testate amoebae (protozoa) to N and P fertilization in an Arctic wet sedge tundra. Arct. Antarct. Alp. Res. 36: 77-82. Mitchell, E. A. D. and Gilbert, D. 2004. Vertical microdistribution and response to nitrogen deposition of testate amoebae in Sphagnum. J. Eukaryot. Microbiol. 51: 480-490. Mitchell, E. A. D. and Meisterfeld, R. 2005. Taxonomic confusion blurs the debate on cosmopolitanism versus local endemism of free-living protists. Protist 156: 263-267 Mitchell, E. A. D., Borcard, D., Buttler, A. J., Grosvernier, Ph., Gilbert, D., and Gobat, J.-M. 2000a. Horizontal distribution patterns of testate amoebae (protozoa) in a Sphagnum magellanicum carpet. Microbial Ecol. 39: 290-300. Mitchell, E. A. D., Buttler, A., Grosvernier, P., Rydin, H., Albinsson, C., Greenup, A. L., Heijmans, M. M. P. D.,

Chapter 2-5: Protozoa: Peatland Rhizopods

Hoosbeek, M. R., and Saarinen, T. 2000b. Relationships among testate amoebae (protozoa), vegetation and water chemistry in five Sphagnum-dominated peatlands in Europe. New Phytol. 145: 95-106. Mitchell, E. A. D., Buttler, A. J., Warner, B. G., and Gobat, J. M. 1999b. Ecology of testate amoebae (Protozoa: Rhizopoda) in Sphagnum peatlands in the Jura mountains, Switzerland and France. Ecoscience 6: 565-576. Mitchell, E. A. D., Charman, D. J., and Warner, B. G. 2008. Testate amoebae analysis in ecological and paleoecological studies of wetlands: past, present and future. Biodivers. Conserv. 17: 2115-2137. Mitchell, E. A. D., Gilbert, D., Buttler, A., Amblard, C., Grosvernier, P., and Gobat, J.-M. 2003. Structure of microbial communities in Sphagnum peatlands and effect of atmospheric carbon dioxide enrichment. Microbial Ecol. 46: 187-199. Mitchell, E. A. D., Knaap, W. O. van der, Leeuwen, J. F. N. van, Buttler, A., Warner, B. G., and Gobat, J.-M. 2001. The palaeoecological history of the Praz-Rodet bog (Swiss Jura) based on pollen, plant macrofossils and testate amoebae (Protozoa). Holocene 11: 65-80. Mitchell, E. A. D., Payne, R. J., and Lamentowicz, M. 2007. Potential implications of differential preservation of testate amoeba shells for paleoenvironmental reconstruction in peatlands. J. Paleolimnol. 40: 603-618. Nguyen-Viet, H., Gilbert, D., Bernard, N., Mitchell, E. A. D., and Badot, P.-M. 2004. Relationship between atmospheric pollution characterized by NO2 concentrations and testate amoebae density and diversity. Acta Protozool. 43: 233-239. Nguyen-Viet, H., Gilbert, D., Mitchell, E. A. D., Badot, P.-M., and Bernard, N. 2007. Effects of experimental lead pollution on the microbial communities associated with Sphagnum fallax (Bryophyta). Microbial Ecol. 54: 232-241. Nguyen-Viet, H., Bernard, N., Mitchell, E. A. D., Badot, P.-M., and Gilbert, D. 2008. Effect of lead pollution on testate amoebae communities living in Sphagnum fallax: An experimental study. Ecotoxicol. Environ. Safety 69: 130138. Opravilová, V. and Hájek, M. 2006. The variation of testacean assemblages (Rhizopoda) along the complete base-richness gradient in fens: A case study from the Western Carpathians. Acta Protozool. 45: 191-204. Opravilová, V. and Zahrádková, S. 2003. Some information of the testate amoebae of Iceland. Limnologica 33: 131-137. Oye, P. van. 1941. Die Rhizopoden des Sphagnetums bei Krisuvik auf Island. Biologisch Jaarboek Dodonaea (Gent) 8: 284-305. Oye, P. van. 1951. Die Rhizopoden des Sphagnetums bei Krisuvik auf Island. Biologische Jaarboek, pp. 284-327. Paulson, B. 1952-1953. Some rhizopod associations in a Swedish mire. Oikos 4: 151-165. Payne, R. 2007. Laboratory experiments on testate amoebae preservation in peats: implications for palaeoecology and future studies. Protozoology 46: 325-332. Payne, R. J. and Mitchell, E. A. D. 2007. Ecology of testate amoebae from mires in the Central Rhodope Mountains, Greece and development of a transfer function for palaeohydrological reconstruction. Protist 158: 159-171. Payne, R. J., Charman, D. J., Matthews, S., and Eastwood, W. J. 2008. Testate amoebae as palaeohydrological proxies in Sürmene Ağaçbaşi Yaylasi Peatland (Northeast Turkey). Wetlands 28: 311-323. Payne, R. J., Kishaba, K., Blackford, J. J., and Mitchell, E. A. D. 2006. Ecology of testate amoebae (Protista) in south-central

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Alaska peatlands: Building transfer-function models for palaeoenvironmental studies. Holocene 16: 403-414. Robson, T., Ballar, C., Sala, O., Scopel, A., and Caldwell, M. 2001. Biodiversity of microfauna and fungal communities in a Sphagnum bog under two levels of solar UV-B. Abstracts of the 86th meeting of the Ecological Society of America, Madison, WI, 3-8 August 2001. Rose, F. 1953. A survey of the ecology of the British lowland bogs. Proc. Linn. Soc. London 164: 186-211. Ruitenburg, G. J. and Davids, C. 1977. Thecamoeba in succession series of some peat-bog vegetations. Hydrobiol. Bull. (Amsterdam) 11: 22-24. Schnitchen, C., Magyari, E., Tóthmérész, B, Grigorszky, I., and Braun, M. 2003. Micropaleontological observations on a Sphagnum bog in East Carpathian region – testate amoebae (Rhizopoda: Testacea) and their potential use for reconstruction of micro- and macroclimatic changes. Hydrobiologia 506: 45-49. Schönborn, W. 1962. Zur Ökologie der sphagnikolen, bryokolen und terrikolen Testaceen. Limnologica 1: 231-254. Schönborn, W. 1963. Die Stratigraphie lebender Testaceen im Sphagnetum der Hochmoore. Limnologica 1: 315-321. Schönborn, W. 1965. Untersuchungen über die ZoochlorellenSymbiose der Hochmoor-Testaceen. Limnologica 3: 173176. Schoning, K., Charman, D. J., and Wastegoard, S. 2005. Reconstructed water tables from two ombrotrophic mires in eastern central Sweden compared with instrumental meteorological data. Holocene 15: 111-118. Searles, P. S., Flint, S. D., Díaz, S. B., Rousseaux, M. C., Ballaré, C. L. and Caldwell, M. M. 1999. Solar ultraviolet-B radiation influence on Sphagnum bog and Carex fen ecosystems: First field season findings in Tierra del Fuego, Argentina. Global Change Biol. 5: 225-234. Seis, J. 1971. Rhizopodenanalytische untersuchungen an den mooren des pleistozaenen Salzachvorlandgletschers, Vorbericht, Leopoldskroner Moor. Berichte aus dem Haus der Natur in Salzburg, Abteilung B, GeologischMineralogische Sammlungen 2: 10-14. Smith, H. G. and Wilkinson, D. M. 2007. Not all free-living microorganisms have cosmopolitan distributions – the case of Nebela (Apodera) vas Certes (Protozoa: Amoebozoa: Arcellinida). J. Biogeogr. 34: 1822 – 1831. Strüder-Kypke, M. C. and Schönborn, W. 1999. Periphyton and sphagnicolous protists of dystrophic bog lakes (Brandenburg, Germany): II. Characteristic species and trophy of the lakes. Limnologica 29: 407-424. Swindles, G. T. and Roe, H. M. 2007. Examining the dissolution characteristics of testate amoebae (Protozoa: Rhizopoda) in low pH conditions: Implications for peatland palaeoclimate studies. Palaeogeogr. Palaeoclimatol. Palaeoecol. 252: 486496. Thomas, R. 1959. Les Thécamoebiens muscicoles et terricoles: Notions d'écologie générale et comparative. Procès-Verbaux de la Société Linnéenne de Bordeaux 98: 27-53. Tolonen, K. 1966. Stratigraphic and rhizopod analyses on an old raised bog, Varrassuo, in Hollola, South Finland. Ann. Bot. Fenn. 3: 147-166. Tolonen, K. 1994. Ecology of testaceans (Protozoa, Rhizopoda) in Mires in Southern Finland.2. Multivariate-analysis. Arch. Protistenk. 144: 97-112. Tolonen, K., Huttunen, P., and Jungner, H. 1985. Regeneration of two coastal raised bogs in eastern North America: Stratigraphy, radiocarbon dates and rhizopod analysis from

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sea cliffs. Ann. Acad. Sci. Fenn., Ser. A, III. GeologicaGeographica 139: 5-51. Tolonen, K., Warner, B. G., and Vasander, H. 1992. Ecology of testaceans (Protozoa: Rhizopoda) in mires in southern Finland: I. Autecology. Arch. Protistenk. 142: 119-138. Tolonen, K., Warner, B. G., and Vasander, H. 1994. Ecology of testaceans in mires in southern Finland: II. Multivariate analysis. Arch. Protistenk. 144: 97-112. Varga, L. 1956. Adatok a hazai Sphagnum lapok vizi mikrofaunajanaki ismeretéhez. Allatani Köslemények 16: 149-158. Vincke, S., Gremmen, N., Beyens, L., and Vijver, B. Van de. 2004. The moss dwelling testacean fauna of Île de la Possession. Polar Biol. 27: 753-766. Vucetich, M. C. 1975. Tecamebianos muscícolas y esfagnícolas de islas Malvinas (Argentina). Neotropica 21: 11-16. Warner, B. G. 1987. Abundance and diversity of testate amoebae (Rhizopoda, Testacea) in Sphagnum peatlands in southwestern Ontario, Canada. Arch. Protistenk. 133: 173189. Warner, B. G. 1991. Fossil testate amoebae (protozoa) and hydrological history of an ombrotrophic bog in northwestern Ontario, Canada. In: Spigarelli, S.A. (ed.). Proceedings of

an International Symposium on Peat/Peatland Characteristics and Uses. Minnesota, Bemidji State University, pp. 5-14. Warner, B. G. and Charman, D. J. 1994. Holocene changes on a peatland in northwestern Ontario interpreted from testate amoebae (protozoa) analysis. Boreas 23: 270-279. Warner, B. G., Asada, T., and Quinn, N. P. 2007. Seasonal influences on the ecology of testate amoebae (protozoa) in a small Sphagnum peatland in Southern Ontario, Canada. Microbial Ecol. 54: 91-100. Wilmshurst, Janet M. 1998. And in the bog there lived... Protozoan bog dwellers can tell us a lot about past climates. New Zealand Science Monthly Online. Accessed on 28 October 2008 at . Woodland, W. A., Charman, D. J., and Sims, P. C. 1998. Quantitative estimates of water tables and soil moisture in Holocene peatlands from testate amoebae. Holocene 8: 261273. Zygmunt, J., Booth, R., and Jackson, S. 2003. Ecology of testate amoebae in Rocky Mountain peatlands and their application to paleohydrological reconstruction. Abstracts of the 88th meeting of the Ecological Society of America, Savannah, GA, 3-8 August 2003 Accessed on 3 July 2004 at .

Glime, J. M. 2017. Protozoa Ecology. Chapt. 2-6. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. Ebook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 18 July 2020 and available at .

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CHAPTER 2-6 PROTOZOA ECOLOGY TABLE OF CONTENTS General Ecology .................................................................................................................................................. 2-6-2 Epiphytes ..................................................................................................................................................... 2-6-4 Antarctic....................................................................................................................................................... 2-6-4 Nutrient Cycling .................................................................................................................................................. 2-6-5 Habitat Effects..................................................................................................................................................... 2-6-5 Moss Effects on Soil Habitat........................................................................................................................ 2-6-5 Epizoites....................................................................................................................................................... 2-6-5 Soil Crusts .................................................................................................................................................... 2-6-6 Vertical Zonation ................................................................................................................................................ 2-6-7 Zoophagy by Liverworts? ................................................................................................................................... 2-6-8 Dispersal............................................................................................................................................................ 2-6-11 Cosmopolitan .................................................................................................................................................... 2-6-12 Communities as Biological Monitors ................................................................................................................ 2-6-13 Collecting and Sorting....................................................................................................................................... 2-6-14 Collecting ................................................................................................................................................... 2-6-14 Storage and Preservation ............................................................................................................................ 2-6-14 Preservation ........................................................................................................................................ 2-6-14 Long-term Storage of Cysts ................................................................................................................ 2-6-14 Extraction ................................................................................................................................................... 2-6-14 Testate Amoebae................................................................................................................................. 2-6-15 Non-testate Taxa ................................................................................................................................. 2-6-15 Observation ................................................................................................................................................ 2-6-15 Staining ...................................................................................................................................................... 2-6-16 Identification .............................................................................................................................................. 2-6-16 Quantification ............................................................................................................................................ 2-6-16 Summary ........................................................................................................................................................... 2-6-17 Acknowledgments ............................................................................................................................................. 2-6-17 Literature Cited ................................................................................................................................................. 2-6-17

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CHAPTER 2-6 PROTOZOA ECOLOGY

Figure 1. The ciliate protozoan Blepharisma americana inhabits the lobules of the liverwort Pleurozia purpurea. Photo by Sebastian Hess, with permission.

General Ecology Protozoa can probably be found on almost any bryophyte if one just looks carefully (Figure 1). Larger protozoa tend to occur in bog habitats (Chardez 1967; Bovee 1979). As drier habitats are examined, the species are smaller and smaller. Difflugia (Figure 2) species are typical of aquatic mosses; Cyclopyxis species occur on terrestrial mosses. Centropyxis species distribution depends on the habitat, with C. aculeata (Figure 3, Figure 4) in wet locations and C. platystoma in dry ones. Corythion dubium (Figure 5), Assulina muscorum (Figure 6), and Trinema lineare (Figure 7) occur generally on forest mosses (Chardez 1957; Bovee 1979; Beyens et al. 1986), although A. muscorum also is known from the cells of living Sphagnum recurvum (Figure 8) (BioImages 1998). Corythion pulchellum (Figure 9) and Trinema complanatum (Figure 10) occur only on forest mosses (Chardez 1960; Bovee 1979). Nebela collaris (Figure 11), Centropyxis aculeata, and Hyalosphenia papilio (Figure 12) occur on Sphagnum and other bog mosses, but not on forest mosses (Chardez 1960; Chiba & Kato 1969; Bovee 1979).

Figure 2. Difflugia bacillifera with diatoms in the test. Note the small desmid beside it. Photo by Yuuji Tsukii, with permission.

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Figure 6. Assulina muscorum. Photo by Yuuji Tsukii, with permission.

Figure 3. Centropyxis aculeata, a testate amoeba that commonly occurs on bryophyte leaves. Photo courtesy of Javier Martínez Abaigar, with permission.

Figure 7. Test of Trinema lineare. Mitchell, with permission.

Figure 4. Centropyxis aculeata test. Bourland, with permission.

Photo by William

Figure 8. Sphagnum recurvum var. tenue, a peatmoss that supports living protozoa in its hyaline cells. Photo by Jan-Peter Frahm, with permission.

Figure 5. Corythion dubium test. Photo by Yuuji Tsukii, with permission.

Figure 9. Corythion pulchellum. Photo by Yuuji Tsukii, with permission.

Photo by Edward

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Chapter 2-6: Protozoa Ecology

Figure 10. Trinema complanatum. Photo by Yuuji Tsukii, with permission.

Antarctic The role of protozoa is particularly important in the Antarctic. On Elephant Island of the South Shetland Islands in the Antarctic, moss carpets and turf form a major part of the habitat available to protozoa (Smith 1972). Mastigophoran (flagellate) moss inhabitants include 15 species. The Mastigophora are not unique to this habitat. Those that were in most of the moss samples also were in samples of grass/soil, clay, or guano (accumulation of feces). Furthermore, none of the species that was abundant in the other habitats was absent among bryophytes except Tetramitus rostratus, which was abundant only on guano. The Rhizopoda, including the testate amoebae, seemingly avoided the guano on Elephant Island, whereas 16 species occurred in the bryophyte habitats (Smith 1972). Several of those Rhizopoda present in the grass/soil habitat were not found among the moss samples. Fourteen species of Ciliata occurred among mosses.

Figure 11. Nebela collaris. Photo by Yuuji Tsukii, with permission.

Figure 13. Nebela tincta test with living amoeba. Photo by Edward Mitchell, with permission.

Figure 12. Hyalosphenia papilio and H. elegans. by Edward Mitchell, with permission.

Photos

The small number of Elephant Island moss samples (4 in Polytrichum–Chorisodontium turf & 5 in Brachythecium–Calliergon–Drepanocladus carpet) precludes comparison of moss preferences (Smith 1972). The most abundant ciliate, Urotricha agilis (see Figure 14), was abundant in both turf and carpet. In samples of turf, mean numbers per gram of fresh weight ranged 170-4,500. In carpet they ranged 250 to 7,700. On Signey Island species numbers were higher in moss turf (40), whereas on Elephant Island they were higher in moss carpet (37) than in turf.

Protozoa are generally the most numerous invertebrates among the Sphagnum plants (Figure 8; ntham & Porter 1945). In a Canadian study, flagellates were the most numerous, but testate amoebae are often the most numerous. Epiphytes Despite the dryness of aerial habitats, protozoa are common among epiphytic bryophytes, drying and encysting as the bryophytes dry, then reviving, eating, and reproducing when the bryophytes are moist. This habitat may hold many species as yet undiscovered because it is a habitat less frequently studied by protozoologists. Nevertheless, a number of taxa are known from this unique habitat (Golemansky 1967; Casale 1967; Bonnet 1973a, b).

Figure 14. Urotricha platystoma. Photo by Yuuji Tsukii, with permission.

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Nutrient Cycling Protozoa are common predators on bacteria and fungi (Hausmann et al. 2003), having the role of nutrient cyclers (Mitchell et al. 2008). In the Pradeaux peatland in France, the testate Nebela tincta (Figure 13) consumed mostly micro-algae, especially diatoms, associated with mosses (Gilbert et al. 2003). In summer they also consumed large ciliates, rotifers, and other small testate species. Microorganisms collect between leaves and along stems of Sphagnum. When the system is wet, prey organisms are mostly immobile and often dead, but when conditions are drier and the water film is thin, testate fauna are able to ingest more mobile organisms than usual because these prey are slowed down by lack of sufficient free water for rapid swimming. Although we know little about their role among bryophytes, it is likely that at least in peatlands the role of moss-dwelling protozoans in nutrient cycling is significant (Gilbert et al. 1998a, b; Mitchell et al. 2008).

Figure 15. Tardigrade. Photo courtesy of Filipe Osorio.

Habitat Effects When protozoa and other inhabitants live on a host, they can alter the host. Insects are well known for the many forms of galls that develop on the host plant. Gradstein et al. (2018) discovered a white colony of protozoa, resembling gnathifers, in the swollen shoot tips of the liverwort Herbertus sendtneri. This resulted in cessation of the tip growth and subsequent development of innovations below the tip.

Figure 16. Hypsibius oberhaeuseri with Pyxidium tardigradum growing as a symphoriont. Redrawn from Van Der Land 1964.

Moss Effects on Soil Habitat The presence of mosses also affects the microorganisms found in the underlying soil. Miroschnichenko and coworkers (1975) found that the greatest numbers of micro-organisms were under mosses (compared to other soil substrata) in a community in Russia, and Smith and Headland (1983) found similar results for testate rhizopods on the sub-Antarctic island of South Georgia. Smith (1974a, 1986) found protozoa living among the bryophytes in the South Orkney Islands and Adelaide Island of the Antarctic. Ingole and Parulekar (1990) found that the faunal density, including protozoa, was high in mossassociated sediments. These micro-organisms may account for the ability of some macrofauna to remain within the moss mat throughout a major part of their development by serving as a food source (Smith 1974a, 1986). Epizoites Some of the fauna, such as Pyxidium tardigradum (Figure 17), an epizoite, are hitch-hikers. This protozoan is recorded as a symphoriont (organism carried by and often dispersed by its host) on two species of tardigrades (Figure 15) [Hypsibius oberhaeuseri (Figure 16) and Milnesium tardigradum] that live among mosses (Land 1964; Morgan 1976). It can be so common on them (up to 35, but more typically 1-3) as to have negative effects on the tardigrade host that must expend extra energy to carry them around (Vicente et al. 2008). For this reason, Vicente et al. (2008) suggest that it should perhaps be considered a parasite.

Figure 17. Pyxidium tardigradum, a symphoriont. Redrawn from Van Der Land 1964.

tardigrade

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Soil Crusts Protozoan communities associated with cryptogamic soil crusts (Figure 18) have hardly been studied. In a study of only five crusts in southeastern Utah, Bamforth (2008) found 28 species of amoebae, 45 ciliates, and 19 testate amoebae. The number of amoebae ranged 680-2500, ciliates 20-460, and testate amoebae 2400-2500 per gram dry mass of crust. As crusts succeeded from Microcoleus (Cyanobacteria) to lichens to bryophytes, numbers of protozoa increased, perhaps reflecting longer periods of internal moisture in the crusts. Predominant taxa are somewhat different from cosmopolitan ones we have seen elsewhere, comprised mostly of Acanthamoeba (Figure 19), Hartmanella (Figure 20), Vahlkampfidae (Figure 21), two species of Colpoda (Figure 22), several other colpodids, Polyhymenophora sp., and species of Cryptodifflugia (Figure 23) and Difflugiella.

Figure 21. permission.

Valkampfia.

Photo by Yuuji Tsukii, with

Figure 18. Soil crust with the moss Syntrichia ruralis. Photo by Michael Lüth, with permission.

Figure 19. Acanthamoeba showing ingested carmine particles. Photo by Akira Kihara, with permission.

Figure 20. permission.

Hartmanella.

Photo by Yuuji Tsukii, with

Figure 22. Colpoda aspera. Photos by William Bourland, with permission.

Chapter 2-6: Protozoa Ecology

Figure 23. Cryptodifflugia ovaliformis on an alga filament. Photo by Yuuji Tsukii, with permission.

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Nitrogen distribution affects the vertical distribution of at least some testate amoebae in Sphagnum communities, but nitrogen availability does not seem important for most testate amoebae in the upper centimeters of Sphagnum mats in the Swiss Jura Mountains (Mitchell & Gilbert 2004). There were 22 testate taxa among these mosses, although mean diversity of a typical sample was only 6.6. The species richness increased with depth. The moss-dwelling Assulina muscorum (Figure 25) was most abundant in the top 0-1 cm; Phryganella acropodia, Heleopera rosea (see Figure 26), and Nebela militaris (Figure 27) were the most abundant taxa at 3-5 cm depth. In this case, species richness increased with depth in the mat. Only Bullinularia indica (Figure 28) appeared to be more abundant in plots fertilized with nitrogen.

Vertical Zonation Bryophyte suitability as a protozoan habitat differs in both time and space. Bryophytes offer a vertical series of habitats (Figure 24) that differ in temperature, moisture, and light, and presumably food quality and quantity. Horizontally, the substrate or height above the water table can differ, causing species differences. Hence, the microorganisms distribute themselves in different communities both seasonally and spatially, particularly in the Sphagnum peatlands (Schönborn 1963; Heal 1964; Meisterfeld 1977; Mazei and Tsyganov 2007). Figure 25. Assulina muscorum. Photo by Yuuji Tsukii, with permission.

Figure 24. Sphagnum subnitens showing tips and lower branches that create habitat zones for protozoa. Photo by Michael Lüth, with permission.

Figure 26. Heleopera sylvatica showing pseudopods. Photo by Yuuji Tsukii, with permission.

Spaces: Several studies indicate that the sizes of spaces within the bryophyte habitat influence the sizes of organisms and influence the available food (Dalenius 1962; Corbet 1973; Bovee 1979; Robson et al. 2001). Capillary spaces among branches and leaves hold water. Gilbert et al. (2003) suggested that as the Sphagnum becomes drier, ciliate protozoa are easier to catch for food because the thin film of water slows them down. As the moss becomes too dry, rather than migrating to lower, moister areas, many of these taxa, like several invertebrate groups, can encyst, permitting them to survive desiccation (Heal 1962; Gerson 1982). And when the moss resumes activity under the stimulation of rain (or fog), the rhizopods do likewise. Nitrogen: Nitrogen from guano seemingly deterred all the testate amoebae on Elephant Island (Smith 1972).

Figure 27. Nebela militaris. Photo by Yuuji Tsukii, with permission.

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Figure 28. Test of Bullinularia indica. Photo by Edward Mitchell, with permission.

Temperature: The Antarctic fauna is dominated by moss-dwelling micro-organisms, including protozoa, rotifers, nematodes, and tardigrades (Schwarz et al. 1993). Here, temperature may play a role as important as that of moisture. This need for adequate heat results in a vertical zonation of the fauna. For example, at the Canada Glacier, in southern Victoria Land, the majority of moss-dwelling organisms were in the top 5 mm in the post-melt samples, rather than in the pre-melt samples. However, while temperatures differed, so did the available moisture, making it difficult to determine controlling factors. Light: As one might expect, light determines the absence of protozoa with chlorophyllous symbionts in the lower strata (Chacharonis 1956). Only those surface species contain chlorophyll, either as symbiotic algae or that of their own possession. However, some with chlorophyllous symbionts may occur as deep as 6-10 cm in Sphagnum mats (Richardson 1981). Of the 27 species lacking symbionts in a Sphagnum mat, all but two exhibited maximum abundance below 6 cm. But even within the first 5 cm, vertical zonation exists. Mitchell and Gilbert (2004) demonstrated a significant difference in number of species between the first 3 cm and the 3-5 cm depth in Polytrichum strictum (Figure 29) of a Swiss peatland (Figure 30).

Figure 30. Vertical distribution of species richness of testate amoebae in a Polytrichum strictum "carpet" of a Swiss peatland. Redrawn from Mitchell & Gilbert 2004.

Community Differences: As for a number of other moss habitats, the Sphagnum peat mat provides vertical differences in microhabitat that are further expressed as vertical community differences (Meisterfeld 1977; StrüderKypke 1999; Mitchell et al. 2000). Strüder-Kypke found that even in the upper 30 cm of the mat, two very different protistan communities are dictated by the strong vertical zonation. Both light and nutrients differ, causing the upper region to support a denser colonization, mostly of autotrophic cryptomonads and vagile ciliates (able to move about or disperse in a given environment). On the other hand, deeper samples exhibited heterotrophic flagellates and sessile peritrich ciliates. Presence of testate amoebae at greater depths within the moss mat does not always indicate a retreat to a location of greater moisture. Schönborn (1977) demonstrated that 15% of the shells can be transported to lower depths by 550 mm rainfall, but 400 mm generally does not seem to cause a noticeable downward loss.

Zoophagy by Liverworts?

Figure 29. Polytrichum strictum. Photo by Michael Lüth, with permission.

Carnivorous plants are well known among the flowering plants, but the ability of bryophytes to attract and trap organisms has been questionable. Who would guess that these seemingly primitive organisms can attract their own prey? But one interpretation is that the leafy liverwort genera Colura (Figure 31, Figure 32) and Pleurozia (Figure 33) have lobules (water sacs) that do just that (Hess et al. 2005). And this is not an isolated example. In the Aberdare Mountains, Kenya, Chuah-Petiot and Pócs (2003) found many protozoa inhabiting the lobules of the epiphytic Colura kilimanjarica (Figure 31, Figure 32).

Chapter 2-6: Protozoa Ecology

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Figure 33. Underside of Pleurozia purpurea showing lobules where invertebrates often live – and die. Photo by Sebastian Hess, with permission.

Figure 31. Upper: The leafy liverwort, Colura. Lower: This lobule of Colura houses the ciliate protozoan Blepharisma americana. Photos by Jan-Peter Frahm, with permission.

Figure 32. Upper: SEM of lobule of Colura. Lower: Living lobule. These lobules of Colura are inhabited by the reddish ciliate protozoan Blepharisma americana. Photos by Jan-Peter Frahm, with permission.

Lobules are usually considered to be water storage organs. However, in these genera, they might also serve as traps. Goebel (1888, 1893, 1915) did not consider it likely that these were real traps. He argued that insectivorous plants have attractants in order to lure their prey into their traps. Although the lobule resembles the trap of the bladderwort, Utricularia, Goebel argued that that does not mean it is used the same way. He furthermore argued that the benefit gained by the excrement from animals (and dead animals?) would be less than that gained from the water. Since having the animals does not preclude also providing a water reservoir, it would seem that zoophagy would simply be an added benefit. Schiffner (1906) even reported chironomid larvae in the lobules, suggesting an even larger source of fecal matter. But the openings in Pleurozia are small, only about 300 µm, and closed by a round "lid" of hyaline cells (Hess et al. 2005). What causes these organisms to enter in the first place?

Figure 34. Pleurozia purpurea, a leafy liverwort with lobules that can house a variety of invertebrates, including the ciliate Blepharisma americana. Photo by Sebastian Hess, with permission.

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see if the dispersion of the protozoan remained random. Indeed, the protozoa gradually accumulated around the Pleurozia! Within only 30 minutes, 86% of the lobules contained the protozoa. After several hours, up to 16 protozoans were trapped, and further observation failed to reveal any that escaped. The mode of attraction is only speculation. Barthlott et al. (2000) found that older parts of Colura were more effective at attracting Blepharisma americana (Figure 37, Figure 38) than were younger parts, suggesting that concentrations of bacteria may have been a factor. In fact, in experiments on Colura, Barthlott et al. (2000) found that B. americana moves over the bryophyte surface "like a vacuum cleaner," devouring the bacteria.

Figure 37. A stained Blepharisma americana. Photo by Yuuji Tsukii, with permission. Figure 35. Upper: Lobule of Pleurozia purpurea showing lid. Photo by Sebastian Hess, with permission. Lower: Lobule redrawn from Hess et al. (2005). This lobule of Pleurozia purpurea serves as home and apparently ultimately as a trap for a wide range of protozoa and invertebrates.

Barthlott et al. (2000), using feeding experiments with the ciliate protozoan Blepharisma americana (Figure 1, Figure 36-Figure 38), demonstrated that Colura does indeed catch protozoa with its lobules. Hess and coworkers (2005) set out to determine if Pleurozia purpurea (Figure 33-Figure 35) is likewise carnivorous.

The shade provided by the plants could also contribute to the higher concentrations of protozoa near the branches of Pleurozia purpurea (Hess et al. 2005), but if so, the liverwort would probably be less effective as a refuge in the field where other mosses were also present. Hess and coworkers (2005) claim that the large number of organisms in the lobules in such a short time is too great to be attributed to chance. However, they fail to provide any statistical evidence or probability to support this claim, for example, alternative liverworts or mosses. They furthermore state that the organisms die there, but they provide no data on the deaths of the organisms. They do point out that there is no direct evidence that any nutrients provided by the organisms are used by the liverworts, but there is likewise no evidence to the contrary. In any case, the liverworts could benefit from the cleaning of bacteria that block light and compete for nutrients.

Figure 36. The ciliate Blepharisma americana that inhabits "zoophagous" liverworts. Photo by Yuuji Tsukii, with permission.

Again using Blepharisma americana, a cohabitant of Sphagnum mats with Pleurozia purpurea, Hess et al. (2005) performed dozens of experiments in Petri dishes to

Figure 38. SEM photo of Blepharisma demonstrating small cell on top and large, cannibalistic cell below. Under starvation conditions, larger individuals become cannibalistic. Photo by Pauline Gould, with permission.

Chapter 2-6: Protozoa Ecology

Zoophagy is the process of eating animals (phag = eat, devour; Hanson 1962; Lincoln et al. 1998). There is a fine distinction in what constitutes just eating compared to true carnivory, wherein living organisms are killed (or not) and digested. In this case, it seems that the animals may be trapped, but there is no real proof that they are consumed by the plant. Does admitting the animals into the trap (lobule) then make the liverworts zoophagous? Hess et al. (2005) argue that animals die in the traps and subsequently release their cell contents, bursting in the case of Blepharisma americana. These dead animals are then decomposed by bacteria. Surely some of the nutrients released are absorbed by the liverworts. Is this not a process parallel to that of the pitcher plant Sarracenia purpurea? Many so-called carnivorous plants, like S. purpurea, seem to lack enzymes to digest all or some of the parts of their prey and depend on resident bacteria to accomplish the task. With this broad definition of carnivory, could we not call the liverworts carnivorous? I think I want more data on whether this is a chance event or true trapping before I make that claim. Such experiments would need controls of leafy liverworts with no "traps" to see if the protozoa simply accumulate wherever there is shelter. On the other hand, I wonder how many leafy liverworts with locules provide preferred housing for protozoa.

Dispersal For any organism to succeed, it must have a means of dispersal. Protozoans can't go very far on their own. They are too small to crawl far on pseudopods or paddle their way with a flagellum or cilia, the common means of transportation for the majority of protozoan moss dwellers. But they can travel reasonable distances as passengers on the mosses, riding on fragments that establish a new home where they land. Sudzuki (1972) conducted experiments using electric fans to determine the success of wind as a dispersal agent, using mosses as one of the sources of invertebrate fauna. He found that the smaller organisms – micro-organisms, including protozoa, were easily dispersed by light breezes as well as wind. Larger organisms such as gastrotrichs, flatworms, rotifers, nematodes, oligochaetes, tardigrades, crustaceans, and arachnomorphs, on the other hand, rarely were dispersed at wind velocities of less than 2 m per second [tornadoes are generally 27-130 m per second (Allaby 1997)]. In the field, colonization progressed from flagellates to ciliates to rhizopods, suggesting that passive dispersal was not the only factor controlling their colonization rates. Once an organism becomes airborne, turbulent air may take them 3,000 to even 17,000 m on thermal drafts, with winds carrying them much higher and farther (Maguire 1963). Puschkarew (1913) found that protozoan cysts average about 2.5 per cubic meter, making these organisms readily available for dispersal and colonization on suitable bryophytes. Smith (1974b) likewise considered that the mosses themselves served as dispersal agents for the protozoa. In particular, moss invasions of volcanic tephra on Deception Island in the Antarctic greatly increased the protozoan fauna. Not only do the mosses provide a great increase in suitable niches, but since they were most likely colonized

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by protozoa in their former locations, fragments arriving on the island could easily carry communities of fauna as passengers. Rain can carry many algae and protozoa (Maguire 1963). Rain-borne organisms seem to originate predominantly from splash, typically from plants and soil, and do not travel far vertically, so that mechanism is most likely only suitable for local habitat travel. In streams, the water movement itself serves as an effective dispersal agent, and aerial dispersal from waterfalls and rapids can carry algae and other Aufwuchs to new locations. Raccoons are very effective in carrying whole communities of organisms, particularly protozoa, and can accomplish distances of at least 60 meters (Maguire 1963). Both terrestrial and aquatic birds contribute to dispersal, and other mammals contribute, but their relative role is not known. Several scientists have discussed the dispersal of micro-organisms by insects (Maguire 1963; Parsons et al. 1966). Such mechanisms could easily contribute to the colonization of bryophytes by their micro-inhabitants. The many aquatic insect inhabitants will be discussed in an upcoming chapter. Consider the activity of insects among bryophytes, especially in streams, and their subsequent relocation due to swimming or stream drift. The Aufwuchs could easily be carried from one location to another by these mobile inhabitants (Figure 39). Emerging insects may also swipe micro-organisms trapped by the surface tension and carry them to resting locations, including bryophytes, on land.

Figure 39. Dragonfly Aeshna grandis female ovipositing and exposing herself to possible transport of protozoa. Photo by David Kitching, with permission.

Although few studies seem to have directly addressed the dispersal of micro-organisms by insects to bryophytes, we can infer at least some possibilities from more general studies on dispersal by insects. Maguire (1963) examined the distance both horizontally and vertically to which organisms were dispersed from a pond in Texas and another in Colorado. Dragonflies (Figure 39) and wasps, in particular, carried several species of protozoa and one species of rotifer. Parsons et al. (1966) found amoeboid and other protozoan cysts on adult Odonata, suggesting the possibility of a relatively long dispersal range. Odonata in

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a short-term experiment dispersed up to 860 m to the farthest pond in the experiment (Conrad et al. 1999). Michiels and Dhondt (1991) estimated that 80% of adult dragonfly Sympetrum danae had migrated 1.75 km or more to their study site. But more importantly, evidence suggests they can migrate 3500 km or more across the Indian Ocean (Anderson 2009). This and other longdistance migrations provide a potential yearly means of dispersal for the micro-organisms.

missing from the sites in Switzerland, Alaska, Sweden, Finland, Netherlands, Britain, Bulgaria, and North America as summarized in Table 1 of Chapter 2-2. The epiphytic community had 34 taxa in 13 genera, whereas the soil mosses had 31 taxa in 13 genera.

Cosmopolitan 'Everything is everywhere, but, the environment selects' (in Wit & Bouvier 2006; O'Malley 2008). This statement, often called the Baas Becking Principle, has been applied to microscopic organisms that are globally distributed by high dispersal, and that lack biogeographic patterns (Fontaneto et al. 2008). But Wit and Bouvier made it clear that the original hypothesis "did not disregard the biogeography of free-living microorganisms." Finlay et al. (1996) extend the concept to suggest global species diversity is inversely related to body size. Therefore, the huge number of protist individuals makes global dispersal inevitable through normal events such as ocean circulations, groundwater connections, hurricanes, damp fur, dust storms, etc. (Weinbauer & Rassoulzadegan 2003). This argument is supported by the fact that the estimated number of free-living ciliates is about 3000, whereas there are about 10,000 species of birds and 120,000 species of Lepidoptera (butterflies and moths) (Lawton 1998). The concept of global distribution describes well the major protozoa associated with bryophytes. This concept does not preclude, however, the presence of cryptic species that differ in less recognizable traits (Richards et al. 2005; Fontaneto & Hortal 2008; Fontaneto et al. 2008; Kooistra et al. 2008), and in recent detailed studies distinct genetic species have been found in disparate parts of the world (Telford et al. 2006; Fontaneto et al. 2008; Kooistra et al. 2008). One consideration to support "everything is everywhere" is the small number of species of protozoa relative to 750,000 species of insects and 280,000 species of other animals (Papke & Ward 2004). Morphological data support the concept that dispersal is worldwide, suggesting there would be fewer than 5000 morphological protozoan species. Could this also be the explanation for the small number of bryophytes relative to other plants? In both cases, molecular evidence is starting to suggest that there may be cryptic species with genetic differences that are not expressed morphologically (Logares 2006), revealing distributions that are much more restricted. Bryophyte protozoan communities are remarkably similar no matter where the bryophytes occur and consist primarily of cosmopolitan species. Davidova (2008) compared the testacean communities of epiphytic bryophytes to those of soil bryophytes in Strandzha Natural Park, South-Eastern Bulgaria, and found them to be quite similar in their taxonomic richness, species diversity, and community structure. The most common taxa in both habitats were Centropyxis aerophila var. sphagnicola, C. aerophila (Figure 40), Phryganella hemisphaerica, Euglypha rotunda (Figure 41), Corythion dubium (Figure 5), Trinema enchelys (Figure 42), and T. lineare (Figure 7). Among these, only Phryganella hemisphaerica is

Figure 40. Centropyxis aerophila test. Tsukii, with permission.

Photo by Yuuji

Figure 41. Euglypha rotunda. Photo by Yuuji Tsukii, with permission.

Figure 42. Trinema enchelys. Photo by Yuuji Tsukii, with permission.

The moss-dweller Nebela (Apodera) vas (Figure 43) has been touted to refute the Baas Becking Principle (Mitchell & Meisterfeld 2005; Smith & Wilkinson 2007). In 89 collections, representing 25 publications, mosses represented 59% of its habitat, with Sphagnum being the most common (Smith & Wilkinson 2007). Its distribution

Chapter 2-6: Protozoa Ecology

is throughout the equatorial region at high altitudes, southern cool-temperate, and sub-Antarctic zones, but it is conspicuously absent in the Holarctic northern hemisphere. Its absence from hundreds of samples from seemingly suitable habitats in the northern hemisphere support the contention that its absence is not a fluke of sampling (Mitchell & Meisterfeld 2005) This distribution is definitely not cosmopolitan, despite its wide pH range (3.86.5) (Smith & Wilkinson 2007). Although it has a rather defined climatic range (temperate to sub-Antarctic), its absence in this climate throughout most of the more frequently studied northern hemisphere cannot support the concept of "everything is everywhere." Evidence such as this has been used to argue that micro-organisms are dispersed following the same principles as macroorganisms (BioMed Central 2007). Genetic differences that are not detectable from morphology suggest that global diversity of micro-organisms may be greater than has been suspected (BioMed Central 2007; Fontaneto et al. 2008). Such evidence suggests that care is needed in assigning names to microbial/protozoan collections.

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Corythion (Figure 5, Figure 9), Euglypha (Figure 41), and Heleopera (Figure 26), as well as Euglena (Figure 44) and Cyanobacteria, in a Sphagnum bog of Tierra del Fuego, South America, were sensitive to UV-B radiation (Robson et al. 2001). But surprisingly the testate amoebae and rotifers were significantly more abundant and had greater species diversity under current levels of UV-B radiation than those that received reduced UV-B. The fungal component likewise had significantly greater abundance and species diversity under the current dosage than under the reduced dosage.

Figure 44. Euglena mutabilis, a common euglenoid among bryophytes, particularly in peatlands. Photo by Yuuji Tsukii, with permission.

Figure 43. SEM view of Apodera (Nebela) vas showing test. Photo by Edward Mitchell, with permission.

Jenkins et al. (2008) have tested the size hypothesis, using 795 data values on dispersal units from published research. They found that active dispersal vs. passive dispersal matters greatly, with active dispersers dispersing significantly farther (p1 cm thick) of mosses, frequently Paraleucobryum sp. (Figure 161) and sometimes species of the leafy liverwort Mylia (Figure 162). This association is typically enriched with detritus. Differing from Barbeyella minutissima (Figure 80) and Colloderma oculatum (Figure 27) that occur almost entirely on the decorticated spruce and fir logs that have coverings of slimy algae and Cyanobacteria, Cribraria cancellata (Figure 163) and Diderma montanum (Figure 164) tend to occur in the cooler valley bottoms, where they produce sporangia on moderately decayed wood of spruce and beech, often on logs with mossy, loose bark.

Figure 163. Cribraria cancellata sporangia, a species that occurs on moderately decayed wood of spruce and beech, often on logs with mossy, loose bark. Photo by Clive Shirley, The Hidden Forest, with permission.

Figure 164. Diderma montanum sporangia, a species that occurs on moderately decayed wood of spruce and beech, often on logs with mossy, loose bark. Photo by Alain Michaud, The Eumycetozoan Project, DiscoverLife.org, with online permission.

Figure 161. Paraleucobryum longifolium, a moss of the moist wood stage of mostly decorticated logs. Photo by Hermann Schachner, through Creative Commons.

Figure 162. Mylia taylorii; the genus Mylia often occurs on the moist wood stage of the mostly decorticated logs. Photo by David T. Holyoak, with permission.

Stephenson and Studlar (1985) concluded that Barbeyella minutissima (Figure 80) and Lepidoderma tigrinum (Figure 83) are bryophilous, being almost invariably associated with bryophytes, and in particular with leafy liverworts. Schnittler et al. (2000) examined collections from 27 localities in the Northern Hemisphere. They concluded that these two species are restricted to decorticated coniferous wood covered by 40-100% leafy liverworts, based on 41 collections. They furthermore noted the importance of a "thin, slimy layer" of algae. Stojanowska and Panek (2004) reported a number of bryophyte-slime mold-log associations from a nature reserve in southwest Poland. Cribraria vulgaris (Figure 78) and Lycogala epidendrum (Figure 67) occur there on moss-covered stumps and logs. Fuligo septica (Figure 1, Figure 66), Lycogala exiguum (Figure 139-Figure 140), Metatrichia vesparia (Figure 141), Stemonitis fusca (Figure 69), S. pallida (Figure 57), Trichia botrytis (Figure 25), T. persimilis (Figure 113-Figure 114), T. varia (Figure 72), and Tubifera ferruginosa (Figure 73) occur on bryophyte-covered stumps. Diderma radiatum (Figure 165-Figure 166) occurs on stumps overgrown with the moss Brachythecium rutabulum (Figure 167). Arcyria cinerea (Figure 32-Figure 33), A. denudata (Figure 62Figure 63), Physarum compressum (Figure 168-Figure 169), Physarum gyrosum (Figure 170-Figure 171), Stemonitis axifera (Figure 24), and Trichia scabra (Figure 92) occur on bryophyte-covered logs. Lepidoderma tigrinum (Figure 83) occurs on decaying logs densely

Chapter 3-3: Slime Molds: Ecology and Habitats – Bark and Logs

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overgrown with Dicranum montanum (Figure 149) (see also Neubert et al. 1993), whereas Badhamia panicea (Figure 172-Figure 173) occurs on bark of a recent log with Brachythecium rutabulum. Reticularia lobata (syn.=Enteridium lobatum; Figure 174) occurs on bryophyte-covered conifer wood. They also mentioned that Lamproderma columbinum (Figure 42) occurs on Tetraphis pellucida (Figure 175), a moss species most typical of decaying stumps, but that also occurs on rocks. The co-occurrence of particular slime molds with specific mosses may reflect a preference of both for the same microclimate.

Figure 168. Physarum compressum on bryophytes. Photo courtesy of Sarah Lloyd.

Figure 165. Diderma radiatum sporangia on log with mosses. Photo by Clive Shirley, The Hidden Forest, with permission.

Figure 169. Physarum compressum fruiting. Photo by Alain Michaud, The Eumycetozoan Project, DiscoverLife.org, with online permission.

Figure 166. Diderma radiatum sporangia, ready for dispersal. Photo from Myxotropic, through Creative Commons.

Figure 167. Brachythecium rutabulum, a common substrate for Diderma radiatum. Photo by Arnoldius, through Creative Commons.

Figure 170. Physarum gyrosum fruiting; this slime mold can be found on logs covered with bryophytes. Photo by Ray Simons, The Eumycetozoan Project, DiscoverLife.org, with online permission.

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Chapter 3-3: Slime Molds: Ecology and Habitats – Bark and Logs

Figure 171. Physarum gyrosum fruiting and dispersing spores. Photo by Dmitry Leontyev, The Eumycetozoan Project, DiscoverLife.org, with online permission.

Figure 172. Badhamia panicea sporangia, a species that occurs on bark of a recent log with the moss Brachythecium rutabulum. Photo by Alain Michaud, The Eumycetozoan Project, DiscoverLife.org, with online permission.

Figure 174. Reticularia lobata, a species of bryophytecovered conifer wood. Photo from The Eumycetozoan Project, DiscoverLife.org, with online permission.

Figure 175. Tetraphis pellucida, a moss that is sometimes a substrate for the slime mold Lamproderma columbinum. Photo by Hermann Schachner, through Creative Commons.

Summary

Figure 173. Badhamia panicea sporangia. Photo by Alain Michaud, The Eumycetozoan Project, DiscoverLife.org, with online permission.

Bark and logs are the two most common substrata for slime molds. And both of these substrates frequently have bryophytes on them. The motile slime molds therefore encounter bryophytes as they move about and may traverse them or stay and form sporangia. On logs in particular, leafy liverworts are common, and these seem to be suitable substrates for a number of slime molds. In some cases, the underlying algae might contribute to this association, providing fixed nitrogen or food. Slime molds that move upward and into the light to produce sporangia may gain some advantage on the slightly elevated bryophytes. This positioning can provide greater access to dispersal agents, including wind and invertebrates. Nevertheless, the bryophytes used are of low stature, with smooth mats being the most frequent. Diderma corrugatum seems to be restricted to moss-covered bark, whereas D. chondrioderma seems only to prefer it. Some of the slime molds seem to be confined to liverworts, including Barbeyella

Chapter 3-3: Slime Molds: Ecology and Habitats – Bark and Logs

minutissima on logs, Licea bryophila on bark, and Licea gloederma on bark. Licea parasitica seems to prefer mosses in its microcyst stage. Colloderma oculatum, Lamproderma columbinum, and Lepidoderma tigrinum are common only associated with Barbeyella minutissima on bryophyte-covered logs, especially with the liverwort Nowellia curvifolia. On the other hand, most of the bryophyte dwellers seem to be accidentals – generalists that tolerate the substrate with no preference for it. Others occur on mossy logs or bark, but not directly on the bryophytes. In some cases, the slime mold seems to start on bark and invade the bryophyte. In other cases, it germinates on the bryophyte and moves onto the bark or wood. In the latter case, the bryophyte might benefit from the greater moisture in the bryophyte mat, in addition to the ability of the bryophyte to trap the spores. Both of bark and logs have periods of drying out, especially tree boles. The slime molds and mosses are both tolerant of these events, but mosses are able to slow the drying process due to their capillary spaces. In addition to moisture, pH seems to be important in separating substrata among slime mold species. Decay stages are likewise important, with different stages providing different moisture levels, but also typically having more bryophytes as they decay more. Slime molds on logs with bryophytes are often also associated with algae and Cyanobacteria, especially Chroococcus tenax, Aphanothece saxicola, and Chlorococcum humicola.

Acknowledgments Marianne Meyer and Isabelle Charissou were very helpful in providing me with pictures of slime molds on mosses and Marianne helped me with identification of some images contributed by others. Harold W. Keller provided additional information and images to support the story on Diachea arboricola.

Literature Cited Adamonyte, G. 2000. New data on Estonian myxomycete biota. Folia Cryptog. Estonica 36: 7-9. Adamonyté, G. 2007. Myxomycetes of the genus Clastoderma in Lithuania. Bot. Lithuanica 13(1): 27-32. Brooks, T. E., Keller, H. W., and Chassain, M. 1977. Corticolous Myxomycetes, VI. A new species of Diderma. Mycologia 69: 179-184. Castillo, A., Moreno, G., and Illana, C. 2009. Myxomycetes from Cabañeros National Park (Spain). Bol. Soc. Micol. Madrid 33: 149-170. Clissmann, F., Fiore-Donno, A. M., Hoppe, B., Krüger, D., Kahl, T., Unterseher, M., and Schnittler, M. 2015. First insight into dead wood protistan diversity: A molecular sampling of bright-spored Myxomycetes (Amoebozoa, slime-moulds) in decaying beech logs. FEMS Microbiol. Ecol. 91(6): fiv050. Coker, P. D. 1966. The destruction of bryophytes by lichens, fungi, Myxomycetes and algae. Trans. Brit. Bryol. Soc. 5: 142-143.

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Compagno, R., Gargano, M. L., La Rosa, A., and Venturella, G. 2016. A contribution to the knowledge of Myxomycetes diversity in volcanic islands. Plant Biosyst. 150: 776-786. Critchfield, R. L. and Demaree, R. S. 1991. Annotated checklist of California Myxomycetes. Madroño 38: 45-56. Döbbeler, P. and Nannenga-Bremekamp, N. E. 1979. Licea gloeoderma, ein neuer Myxomycet aus Bayern. Zeits. Mykol. 45: 235-238. Doidge, E. M. 1950. Myxomycetes. Bothalia 5(1): 83-94. Dudka, I. O. and Romanenko, K. O. 2006. Co-existence and interaction between Myxomycetes and other organisms in shared niches. Acta Mycol. 41: 99-112. Eliasson, U. 1980. Patterns of occurrence of Myxomycetes in a spruce forest in South Sweden. Ecography 4: 20-31. Eliasson, U. H. and Gilert, E. 2007. Additions to the Swedish myxomycete biota. Karstenia 47: 29-36. Everhart, S. E. and Keller, H. W. 2008. Life history strategies of corticolous Myxomycetes: the life cycle, plasmodial types, fruiting bodies, and taxonomic orders. Fungal Divers. 29: 116. Everhart, S. E., Ely, J. S., Keller, H. W. 2009. Evaluation of tree canopy epiphytes and bark characteristics associated with the presence of corticolous Myxomycetes. Botany 87: 509-517. Farr, M. L. 1979. Notes on Myxomycetes II. New taxa and records. Nova Hedw. 31: 103-118. Gilert, E. and Neuendorf, M. 1991. A new species of Lamproderma (Myxomycetes) found in Java. Nord. J. Bot. 10: 661-664. Gradstein, S. R., Reenen, G. B. A. van, and Griffin, D. III. 1989. Species richness and origin of the bryophyte flora of the Colombian Andes. Acta Bot. Neerl. 38: 439-448. Gray, W. D. and Alexopoulos, C. J. 1968. Biology of the Myxomycetes. The Ronald Press, Co., New York. Greene, H. C. 1929. Myxomycetes of western Washington. Mycologia 21(5): 261-273. Haan, M. de. 2016. Myxomycetes growing on epiphytic bryophytes: An opportunity. Sterbeeckia 34: 47-61. Hagelstein, R. 1941. Notes on the Mycetozoa V. Mycologia 33: 294-309. Härkönen, M. 1977. Corticolous Myxomycetes in three different habitats in southern Finland. Karstenia 17: 19-32. Härkönen, M., Rikkinen, J., Ukkola, T., Enroth, J., Virtanen, V., Jaaskelaiiinena, K., Rinnea, E., Hiltunena, L., Piippo, S., and He, X. 2004. Corticolous Myxomycetes and other epiphytic cryptogams on seven native tree species in Hunan Province, China. Syst. Geogr. Plants 74: 189-198. Ing, B. 1967. Myxomycetes as food for other organisms. Proc. S. London Entomol. Nat. Hist. Soc. 1967: 18-23. Ing, B. 1982. Provisional atlas of the Myxomycetes of the British Isles. Biological Records Centre, Institute of Terrestrial Ecology, Monks Wood Experimental Station, Huntingdon, UK. Ing, B. 1983. A ravine association of Myxomycetes. J. Biogeogr. 10: 299-306. Ing, B. 1994. The phytosociology of Myxomycetes. Tansley review no. 62. New Phytol. 126: 175-201. Johannesen, E. W. 1984. New and interesting Myxomycetes from Norway. Nordic J. Bot. 4: 513-520. Joshaghani, A. A., Falahian, F., Nejadsattari, T., Khavarinejad, R. A., and Saadatmand, S. 2013. Two new Myxomycetes records for Iran mycoflora. Pakistan J. Bot. 45: 1813-1816. Kaiser, G. B. 1913. Slime, mould growing on a moss. Bryologist 16: 45.

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Keller, H. W. 2019. Student Team-Based Tree Canopy Biodiversity Research in Great Smoky Mountains National Park. Plant Science Bulletin 65: 28-37. Keller, H. W. and Barfield, K. M. 2017. Great Smokey Mountains National Park: The people's park. Fungi 102: 4464. Keller, H. and Braun, K. 1999. Myxomycetes of Ohio: Their systematics, biology and use in teaching. Ohio Biol. Surv. Bull. New Ser. 13, No. 2. Keller, H. W. and Brooks, T. E. 1975. Corticolous Myxomycetes III: A new species of Badhamia. Mycologia 67: 1218-1222. Keller, H. W. and Skrabal, M. 2002. Discovery of a new obligate tree canopy myxomycete in the Great Smoky Mountains National Park. Inoculum 53(2): 1-4. Keller, H. W., Skrabal, M., Eliasson, U. H., and Gaither, T. W. 2004. Tree canopy biodiversity in the Great Smoky Mountains National Park: Ecological and developmental observations of a new myxomycete species of Diachea. Mycologia. 96: 537-547. Kowalski, D. T. 1971. The genus Lepidoderma. Mycologia 63: 490-516. Kowalski, D. T. 1972. Two new alpine Myxomycetes from Washington. Mycologia 64: 359-364. Kowalski, D. T. and Hinchee, A. A. 1972. Barbeyella minutissima – a common alpine myxomycete. Syesis 5: 9597. Leontyev, D. V. 2010. Plant community preferences of some Myxomycete species in Gomolsha forests. Nauka Studia (Poland). 4(28): 14-24. Liu, C. H., Chen, Y. F., Chang, J. H., and Yang, F. H. 2002. Myxomycetes of Taiwan XVI. One new species and one new record of Physaraceae. Taiwania-Taipei 47(4): 290-297. Lloyd, Sarah. 2018. Tasmanian Myxomycetes. Cribraria species. Accessed 9 May 2019 at . Longton, E. E. 1980. Physiological ecology of mosses. In: Taylor, R. J. and Leviton A. E. (eds.). The Mosses of North America. Pacific Division of the Amer. Assoc. Adv. Sci., San Francisco, California, pp. 77-103. Martin, G. W. and Alexopoulos, C. J. 1969. The Myxomycetes. Univ. Iowa Press, Iowa City, 560 pp. Neubert, H., Nowotny, W., and Baumann, K. 1993. Die Myxomyceten Deutschlands und des angren-zenden Alpenraums unter besonderer Berücksichtingun Österreichs. Vol. 1. Karlheinz Baumann Verlag, Gomaringen. Nissan, B. 1997. Myxomycetes from Israel IV. Mycoscience 38: 87-89. Novozhilov Y. K. 2005. Myxomycetes (class Myxomycetes) of Russia: Taxonomic composition, ecology and geography. Dr of Biol. thesis, 48 pp. (in Russian). Poulain, M., Meyer, M., and Bozonnet, J. 2011. Les Myxomycètes 1. Guide de détermination. Fédération mycologique et botanique Dauphiné-Savoie, Sévrier, 568 pp. Ranade, V. D., Korade, S. T., Jagtap, A. V., and Ranadive, K. R. 2012. Checklist of Myxomycetes from India. Mycosphere 3: 358-390. Robbrecht, E. 1974. The Genus Arcyria Wiggers (Myxomycetes) in Belgium. Bull. Jardin Bot. Natl. Belgique / Bull. Natl. Plant. België 44: 303-353. Rojas, C. and Stephenson, S. L. 2007. Distribution and ecology of myxomycetes in the high-elevation oak forests of Cerro Bellavista, Costa Rica. Mycologia 99: 534-543.

Rollins, A. W. and Stephenson, S. L. 2011. Global distribution and ecology of Myxomycetes. Plant Biol. 12: 1-14. Rosing, W. C., Mitchell, D. W., and Stephenson, S. L. 2007. Corticolous Myxomycetes from Victoria. Australasian Mycol. 26: 9-15. Schnittler, M. 2001. Foliicolous liverworts as a microhabitat for Neotropical Myxomycetes. Nova Hedw. 72: 259-270. Schnittler, M. and Novozhilov, Y. 1996. The Myxomycetes of boreal woodlands in Russian northern Karelia: A preliminary report. Karstenia 36: 19-40. Schnittler, M. and Novozhilov, Y. K. 1998. Late-autumn Myxomycetes of the Northern Ammergauer Alps. Nova Hedw. 66: 205-222. Schnittler, M. and Stephenson, S. L. 2000. Myxomycete biodiversity in four different forest types of Costa Rica. Mycologia 92: 626-637. Schnittler, M., Stephenson, S. L., and Novozhilov, Y. K. 2000. Ecology and world distribution of Barbeyella minutissima (Myxomycetes). Mycolog. Res. 104: 15181523. Schnittler, M., Lado, C., and Stephenson, S. L. 2002. Rapid biodiversity assessment of a tropical myxomycete assemblage - Maquipucuna Cloud Forest Reserve, Ecuador. Fung. Divers. 9: 135-167. Schnittler, M., Unterseher, M., and Tesmer, J. 2006. Species richness and ecological characterization of Myxomycetes and myxomycete-like organisms in the canopy of a temperate deciduous forest. Mycologia 98: 223-232. Schuster, R. M. 1957. Boreal Hepaticae, a manual of the liverworts of Minnesota and Adjacent regions. II. Ecology. Amer. Midl. Nat. 57: 203-299. Singer, H., Moreno, G., Illana, C., and Lizarraga, M. 2005. Mountainous and nivicolous Myxomycetes described by Charles Meylan. A SEM-study. Österr. Z. Pilzk 14: 11-29. Smith, T. and Stephenson, S. L. 2007. Algae associated with Myxomycetes and leafy liverworts on decaying spruce logs. Castanea 72: 50-57. Snell, K. L. and Keller, H. W. 2003. Vertical distribution and assemblages of corticolous Myxomycetes on five tree species in the Great Smoky Mountains National Park. Mycologia 95: 565-576. Stephenson, S. L. 1985. New records of Myxomycetes from West Virginia. III. Castanea 50: 262-264. Stephenson, S. L. and Studlar, S. M. 1985. Myxomycetes fruiting upon bryophytes: Coincidence or preference? J. Bryol. 13: 537-548. Stephenson, S. L., Kalyanasundaram, I., and Lakhanpal, T. N. 1993. A Comparative biogeographical study of Myxomycetes in the mid-Appalachians of Eastern North America and two regions of India. J. Biogeogr. 20: 645-657. Stephenson, S. L., Novozhilov, Y. K., and Schnittler, M. 2000. Distribution and ecology of Myxomycetes in high-latitude regions of the Northern Hemisphere. J. Biogeogr. 27: 741754. Stojanowska, W. and Panek, E. 2004. Myxomycetes of the nature reserve near Wałbrzych (SW Poland). II. Dependence on the substrate and seasonality. Acta Mycol. 39: 147-159. Studlar, S. M. 1982. Host specificity of epiphytic bryophytes near Mountain Lake, Virginia. Bryologist 85: 37-50. Ukkala, T., Härkönen, M., and Zeng, Z. 2001. Myxomycetes of Hunan, China. I. Ann. Bot. Fennici 38: 305-328. Wolf, J. H. D. 1993. Diversity patterns and biomass of epiphytic bryophytes and lichens along an altitudinal gradient in the northern Andes. Ann. Missouri Bot. Gard. 80: 928-960.

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Yatsiuk, I. I., Leontyev, D. V., and Shlakhter, M. L. 2018. Myxomycetes of the National Nature Park Slobozhanskiy (Ukraine): Biodiversity and noteworthy species. Nord. J. Bot. 36(1-2), njb-01605. Zabka, G. G. and Lazo, W. R. 1962. Reciprocal transfer of materials between algal cells and myxomycete plasmodia in intimate association. Amer. J. Bot. 49: 146-148.

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Glime, J. M. 2019. Slime Molds: Ecology and Habitats – Lesser Habitats. Chapt. 3-4. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. Ebook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 18 July 2020 and available at .

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CHAPTER 3-4 SLIME MOLDS: ECOLOGY AND HABITATS – LESSER HABITATS TABLE OF CONTENTS Epiphyllous Leafy Liverwort Associations ......................................................................................................... 3-4-2 Non-Epiphyllous Liverwort Associations ........................................................................................................... 3-4-5 Leaf Litter ........................................................................................................................................................... 3-4-7 Soil Associations ................................................................................................................................................. 3-4-8 Rock Assoiations............................................................................................................................................... 3-4-15 Sand Dunes ....................................................................................................................................................... 3-4-16 Alpine and Polar ................................................................................................................................................ 3-4-16 Wet-Habitat Associations.................................................................................................................................. 3-4-22 Ravines....................................................................................................................................................... 3-4-22 Wet Rocks .................................................................................................................................................. 3-4-27 Sphagnum Dwellers ................................................................................................................................... 3-4-28 Summary ........................................................................................................................................................... 3-4-31 Acknowledgments ............................................................................................................................................. 3-4-31 Literature Cited ................................................................................................................................................. 3-4-31

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CHAPTER 3-4 SLIME MOLDS: ECOLOGY AND HABITATS – LESSER HABITATS

Figure 1. Lophocolea heterophylla with slime molds. Photo by Sture Hermansson, with online permission.

Epiphyllous Leafy Liverwort Associations

habitat appears to be less than ideal, as evidenced by the atypically small sporocarps.

In the tropics, epiphyllous (growing on leaves) liverworts (Figure 2) are common, typically associated with lichens, fungi, algae, and bacteria. Mosses are rare in this association. But some associations also include slime molds. Schnittler (2001) found eleven species of slime molds associated with epiphyllous liverworts (Figure 2) in Ecuador, Costa Rica, and Puerto Rico. He found 11 species, with 97% of the 131 cultures producing growths of slime molds. One of his finds, Arcyria afroalpina (Figure 3-Figure 4), was a new find for the Neotropics (Schnittler et al. 2002). When samples of 15 leaf pieces were cultured in moist chambers, the most frequent slime mold species (59-66%) were Arcyria cinerea (Figure 5), Didymium iridis (Figure 6), and D. squamulosum (Figure 7). These most likely occur with the epiphylls as myxamoebae. Lowland rainforests that have a high annual rainfall provide the greatest numbers of slime molds. However, the

Figure 2. Leptolejeunea epiphylla on leaf. Photo by Tom Thekathyil, with permission.

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7). At least the three most common species of slime molds (Arcyria cinerea, Didymium iridis, and D. squamulosum) are very probably regular inhabitants of liverwort-covered leaves. Several lines of evidence seem to support this. First, all three species were found with very scattered and often solitary sporocarps considerably smaller than typical for fructifications of these species in other microhabitats. In addition, tiny phaneroplasmodia (conspicuous plasmodia, as in the Physarales; Figure 8), 1-3 mm in extent were frequently observed in the first two weeks of culture. Plasmodia migrating from the litter layer to fruit on living plants are much larger.

Figure 3. Arcyria afroalpina spores and capillitia. Photo by Yuri Novozhilov, Eumycetozoan Project, DiscoverLife.org, with online permission.

Figure 6. Didymium iridis sporangia, one of the most frequent epiphyllous species of slime molds cultured from leaves with epiphyllous liverworts. Photo through Creative Commons.

Figure 4. Arcyria afroalpina spore, SEM. Photo by Yuri Novozhilov, The Eumycetozoan Project, DiscoverLife.org, with online permission.

Figure 7. Didymium squamulosum. Photo by John Shadwick, The Eumycetozoan Project, DiscoverLife.org, with online permission.

Figure 5. Arcyria cinerea, one of the most frequent epiphyllous species of slime molds cultured from leaves with epiphyllous liverworts. Photo by Kim Fleming, through Creative Commons.

On the other hand, all six sites clearly share an assemblage of common species (Fig. 2) (Schnittler 2001). The average frequency of the three most common species on epiphyllous liverwort covers was surprisingly high, with 0.59 for Arcyria cinerea (Figure 5) and 0.66 for both Didymium iridis (Figure 6) and D. squamulosum (Figure

Figure 8. Phaneroplasmodium. Photo by Sarah Lloyd, with permission.

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There is a potential for direct leaf-to-leaf dispersal of myxamoebae as well as their dormant stages (microcysts) by rainwater or leaf-dwelling insects (Schnittler 2001). Occasional cultures produce growths of Diderma effusum (Figure 9), D. hemisphaericum (Figure 10), Lamproderma scintillans (Figure 11), and Physarum compressum (Figure 12); all other recorded slime molds are rare. None of the slime molds found in this study seems to be specialized for living leaves as a microhabitat. The leaf microflora most likely supplies ample food for successful colonization. However, some differ sufficiently from nonepiphyllous populations that they might be separate races.

Figure 11. Lamproderma scintillans sporangia, a slime mold that occasionally occurs with epiphyllous liverworts. Photo by Ray Simons, The Eumycetozoa Project, DiscoverLive.com, with online permission.

Figure 12. Physarum compressum, a slime mold that occasionally occurs with epiphyllous liverworts. Photo by David Mitchell, The Eumycetozoan Project, DiscoverLife.org, with online permission. Figure 9. Diderma effusum on moss, a slime mold that occasionally occurs with epiphyllous liverworts. Photo by Ray Simons, The Eumycetozoan Project, DiscoverLife.org, with online permission.

Camino et al. (2008) reported on the slime molds in the mountains of central Cuba. There they found two species associated with epiphyllous liverworts: Arcyria afroalpina (Figure 4-Figure 3) and Comatricha laxa (Figure 13).

Figure 10. Diderma hemisphaericum, a slime mold that occasionally occurs with epiphyllous liverworts. Photo by Clive Shirley, The Hidden Forest, with permission.

Figure 13. Comatricha laxa sporangia on decaying log, a species known to also associate with epiphyllous leafy liverworts. Photo by Clive Shirley, The Hidden Forest, with permission.

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Non-Epiphyllous Liverwort Associations Stephenson and Studlar (1985) reported Arcyria cinerea (Figure 5), Physarum viride (Figure 14), Stemonitis axifera (Figure 15-Figure 16), Trichia decipiens (Figure 17), and T. favoginea (Figure 18) associated with non-epiphyllous leafy liverworts, but they were not restricted to this substrate. As already noted, Barbeyella minutissima (Figure 19) and Lepidoderma tigrinum (Figure 20) exhibited a preference for leafy liverworts on rotten conifer logs. In fact, the rare B. minutissima is mostly known from the leafy liverworts Nowellia curvifolia (Figure 19, Figure 21), Lepidozia reptans (Figure 22), and Cephalozia lunulifolia (Figure 23-Figure 24).

Figure 16. Stemonitis axifera with liverworts, a species that can be associated with leafy liverworts on logs and elsewhere. Photo by Clive Shirley, Hidden Forest, with permission.

Figure 14. Physarum viride sporangia, a species that can be associated with leafy liverworts on logs and elsewhere. Photo by Sarah Lloyd, with permission. Figure 17. Trichia decipiens sporangia, a species that can be associated with leafy liverworts on logs and elsewhere. Photo by Fungi07, through public domain.

Figure 15. Stemonitis axifera plasmodium starting to produce sporophytes, a species that can be associated with leafy liverworts on logs and elsewhere. Photo by Clive Shirley, The Hidden Forest, with permission.

Figure 18. Trichia favoginea on log with liverworts. Photo by Jerry Cooper, through Creative Commons.

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Figure 19. Barbeyella minutissima sporangia on the leafy liverwort Nowellia curvifolia. Photo by Randy Darrah, The Eumycetozoan Project, DiscoverLife.org, with online permission. Figure 22. The liverwort Lepidozia reptans. Michael Lüth, with permission.

Figure 20. Lepidoderma tigrinum with sporangia on moss, a species that is more common on leafy liverworts. Photo by Alain Michaud, The Eumycetozoan Project, DiscoverLife.org, with online permission.

Photo by

Figure 23. Cephalozia lunulifolia, a suitable substrate for a number of species of slime molds. Photo by Michael Lüth, with permission.

Figure 24. Cephalozia lunulifolia, a suitable substrate for a number of species of slime molds. Photo by Hermann Schachner, through Creative Commons.

Figure 21. Nowellia curvifolia on log, a suitable substrate for a number of species of slime molds. Photo by Bernd Haynold, through Creative Commons.

Nowellia curvifolia (Figure 19, Figure 21) is the most common slime mold associate (Stephenson & Studlar 1985); it is a liverwort found almost exclusively on rotten logs (Schuster 1957). Hence, the preference in the rotting log habitat for leafy liverworts may simply be that leafy

Chapter 3-4: Slime Molds: Ecology and Habitats – Lesser Habitats

liverworts are common on rotting logs. The mosses Tetraphis pellucida (Figure 25) and Dicranum montanum (Figure 26-Figure 27) are also common associates of slime molds, and likewise are characteristic of rotting wood (Stephenson & Studlar 1985). It is likely that the slime molds are opportunists or simply have broad enough habitat requirements to permit their survival on the potentially competing bryophytes.

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Leaf Litter Some moss dwellers are also litter slime molds. Compagno et al. (2016) reported Didymium melanospermum (Figure 28) on mosses or litter. Moreno et al. (2018) found Didymium nigripes (Figure 29) on moss debris in Spain. Doidge (1950) reported Diderma subdictyospermum on moss and dead leaves. Similarly, Ranade et al. (2012) reported Diderma alpinospumarioides on dead leaves and twigs, but sometimes on living moss in India. Renade and coworkers found that Physarum melleum (Figure 30) occurs on dead leaves as well as among living mosses. Sarah Lodge photographed Collaria aff. rubens (Figure 31) on mosses; this is a species that typically is associated on deciduous leaf litter (Takahashi 2015).

Figure 25. Tetraphis pellucida with gemmae, a common rotten wood moss. Photo by Hermann Schachner through Creative Commons.

Figure 28. Didymium melanospermum on leaves of a soil moss (Mniaceae). Photo by Armand Turpel, through Creative Commons.

Figure 26. Dicranum montanum, a suitable substrate for some slime molds, on rotting log. Photo by Janice Glime.

Figure 27. Dicranum montanum showing the curly leaves when dry. Photo by Janice Glime.

Figure 29. Didymium nigripes sporangia, a species known from moss debris. Photo by Christophe Quintin, with online permission.

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Figure 30. Physarum melleum sporangia, a species of dead leaves and living mosses. Photo by Clive Shirley, The Hidden Forest, with permission.

Figure 31. Collaria aff. rubens on mosses, a species associated with leaf litter. Photo by Sarah Lloyd, with permission.

Figure 32. Polytrichum sp. on the forest floor, habitat for Fuligo muscorum and several species of Physarum. Photo by Janice Glime.

Figure 33. Dicranum scoparium on the forest floor, habitat for Fuligo muscorum and several species of Physarum. Photo by Janice Glime.

Soil Associations Soil associations between bryophytes and slime molds seem to be much less common than associations in other habitats. In temperate forests, mosses of Polytrichaceae (Figure 32, Figure 36), Dicranaceae (Figure 33-Figure 34), and Hypnaceae (Figure 35) are common, with the slime molds Fuligo muscorum (Figure 36), Physarum citrinum, P. confertum (Figure 37), and P. virescens (Figure 38Figure 39) occasionally occurring on them (Ing 1994). One very rare slime mold (Elaeomyxa cerifera – Figure 40Figure 41) is known from the soil-dwelling thallose liverwort Pellia epiphylla (Figure 42) (Hadden 1921; Ing 1994) and from decaying wood, usually in association with bryophytes (Steven Stephenson, pers. comm. 1 June 2019).

Figure 34. Dicranum scoparium, habitat for Fuligo muscorum and several species of Physarum. Photo by Janice Glime.

Chapter 3-4: Slime Molds: Ecology and Habitats – Lesser Habitats

Figure 35. Hypnum curvifolium, a species of the forest floor and logs and a common substrate for moss-dwelling slime molds. Photo by Bob Klips, through Creative Commons.

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Figure 38. Physarum virescens in early fruiting stage on moss. Photo by Alexey Sergeev, with permission.

Figure 39. Physarum virescens on the moss Dicranum. Photo by Alexey Sergeev, with permission.

Figure 36. Fuligo muscorum on Polytrichaceae. Photo by James K. Lindsey, with permission.

Figure 37. Physarum confertum, a slime mold species that occurs on forest mosses in the families Polytrichaceae, Dicranaceae, and Hypnaceae. Photo from The Eumycetozoan Project, DiscoverLife.org, with online permission.

Figure 40. Elaeomyxa cerifera with sporangia on mosses. Photo by Sarah Lloyd, with permission.

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Figure 43. Fuligo intermedia on Polytrichum. Photo by David Mitchell, The Eumycetozoan Project, DiscoverLife.org, with online permission.

Figure 41. Elaeomyxa cerifera sporangium beginning to dehisce. Photo from Myxotropic.org, through Creative Commons.

Figure 44. Pogonatum aloides (Polytrichaceae), one of the substrates for the slime mold Fuligo intermedia. Photo by Hermann Schachner, through Creative Commons.

Figure 42. Pellia epiphylla with capsules, substrate for Elaeomyxa cerifera. Photo by Li Zhang, with permission.

Pant and Tewari (1982) described the growth of Fuligo intermedia (Figure 43) on mosses in Nainital in the Himalayan region of India. These slime molds occurred on the mosses Atrichum obtusulum, Pogonatum aloides (Figure 44), Barbula sp. (Figure 45), and Leucodon secundus. Only the green tips of the mosses appeared above the yellowish-white of the Fuligo intermedia (Figure 43). They suspected that the growth of the mosses was retarded. A related species, Fuligo cinerea (Figure 46-Figure 47) occurs on dead leaves, yeast, and rotten cloth pieces, as well as on mosses and lichens.

Figure 45. Barbula convoluta; the genus Barbula is one of the substrates for the slime mold Fuligo intermedia. Photo by Dale A. Zimmerman Herbarium, Western New Mexico University, with permission from Russ Kleinman and Karen Blisard.

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Figure 46. Fuligo cinerea on lichens and leafy liverworts on bark. Photo by Alexey Sergeev, with permission.

Figure 48. Typhula lutescens with sporangia on mosses. Photo by Tomasz Pachlewski, with permission.

Figure 47. Fuligo cinerea on a mossy forest floor. Photo by Ramsés Pérez, through Creative Commons.

It is not unusual to find that species cannot be put into their proper substrate heading when using the descriptions. This is not necessarily the fault of the author. Information is often based on herbarium labels and material present with the specimen, but not seen in the field by the author(s). Physarum citrinum occurs on terrestrial mosses in woodlands, but were the mosses on soil (Ing 1982)? Later, Ing (1994) reported this species from soil. Ing (1982) was able to be more specific in reporting Physarum virescens (Figure 38-Figure 39) as mostly on terrestrial mosses in woodlands and characteristic of sessile oakwoods, a species that elsewhere is also almost always associated with bryophytes (Steven Stephenson, pers. comm, 1 June 2019). In Spain, Physarum bivalve (Figure 49) occurs on mosses (Castillo et al. 2009), but in what habitat?

Figure 49. Physarum bivalve, a species known from mosses in Spain. Photo by Rod Nelson, DiscoverLife.org, with online permission.

Schnittler and Novozhilov (1996) described several slime mold-bryophyte associations that appear to be on soil in their study of the northern Karelia of Russia. One they noted as a very scanty collection of Physarum cf. carneum on mosses. They were more specific in noting Physarum virescens (Figure 38-Figure 39) as preferring big moss tussocks on the ground, especially Dicranum (Figure 103). Stemonitis fusca (Figure 50) was represented by a single collection on moss tussocks in a spruce-birch-aspen woodland. Didymium melanospermum (Figure 28) typically occurs on thick moss tussocks on soil, but it also occurs at the base of rocks, or even more rarely on litter.

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Similarly, Leocarpus fragilis (Figure 51-Figure 52) can grow on the ground, on mosses, and on litter, but it can only be located in autumn.

Ranade et al. (2012) reported several species that are likely to be associated with soil or litter. Cribraria intricata (Figure 53; syn.=C. dictydioides) occurs not only on rotten wood, but also on roots and dead mosses. Cribraria languescens (Figure 54-Figure 55) occurs on rotten stems and mosses, presumably on the ground. They reported that Physarum didermoides (Figure 56; syn.=Diderma spumarioides) occurs on living moss, presumably on soil mosses. Collaria arcyrionema (Figure 57; syn.=Lamproderma arcyrionema) occurs not only on wood, but also on dead leaves and mosses. Lamproderma echinulatum (Figure 58) and Metatrichia floriformis (Figure 59; syn.=Trichia floriformis) likewise occur on mosses, presumably on the forest floor. Physarum brunneolum (Figure 60) occurs not only on mosses, but also on lichens and decaying wood; again, the substrate of the mosses and lichens is not provided. The most unusual substrate is that of Stemonitis flavogenita (Figure 61) on a dead archegoniophore of the thallose liverwort Marchantia sp. (Figure 62), presumably with the latter growing on soil.

Figure 50. Stemonitis fusca with sclerotia and sporangia on mosses. Photo by Deryni, through Creative Commons.

Figure 53. Cribraria intricata sporangia on bark with a few mosses. Photo by Fluff Berger, through Creative Commons.

Figure 51. Leocarpus fragilis on moss. Photo by Matt Goff, Sitka Nature, with permission.

Figure 52. Leocarpus fragilis on a soil moss in the Polytrichaceae. Photo by Boris Loboda, with permission.

Figure 54. Cribraria languescens, a species that occurs on rotten wood, roots, and dead mosses. Photo from Myxotropic, through Creative Commons.

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Figure 58. Lamproderma echinulatum sporangia on bryophytes. Photo by Clive Shirley, The Hidden Forest, with permission.

Figure 55. Cribraria languescens sporangium. Photo from Myxotropic, through Creative Commons.

Figure 56. Physarum didermoides on mosses. Photo by Andrew Khitsun, with online permission.

Figure 57. Collaria arcyrionema, a species that occurs on dead wood and mosses. Photo by Taibif.tw, through Creative Commons.

Figure 59. Metatrichia floriformis with mosses on bank. Photo by David Mitchell, The Eumycetozoan Project, DiscoverLife.org, with online permission.

Figure 60. Physarum brunneolum, a species of mosses, lichens, and decaying wood. Photo from The Eumycetozoan Project, DiscoverLife.org, with online permission.

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Figure 63. Diachea leucopodia on leaf litter, a species that also occurs on mosses. Photo by Rosser1954, through Creative Commons.

Figure 61. Stemonitis flavogenita, a species that has been found on a dead archegoniophore of Marchantia. Photo by Malcolm Storey, DiscoverLife.org, with online permission.

Figure 64. Diderma testaceum on leaf litter, a species that also occurs on mosses. Photo by Alain Michaud, The Eumycetozoan Project, DiscoverLife.org, with online permission.

Figure 62. Marchantia polymorpha archegoniophores, one of the substrates for Stemonitis flavogenita. Photo by Janice Glime.

Joshaghani et al. (2013) reported Badhamia ovispora as occurring on forest mosses in Iran. This suggests that they grew on soil mosses as the other records were more specific in referring to wood or rotten wood. Stojanowska and Panek (2004) were specific about a number of species of slime molds that occurred on mosses on logs or stumps, but they reported some simply from mosses. Presumably, these were forest floor mosses, including Diachea leucopodia (Figure 63), Diderma testaceum (Figure 64), and Physarum virescens (Figure 38-Figure 39) (plasmodial stage). They described Diderma deplanatum (Figure 65) as surrounding mosses. Lamproderma columbinum (Figure 66) occurred on the moss Tetraphis pellucida (a species of rocks and decaying wood; Figure 25), but also on the moss Dicranum scoparium (Figure 33-Figure 34) – a moss that could occur on soil, rocks, logs, or tree bases.

Figure 65. Diderma deplanatum on mosses. Photo by The Eumycetozoan Project, DiscoverLife.org, with online permission.

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rocks, where "it prefers medium-wet places between the pure slimy algae layers and the big moss tussocks."

Figure 66. Lamproderma columbinum, with fruiting bodies of slime mold on bryophytes. Photo from The Eumycetozoan Project, DiscoverLife.org, with online permission.

Rock Associations Among the earliest moss-slime mold associations reported is that of Kaiser (1913). Brown capsules of the slime mold Leocarpus fragilis (Figure 51) occurred on the moss Dicranum fulvum (Figure 67) in the southern Catskill Mountains of New York. The substrate was not reported, but this moss commonly occurs on sandstone rocks (Seltzer & Wistendahl 1971). The slime mold is not bryophilous, being common on dead leaves (Kaiser 1913).

Figure 68. Physarum album sporangia on decaying wood, a generalist that also occurs on mosses. Photo by George Shepherd, through Creative Commons.

On granite rocks Schnittler and Novozhilov (1996) found two subassociations of slime molds. One prefers the thicker tussocks (> 0.5 cm), especially the mosses Sanionia uncinata (Figure 69), Dicranum fuscescens (Figure 70), and Cynodontium strumiferum (Figure 71). These tussocks have dry leaf tips, but the tussocks have a wet interior and are enriched with small particles of detritus. The slime molds Lamproderma columbinum (Figure 66), L. sauteri (Figure 72), and Didymium melanospermum (Figure 28) fruit here, the latter often at the bases of the rocks. The second sub-association occurs in thin water films and will be discussed below under the Wet Habitat Associations.

Figure 67. Dicranum fulvum, sometimes a substrate for the slime mold Leocarpus fragilis. Photo by Bob Klips, with permission.

Schnittler and Novozhilov (1996) reported on a number of slime molds using bryophytes as a substrate in the northern Karelia of Russia. One of the most common species, Physarum album (Figure 68) appears to be a generalist and includes moss tussocks on rocks among its substrata. Physarum viride (Figure 14) likewise accepts a number of substrata, including moss and liverwort layers of

Figure 69. Sanionia uncinata, a species forming thick mats with dry tips but moist interiors and collections of detritus. It serves as substrate for the slime molds Lamproderma columbinum, L. sauteri, and Didymium melanospermum. Photo by Hermann Schachner, through Creative Commons.

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Figure 70. Dicranum fuscescens, a rock-dwelling moss that serves as substrate for the slime molds Lamproderma columbinum, L. sauteri, and Didymium melanospermum. Photo by Michael Lüth, with permission.

Diderma lucidum seems to be restricted to mossy rocks (Brooks et al. 1977). Few studies seem to have included the rock habitat. Schnittler and Novozhilov (1996), studying the boreal woodlands of northern Karelia in Russia, have contributed a number of records of slime molds that seemingly are able to live on rocks by using bryophytes as their immediate substrate. Lamproderma columbinum (Figure 66) occurred almost exclusively on moss-covered rocks, where it was often accompanied by L. sauteri (Figure 72) and Colloderma oculatum (Figure 73), but preferring drier and thicker moss tussocks than the substrate preferred by these two slime molds. Lamproderma columbinum forms large and conspicuous colonies on thick moss beds on rocks (as well as on moss-covered logs). Lepidoderma tigrinum (Figure 20) fruits in autumn after the first frosts and snowfalls, when it is visible in a rock association of very wet, thin liverwort and algae mats. In summer the plasmodia are visible.

Figure 73. Colloderma oculatum on bryophytes. Photo by David Mitchell, The Eumycetozoan Project, DiscoverLife.org, with online permission.

Figure 71. Cynodontium polycarpon with capsules, a rockdwelling moss that serves as substrate for the slime molds Lamproderma columbinum, L. sauteri, and Didymium melanospermum. Photo by Štĕpán Koval, with permission.

Sand Dunes Sand dunes are inhospitable habitats for both bryophytes and slime molds. But where there is a niche, some bryophyte will usually fill it. Hence, the slime mold Physarum didermoides (Figure 56; syn.=Diderma spumarioides) is common in sand dunes and often forms "plaques of sporangia up to a square meter" on carpets of the moss Syntrichia ruralis (Ing 1994).

Alpine and Polar

Figure 72. Lamproderma sauteri sporangia that can occur on moss-covered rocks. Photo by The Eumycetozoan Project, DiscoverLife.org, with online permission.

When investigating the alpine and Arctic/Antarctic areas, researchers have often been surprised at the low diversity of slime molds. They are both less abundant and exhibit fewer species than in other areas, but some rarer species elsewhere can be present more commonly in the Arctic (Stephenson et al. 2000). Although the cold regions do not appear to be friendly toward slime molds, the most bryophyte-exclusive (perhaps leafy liverwort-exclusive) slime mold, Barbeyella minutissima (Figure 19) is a common alpine slime mold (Kowalski & Hinchee 1972). Similarly, Kowalski (1972) found that in the mountains of Washington, USA, Licea

Chapter 3-4: Slime Molds: Ecology and Habitats – Lesser Habitats

hepatica seems to be restricted to leafy liverworts, a species that seems to be unknown from other substrata (Steven Stephenson, pers. comm. 1 June 2019). This may cause us to be hopeful of special bryophyte associations high in the mountains, but beyond these two limited cases, that does not appear to be the case. Elaeomyxa australiensis (Figure 74) is known from an alpine snowbank habitat in Australia (Moreno et al. 2009; Stephenson & Shadwick 2009). There it grows on litter in association with bryophytes, with only 3 collections out of 300 actually occurring on bryophytes (Stephenson & Shadwick 2009). In these Australian alpine areas, Meriderma cribrarioides (reported as Lamproderma atrosporum; Figure 75) also occurs on bryophytes.

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Stephenson et al. (2000) set out to determine what factors limit slime mold distribution in high-latitude and cold-dominated regions in the Northern Hemisphere. They collected 938 specimens and cultured 1453 substrate samples from 12 study areas in Iceland, northern Russia, Alaska, and Greenland. They identified 150 species, with 33 being widely distributed in at least five study areas. With only 41 species having a frequency greater than 1%, most of the species seemed to have only limited distribution or low frequency. Although the Arctic species seem to have a depauperate representation of species known from the temperate region, as already noted, some species that are considered rare in temperate areas are common in the Arctic, supporting the conclusion that the Arctic slime mold communities are different from those in temperate regions. Novozhilov et al. (1999) reported 56 species of slime molds from the Taimyr Peninsula in north-central Siberia. Among these, only two species apparently were found ever associated with bryophytes. Didymium melanospermum (Figure 28) typically occurs on mossy coarse woody debris. Mucilago crustacea (Figure 76) is even less associated, occurring in a moss- and grass-rich, open patch of the forest tundra. It is notable that slime mold species numbers decrease progressively from the northern taiga, northward to the tundra subzone. This study supports the contention that the tundra is represented by an impoverished flora from the northern taiga subzone.

Figure 74. Elaeomyxa cf. australiensis, an alpine snowbank species that grows with litter in association with bryophytes. Photo by Sarah Lloyd, with permission.

Figure 76. Mucilago crustacea, a species that occurs in moss-rich habitats in the forest tundra. Photo by Alexey Sergeev, with permission.

Figure 75. Meriderma cribrarioides sporangium, a species that sometimes occurs on bryophytes in alpine areas of Australia. Photo by Alain Michaud, The Eumycetozoan Project, DiscoverLife.org, with online permission.

Stephenson et al. (1991) expressed their disappointment at the small number of species they were able to find on the soils of the Alaskan tundra. After collecting from nine different study sites, their cultures yielded only Dictyostelium mucoroides (Figure 77; Dictyosteliomycetes) and D. sphaerocephalum (Figure 78). The total number of slime mold colonies per gram of wet soil averaged more than 100 for all samples and was more than 200 at three of the four Arctic tundra sites. These values are similar to those they found for forest soils in two spruce study sites of interior Alaska.

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Figure 77. Dictyostelium mucoroides (Dictyosteliomycetes) plasmodial slug, a tundra species. Photo by Dmitry Leontyev, The Eumycetozoan Project, DiscoverLife.org, with online permission.

68% D. mucoroides when appearing in cultures. Cavender (1980, 1983) concluded that the altitudinal distribution of slime molds is similar to that of latitude. In the Appalachian Mountains, eastern USA, Cavender (1980) found that the dictyostelid slime molds predominate, with 15 species. The greatest Dictyostelium richness occurred at 590 - 820 m. Landolt et al. (1992) found Dictyostelium mucoroides (Figure 77) and D. sphaerocephalum (Figure 78) to be overwhelmingly dominant in the Kantishna Hills of Denali National Park (formerly Mt. McKinley), Alaska, USA, with the number of clones per gram of wet soil ranging 0-1203. Some of these sites were restoration sites; the natural sites had far greater slime mold density. The mean number of clones per gram of wet soil was 259 clones for the 14 study sites, with the seven natural sites having a mean of 430. Dictyostelium mucoroides was the dominant species (5998%) in the natural sites. In the restoration sites, D. sphaerocephalum was dominant (50-100% of all clones) in the six restoration study plots where slime molds were found. But none of the preceding studies reported any Dictyostelium species on bryophytes. Emphasizing the paucity of species in these cold habitats, Kanda and Sato (1982) were unable to find any cellular slime molds in the alpine tundra of Mt. O-Akan, Hokkaido, Japan. Hence, we should not be surprised that most of these polar and alpine studies did not report any slime molds growing on bryophytes. In the Carpathians of Poland, other species emerge as nivicolous species (Ronikier et al. 2008). These include 18 species, of which 10 are reported for the first time in Poland. Diderma niveum (Figure 79), Lepidoderma chailletii (Figure 80), and Lamproderma ovoideum (Figure 81) are very abundant, particularly in the spring in glades and shrub communities. Diderma alpinum (Figure 82) and D. niveum occur on mosses.

Figure 78. Dictyostelium sphaerocephalum fruiting body, sometimes the only slime mold present in the Alaskan tundra. Photo by Andy Swanson, with permission, image provided by Steve Stephenson.

The report from Stephenson et al. (1991) is similar to that of Benson and Mahoney (1977). But the latter authors considered Dictyostelium mucoroides (Figure 77) to be conspecific with D. sphaerocephalum (Figure 78). They found the latter inclusive species to be dominant above 1700 m in Southern California. Cavender conducted a number of studies in Arctic and high altitude locations. He found a new Alaskan tundra species of Dictyostelium, D. septentrionale, along with D. mucoroides (Figure 77), D. sphaerocephalum (Figure 78), and D. giganteum in that tundra habitat (Cavender 1978). He considered D. sphaerocephalum and D. mucoroides to have sufficiently large populations to play a role in tundra ecology. When Cavender (1983) sampled slime molds in the Rocky Mountains, USA, he found that the soil slime molds were 29.5% Dictyostelium sphaerocephalum and

Figure 79. Diderma cf. niveum sporangia on mosses. Photo by Tom Thekathyil, with permission.

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Lepidoderma crustaceum (Figure 83) is among the bryophyte dwellers found on the subAntarctic Macquarie Island in the Antarctic region (Stephenson et al. 2007a). Lamproderma ovoideum (Figure 84) similarly occurs on the leafy liverwort Lepidozia sp. (Figure 22) on Macquarie Island (Stephenson et al. 1992). But most of the species in the Antarctic region are niveal (subject to actions of snow and ice) species, and their fruiting is associated with winter snow packs. Lamproderma ovoideum is typical of such habitats in alpine areas. Whereas only 6 slime mold species were known in 1990 from the Antarctic region, 32 were known from Iceland and 54 from Greenland (Gøtzsche 1989, 1990). In an intensive study, Stephenson et al. (2007b) located 22 species on Macquarie Island. Figure 80. Lepidoderma chailletii sporangia. Photo by Alain Michaud, The Eumycetozoan Project, DiscoverLife.org, with online permission.

Figure 81. Lamproderma ovoideum sporangia. Photo by Alain Michaud, The Eumycetozoan Project, DiscoverLife.org, with online permission.

Figure 82. Diderma alpinum sporangia, a species that occurs on mosses in the Carpathian Mountains. Photo by The Eumycetozoan Project, DiscoverLife.org, with online permission.

Stephenson et al. (1992) noted the paucity of reports of slime molds from Antarctica and the subAntarctic islands. Several genera occurring there are known from bryophytes elsewhere, but many of the Antarctic species are different. Diderma effusum (Figure 9) is known from mosses in the Antarctic (unpublished record from Steven Stephenson, pers. comm. 1 June 2019).

Figure 83. Lepidoderma crustaceum sporangia, one of the bryophyte dwellers on Macquarie Island. Photo from Myxotropic, through Creative Commons.

Figure 84. Lamproderma ovoideum sporangia, a late snowmelt species in alpine areas, sometimes occurring on bryophytes. Photo by Alain Michaud, The Eumycetozoan Project, DiscoverLife.org, with online permission.

Stephenson et al. (2007b) reported a more diverse slime mold fauna on Macquarie Island, including several that occurred on bryophytes. These bryophyte dwellers included 6 of 80 collections of Trichia verrucosa (Figure 85), 1 of 78 of Diderma alpinum (Figure 86-Figure 87), 2 of 59 of Craterium leucocephalum (Figure 88), 2 of 48 Didymium cf. dubium (Figure 89-Figure 90), 7 of 15 Lamproderma arcyrioides (Figure 91-Figure 92), and 13 of 68 of all other species. Diderma radiatum (Figure 93-

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Figure 94) had a higher ratio, but poor representation, with 1 of the 3 collections being on bryophytes. Lamproderma ovoideum (Figure 84) is considered nivicolous (associated with snow), but the only collection of this species was on bryophytes. Lepidoderma crustaceum (Figure 84) also was reported from bryophytes. The most common bryophytes serving as slime mold substrates on Macquarie Island are the mosses Brachythecium salebrosum (Figure 95), Achrophyllum dentatum (Figure 96-Figure 97), and the leafy liverwort Lophocolea bidentata (Figure 98).

Figure 87. Diderma alpinum spores and capillitium. Photo from The Eumycetozoan Project, DiscoverLife.org, with online permission.

Figure 85. Trichia verrucosa mature and dispersing sporangia, a Macquarie Island slime mold that occasionally fruits on bryophytes. Photo by Clive Shirley, The Hidden Forest, with permission.

Figure 88. Craterium leucocephalum, a slime mold that occasionally appears on bryophytes on Macquarie Island in the Antarctic. Photo by Clive Shirley, the Hidden Forest, with permission.

Figure 86. Diderma alpinum sporangia, a Macquarie Island slime mold that occasionally fruits on bryophytes. Photo from The Eumycetozoan Project, DiscoverLife.org, with online permission.

Figure 89. Didymium dubium on leaf litter, a species that can also occur on bryophytes on Macquarie Island. Photo from The Eumycetozoan Project, DiscoverLife.org, with online permission.

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Figure 90. Didymium dubium spore SEM. Photo from The Eumycetozoan Project, DiscoverLife.org, with online permission.

Figure 93. Diderma radiatum sporangia with mosses on decaying wood, a slime mold that occasionally appears on bryophytes on Macquarie Island in the Antarctic. Photo by Clive Shirley, The Hidden Forest, with permission.

Figure 91. Lamproderma arcyrioides sporangia with moss, sometimes a bryophyte inhabitant on Macquarie Island. Photo by James K. Lindsey, with permission.

Figure 94. Diderma radiatum after the capsules dehisce. Photo by Clive Shirley, The Hidden Forest, with permission.

Figure 92. Lamproderma arcyrioides mature sporangia. Photo by Randy Darrah, The Eumycetozoan Project, DiscoverLife.org, with online permission.

Figure 95. Brachythecium salebrosum, one of the preferred bryophyte substrates for slime molds on Macquarie Island. Photo by Michael Lüth, with permission.

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Wet-Habitat Associations Lindley et al. (2007) remarked on the paucity of information on slime molds in aquatic habitats. They found that the distributions of slime molds above and below the water level were different.

Figure 97. Achrophyllum dentatum with leaf gemmae. Photo by Des Callaghan, through Creative Commons.

Ravines Krziemiewska (1934) reported Colloderma oculatum (Figure 73; as C. dubium) from wet wood covered with mosses and liverworts in her study in the Zaroœlak forest, eastern Carpathians. But studies that concentrate on ravine slime molds are still very limited. One reason for the lack of study in this interesting habitat is that they can only be identified during their fruiting season. In most habitats, bark and other substrate samples can be taken to the lab and cultured. But Novozhilov et al. (2000) lamented the difficulty of culturing the slime molds that prefer the trickling water of humid ravines. This lack of success forces researchers to be in the field when the slime molds are producing sporangia, noting that this is predominately in the late autumn, a time when most slime mold specialists, who are also academicians, are busy with their educational responsibilities. With all this difficulty in being at the right place at the right time, Novozhilov and coworkers estimate that less than 5% of the species occur in such habitats. Whereas most of the slime molds seem to prefer rotting logs, some prefer more moist or even wet habitats. One reason for this may be the associated algae that can serve as a food source. Ing (1994) noticed that algae were typically abundant in association with the mats of bryophytes that served as substrate for slime mold fruiting bodies in cool, moist ravines of the western British Isles (Ing 1983). In another European study, Schnittler and Novozhilov (1998) reported the slime molds Colloderma oculatum (Figure 73) fruiting on wet, moss-covered rock surfaces that presented a continuous layer of algae. Craterium muscorum (Figure 99; syn.=Badhamia rubiginosa var. globosa) and Diderma lucidum are rare Atlantic species that can be found on moss-covered rocks in wooded ravines (Ing 1982). Lamproderma columbinum (Figure 66) and Lepidoderma tigrinum (Figure 20), both species noted elsewhere from bryophytes, are characteristic of ravines. Fuligo muscorum (Figure 100) occurs in wet, terrestrial mossy habitats.

Figure 98. Lophocolea bidentata, one of the preferred bryophyte substrates for slime molds on Macquarie Island. Photo by Hermann Schachner, through Creative Commons.

Figure 99. Craterium muscorum sporangia on mosses, a species that occurs in wet, terrestrial mossy habitats. Photo by Janet Graham, through Creative Commons.

Figure 96. Achrophyllum dentatum, one of the preferred bryophyte substrates for slime molds on Macquarie Island. Photo by David Tng, through Creative Commons.

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reptans (Figure 22), Plagiochila asplenioides (Figure 110), P. spinulosa (Figure 111), Saccogyna viticulosa (Figure 112), and Scapania gracilis (Figure 113). The most common slime molds that occur on these ravine bryophytes are Craterium muscorum (Figure 99), Diderma lucidum, D. ochraceum (Figure 114), Lamproderma columbinum (Figure 66), and Lepidoderma tigrinum (Figure 20).

Figure 100. Fuligo muscorum on the moss Hypnum. Photo by Charles Hipkin, with permission from Barry Stewart.

Lamproderma sauteri (Figure 72) occurs on bryophyte layers on rocks and boulders where there is running water (Novozhilov et al. 2000). These occurrences seem to be mostly in association with the Arctic-alpine leafy liverwort, Gymnomitrion concinnatum (Figure 101). Colloderma oculatum (Figure 73) and Lepidoderma tigrinum (Figure 20) seem to benefit from living on thin, slimy layers of liverworts under a thick cover of mosses and having a covering of water film.

Figure 101. Gymnomitrion concinnatum, an Arctic-alpine leafy liverwort that serves as substrate for Lamproderma sauteri. Photo by Michael Lüth, with permission.

In his 1983 study of ravines in the UK, Ing found that slime molds were associated with the moist bryophytes near waterfalls and dripping areas that kept the mosses moist. Novozhilov et al. (2000) reported a similar relationship on wood and rocks near trickling water in humid ravines. In fact, Lamproderma columbinum (Figure 66; Stemonitidaceae) is an ecotype that is associated with mosses in such habitats. Ing (1983) found that sporangia of slime molds occur most commonly on the mosses Cratoneuron commutatum (Figure 102), Dicranum majus (Figure 103), D. scoparium (Figure 33Figure 34), Hyocomium armoricum (Figure 104), Hypnum cupressiforme (Figure 105), Isothecium myosuroides (Figure 106), Plagiothecium undulatum (Figure 107), and Rhytidiadelphus loreus (Figure 108), and the liverworts Bazzania trilobata (Figure 109), Lepidozia

Figure 102. Cratoneuron commutatum, one of the more common mosses serving as substrate for fruiting slime molds. Photo by Michael Lüth, with permission.

Figure 103. Dicranum majus, a large Dicranum where slime molds commonly form sporangia. Photo by Michael Lüth, with permission.

Figure 104. Hyocomium armoricum, one of the more common mosses serving as substrate for fruiting slime molds. Photo by Michael Lüth, with permission.

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Figure 105. Hypnum cupressiforme, one of the more common mosses serving as substrate for fruiting slime molds. Photo by Michael Lüth, with permission.

Figure 106. Isothecium myosuroides, one of the more common mosses serving as substrate for fruiting slime molds. Photo by Michael Lüth, with permission.

Figure 107. Plagiothecium undulatum, one of the more common mosses serving as substrate for fruiting slime molds. Photo by Michael Lüth, with permission.

Figure 108. Rhytidiadelphus loreus, one of the more common mosses serving as substrate for fruiting slime molds. Photo by Michael Lüth, with permission.

Figure 109. The leafy liverwort Bazzania trilobata, one of the more common liverworts serving as substrate for fruiting slime molds. Photo by Michael Lüth, with permission.

Figure 110. Plagiochila asplenioides, one of the more common liverworts serving as substrate for fruiting slime molds. Photo by Michael Lüth, with permission.

Chapter 3-4: Slime Molds: Ecology and Habitats – Lesser Habitats

Figure 111. Plagiochila spinulosa, one of the more common liverworts serving as substrate for fruiting slime molds. Photo by Michael Lüth, with permission.

Figure 112. Saccogyna viticulosa, one of the more common liverworts serving as substrate for fruiting slime molds. Photo by Michael Lüth, with permission.

Figure 113. Scapania gracilis, one of the more common liverworts serving as substrate for fruiting slime molds. Photo by Michael Lüth, with permission.

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Figure 114. Diderma ochraceum sporangia on moss, a common slime mold on ravine bryophytes. Photo by Alain Michaud, The Eumycetozoan Project, DiscoverLife.org, with online permission.

A very detailed study of slime molds in ravines and their associated bryophytes, using 127 small-scale relevés, is that of Schnittler et al. (2010) in sandstone gorges of Switzerland. They followed the methods developed by Holz (1997) for ravine bryophyte communities. Only five taxa account for 87% of the records, and all of these except Lamproderma puncticulatum (Figure 115-Figure 116) are reported elsewhere in this chapter from bryophyte associations: Colloderma robustum (Figure 117), Diderma ochraceum (Figure 114), Lamproderma columbinum (Figure 66), L. puncticulatum agg., and Lepidoderma tigrinum (Figure 20). They determined that the community is relatively unique, occurring only in the deep, narrow ravines on nearly vertical rocks, mostly on northern exposures. The substrate has a very acidic pH with a mean of 3.35. The fruiting season, in the beginning of October, has a very constant microclimate with nearly 100% relative humidity and ~10ºC. Green algae, most commonly Coccomyxa confluens (Figure 118), were associated with all the slime mold collections. The mosses Dicranodontium denudatum (Figure 119) (59%) and Tetraphis pellucida (Figure 25) (50%) and leafy liverworts Mylia taylorii (Figure 120) (64%) and Diplophyllum albicans (Figure 121) (40%) had high indicator values for the community. Nevertheless, the five most common slime molds had high niche overlap values, but low niche width values, indicating their high degree of specialization. I have to wonder if these slime molds were cryptospecies because they are relatively well known outside ravines and are among species more frequently cited as associated with bryophytes. For example, Hoffmann (1795) originally described Diderma ochraceum from mosses. On the other hand, sufficient habitat information is often lacking.

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Figure 115. Lamproderma puncticulatum immature sporangia on bryophytes. Photo by Mireille Lenne, courtesy of Marianne Meyer.

Figure 118. Coccomyxa confluens on mosses. Photo by James K. Lindsey, with permission.

Figure 119. Dicranodontium denudatum, a common substrate for slime molds in ravines. Photo by David T. Holyoak, with permission.

Figure 116. Lamproderma puncticulatum on the liverwort Pellia. Photo courtesy of Isabelle Mazaud.

Figure 117. Colloderma robustum, a species associated with ravine bryophytes. Photo by Sarah Lloyd, with permission.

Figure 120. Mylia taylorii, a common ravine substrate for slime molds. Photo by Hermann Schachner, through Creative Commons.

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pellucida]. Lepidoderma tigrinum (Figure 20) prefers Dicranodontium denudatum (Figure 119) (74% of all records) and Mylia taylorii (65%), but occurred several times on Sphagnum (Figure 128-Figure 129) tufts at the base of large rocks; Diderma umbilicatum (Figure 122) had a similar preference for these two species. Physarum album (Figure 123) was less common, with only three records on Tetraphis pellucida and one on Dicranodontium denudatum. Overall, the slime molds seem to prefer the closed turfs of Mylia taylorii and Dicranodontium denudatum, but not the common pure short turfs of Tetraphis pellucida.

Figure 121. Diplophyllum albicans, a common ravine substrate for slime molds. Photo by David T. Holyoak, with permission.

Ing (1983) described a ravine slime mold community having a preference for bryophytes on rocks in numerous Atlantic locations in the British Isles. But the species differed somewhat from those in Switzerland: Craterium muscorum (Figure 99), Diderma lucidum, *D. ochraceum (Figure 114), *Lamproderma columbinum (Figure 66), and *Lepidoderma tigrinum (Figure 20), with *species being common in ravines of both countries. Later he (Ing 1994) recognized the ravine slime mold community as a distinct community. Schnittler et al. (2010) did note that even when the inclination was suitable, pure turfs of Tetraphis pellucida (Figure 25) rarely had slime molds, but also tended to have less trickling water or algae. The leafy liverwort Mylia taylorii (Figure 120), on the other hand, is a good indicator organism for the presence of ravine slime molds. These researchers concluded that most of the ravine species are rare outside the ravines, citing Colloderma robustum (Figure 117) and Diderma ochraceum (Figure 114), two species closely associated with Mylia taylorii. Lamproderma puncticulatum (Figure 115-Figure 116) agg. was likewise closely associated with M. taylorii. Other common ravine species, specifically Lamproderma columbinum (Figure 66) and Lepidoderma tigrinum (Figure 20), occur elsewhere in forests with constantly humid conditions; in the British ravines they are closely associated with Tetraphis pellucida (Figure 25). As noted earlier in this chapter, they may be true bryophiles. Diderma umbilicatum (Figure 122) was always "in close neighborhood" with Mylia taylorii and Dicranodontium denudatum (Figure 119), suggesting that this slime mold preferred similar conditions to these two bryophytes. The moving plasmodia of D. umbilicatum were a conspicuous bright yellow. These segregate to form distinct sporangia on the tips of the bryophyte shoots, often forming a doughnut shape around the narrow leaves of Dicranodontium. Other species preferring Tetraphis pellucida (Figure 25) in ravines include Diderma lucidum and Lamproderma columbinum (Figure 66), the latter occurring there in 73% of the Tetraphis turf records where green algae were present in Saxonian Switzerland (Schnittler et al. 2010). Lamproderma puncticulatum (Figure 115-Figure 116) prefers thicker bryophyte tufts [64% with Mylia taylorii (Figure 120), 56% with Tetraphis

Figure 122. Diderma umbilicatum on mosses, a species often near bryophytes in ravines. Photo by Alain Michaud, The Eumycetozoan Project, DiscoverLife.org, with online permission.

Figure 123. Physarum album, a species that occasionally occurs on mosses in ravines. Photo by Sarah Lloyd, with permission.

Schnittler et al. (2010) agreed with Ing (1994) that nitrogen-fixing activity of the Cyanobacteria may be beneficial in some way to the slime molds, possibly as nutrients for their food source, or directly as a food source. But experimental evidence to support this is lacking. They in fact suggested that bryophilous slime molds may instead be phycophilous. Wet Rocks One of the early reports on slime mold-bryophyte associations in wet habitats is that of Lister (1918) in the UK. He found Lamproderma scintillans (Figure 11) on stones in a shallow stream. He surmised that they had migrated to these rocks from mosses and leaf litter on the stream bank.

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Schnittler and Novozhilov (1996) described a granite rock community that is comprised of Colloderma oculatum (Figure 73) and Lepidoderma tigrinum (Figure 20). These two species fruit on very thin (< 0.5 cm), slimy layers of liverworts, covered with a water film. These microhabitat films are found at 1-3 m height on rocks that are provided with trickling water. The large moss tussocks on the upper margins of the rocks can function as a water reservoir. Both slime mold species produce sporangia directly on the water film of the liverworts. The researchers assumed that the plasmodia lived within the bryophyte layers because of their location on the rocks. The huge colonies, especially of Colloderma oculatum, suggest that moss layers are a normal microhabitat. The Cyanobacteria (Figure 124Figure 127) present are a possible food source for the plasmodia. In the northern Ammergauer Alps, Schnittler and Novozhilov (1998) also found Colloderma oculatum on wet rock surfaces where they were associated with mosses and a continuous layer of algae (probably including Cyanobacteria). One such bryophyte dweller that may really be an algae/Cyanobacteria dweller is Physarum viride (Figure 14). This species occurs on two substrate types, one of which is on the moss and liverwort layers of rocks (Schnittler & Novozhilov 1996). It prefers medium-wet places between the pure slimy algae layers and the big moss tussocks. One advantage to living on a wet rock is the presence of Cyanobacteria. Not only do the rocks present slimy layers of these nitrogen-fixing organisms, but so also do the bryophytes (Ing 1994). In the study by Ing, these encrustations are predominantly Nostoc muscorum (Figure 124-Figure 125) or N. commune (Figure 126-Figure 127). For the slime molds, these can be a food source, whereas for the bryophytes, they may improve the nitrogen availability. The beneficial aspects of this association are supported by the frequency with which this assemblage of species coincides with the Nostoc growths. In this case, the rocks are base-rich, and Ing hypothesized that the nitrogenfixing activity of the Nostoc, enhanced by a high pH, may be beneficial for the slime molds. Craterium muscorum (Figure 99), Lamproderma columbinum (Figure 66), and Lepidoderma tigrinum (Figure 20) typically develop plasmodia that have close contact with the Nostoc on these wet rocks.

Figure 125. Nostoc muscorum individual filaments. Photo by Charles Krebs, with online permission.

Figure 126. Nostoc commune on mosses. through Creative Commons.

Yamamaya,

Figure 127. Nostoc commune individual filaments. Photo by David Wagner, with permission.

Figure 124. Nostoc muscorum gelatinous ball, a Cyanobacterium frequently associated with wet bryophytes and of likely benefit to slime molds. Photo from Protist Information Server, with permission.

Sphagnum and peatland Dwellers Sphagnum (Figure 128) offers both a habitat modifier that maintains a high moisture level, and a substrate. Carr (1939) provided an early record of Didymium iridis (Figure 6; as Didymium nigripes var. xanthopus) growing in abundance on Sphagnum.

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Figure 130. Symphytocarpus trechispora on moss. Photo by Thomas Laxton, through Creative Commons.

Figure 128. Sphagnum fallax with capsules. David T. Holyoak, with permission.

Photo by

Schnittler and Novozhilov (1996) noted species of slime molds that were in some way associated with Sphagnum (Figure 128) in the northern Karelia of Russia. Nevertheless, they observed that the Sphagnum-rich spruce (Picea; Figure 129) woodland, despite its nearly continuously moist environment, served as a poor habitat for slime molds. Only Physarum virescens (Figure 38Figure 39) appeared to be adapted sufficiently to live on the large moss tussocks. Figure 131. Polytrichum commune, a common substrate for Symphytocarpus trechispora. Photo by Christopher Tracey through Creative Commons.

In the same study, Ing (1994) found that two bryophiles, Lamproderma columbinum (Figure 66, Figure 132) and Lepidoderma tigrinum (Figure 20), occur on Sphagnum (Figure 128) as well as other bryophytes. Diderma simplex (Figure 133) is a moorland species that includes bog mosses among its substrates. Hagelstein (1941) reported Paradiachea caespitosa (Figure 134) growing on the tips of Sphagnum. But Ing (1994) concludes that in general, the low pH and low oxygen availability make many mires and bogs unsuitable for the growth of slime molds.

Figure 129. Sphagnum in spruce forest. Photo courtesy of Kim Barton.

In his examination of mosses of wet habitats, Ing (1994) found two slime molds that are mostly restricted to growing on Sphagnum (Figure 128). These are Symphytocarpus trechispora (Figure 130) and Amaurochaete trechispora. On the other hand, Salamaga et al. (2014) concluded that in Poland S. trechispora is acidophilic. Whereas it frequently occurs on Sphagnum, it is not restricted to that substrate. They reported it also from Polytrichum sp. (Figure 131) (growing with Sphagnum fallax – Figure 128). It is also known from Sphagnum in Scotland, England, and Germany (Ing 1999; Schnittler et al. 2011).

Figure 132. Lamproderma cf. columbium, on Sphagnum, Catfield Fen. Photo courtesy of Isabelle Masaud.

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Chapter 3-4: Slime Molds: Ecology and Habitats – Lesser Habitats

Figure 135. Didymium ovoideum sporangium on wood, a species that sometimes occurs on Sphagnum. Photo by Thomas Laxton, through Creative Commons. Figure 133. Diderma simplex, a species that can grow on bog mosses. Photo by Bruce Watt, University of Maine, Bugwood.org, through Creative Commons.

Figure 136. Stemonitis axifera sporangia on decorticated log, a species that also occurs on Sphagnum and Polytrichaceae. Photo by Clive Shirley, The Hidden Forest, with permission.

Figure 134. Paradiachea caespitosa, a species that grows at the tip of Sphagnum. Photo by Sarah Lloyd, with permission.

Cavender et al. (2005) reported a new species of cellular slime mold, Dictyostelium quercibrachium (Dictyosteliomycetes), from the margin of a small bog in Ohio, USA. Cavender and Vadell (2006) likewise reported the cellular slime mold Acytostelium magniphorum from the margin of a small bog in Ohio. Landolt et al. (2006) suggested that bog margins provide relict habitats that have been under explored for slime molds and therefore may hold more unknown species or range extensions. In a more recent study in the Ukraine, Yatsiuk et al. (2018) found Didymium ovoideum (Figure 135) on Sphagnum (Figure 128). Didymium melanospermum (Figure 28) and Stemonitis axifera (Figure 136) occurred on species of Sphagnum and Polytrichaceae (Figure 131). Didymium melanospermum typically occurs on acid substrates, including mosses (Stephenson & Studlar 1985; Nannenga-Bremekamp 1991; Ing 1994). On the other hand, Stemonitis axifera does not appear to be bryophilous in most locations.

In Sphagnum (Figure 128) bogs, Badhamia lilacina (Figure 137-Figure 138) seems to prefer aquatic areas, but their fruiting occurs on moss leaves (Tamayama & Keller 2013). Others, like the Leocarpus fragilis (Figure 139) in occur in peatlands but seem to avoid the Sphagnum. Only one tiny patch of this one is on the moss.

Figure 137. Badhamia lilacina plasmodium on Sphagnum. Photo from , with implied permission.

Chapter 3-4: Slime Molds: Ecology and Habitats – Lesser Habitats

Figure 138. Badhamia lilacina on Sphagnum. Photo by Janet Graham, through Creative Commons.

Figure 139. Leocarpus fragilis on Sphagnum and twigs. Photo by Boris Loboda, with permission.

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Ravines provide a unique assemblage of species, and many of these occur on bryophytes, probably in part because bryophytes provide a high cover there. Craterium muscorum, Diderma lucidum, D. ochraceum, Lamproderma columbinum, and Lepidoderma tigrinum are common on bryophytes there. The presence of Mylia taylorii is a good indicator organism for the presence of ravine slime molds, and many also occur on the moss Dicranodontium denudatum. The Cyanobacteria Nostoc muscorum and N. commune are common associates on wet rocks and may provide food for the slime molds. Slime molds occurring in peatlands in association with Sphagnum may be there because of the low pH. Of the 79 genera of slime molds in the Mxyomycetes, Dictyosteliomycetes, and Ceratiomyxomycetes listed by nomen.eumycetozoa.com as of 5 May 2019, 44 have at least one member that has been found on a bryophyte. I have found no records among the protostelids. Summarizing this chapter raises more questions than answers. Do either the bryophytes or the slime molds, or both, benefit from their association? If so, how? Do the bryophytes and slime molds simply prefer the same environmental conditions? It seems likely that moisture is a major factor, but experiments are needed on a sponge or other non-biological material to provide moisture with no nutrients. Do some bryophytes inhibit the growth of slime molds? Do some provide food through the microflora and fauna of the bryophyte, and do others fail to provide it because of growing conditions or inhibitors? Are some slime molds inhibited while others are not by the same bryophyte species? Experiments with bryophyte extracts on cultures of slime molds could be illuminating.

Acknowledgments Summary Habitats for the slime molds are arguably as diverse as those of bryophytes. Some of the "less important" habitats, in terms of number of species, are on epiphyllous leafy liverworts, on liverworts elsewhere, on leaf litter, on soil, on rocks, on sand dunes, in alpine and polar regions, in ravines, on wet rocks, and in peatlands, including on Sphagnum. These habitats contrast with the higher richness and abundance on bark and rotting wood. In all of these habitats, some slime molds exist on bryophytes. Our understanding of this slime mold-bryophyte relationship is almost non-existent. The presence of plasmodia on bryophytes is even less well understood than the presence of sporangia. In contrast to the bryophytes, the species richness and abundance changes of slime molds with increasing elevation mimic those seen for increasing altitude. Alpine areas seem have some of the bryophiles, such as Barbeyella minuta. Polar regions, on the other hand, are often dominated by Dictyosteliomycetes. Records of bryophyte dwellers are rare or non-existent in the polar regions.

Marianne Meyer and Isabelle Charissou were very helpful in providing me with pictures of slime molds on mosses and Marianne helped me with identification of some images contributed by others. Sarah Lloyd and Steve Stephenson were very helpful in providing images, references, and critiques.

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Schnittler, M., Kummer, V., Kuhnt, A., Krieglsteiner, L., Flatau, L., and Müller, H. 2011. Rote Liste und Gesamtartenliste der Schleimpilze (Myxomycetes) Deutschlands. Naturschutz und Biologische Vielfalt. In: Ludwig, G. and MatzkeHajek, G. (eds.). Rote Liste gefährdeter Tiere, Pflanzen und Pilze Deutschlands 6(2), Landwirtschaftsverlag GmbH, Bonn, pp. 125-234. Schuster, R. M. 1957. Boreal Hepaticae, a manual of the liverworts of Minnesota and adjacent regions. II. Ecology. Amer. Midl. Nat. 57: 203-299. Seltzer, R. C. and Wistendahl, W. A. 1971. Some environmental factors related to the occurrence of Dicranum fulvum in Southeastern Ohio. Bryologist 74: 28-32. Stephenson, S. L. and Shadwick, J. D. 2009. Nivicolous myxomycetes from alpine areas of south-eastern Australia. Austral. J. Bot. 57: 116-122. Stephenson, S. L. and Studlar, S. M. 1985. Myxomycetes fruiting upon bryophytes: Coincidence or preference? J. Bryol. 13: 537-548. Stephenson, S. L., Landolt, J. C., and Laursen, G. A. 1991. Cellular slime molds in soils of Alaskan tundra, U.S.A. Arct. Alp. Res. 23: 104-107. Stephenson, S. L., Seppelt, R. D., and Laursen, G. A. 1992. The first record of a Myxomycete from sub Antarctic Macquarie Island. Antarct. Sci. 4: 431-432.

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Stephenson, S. L., Novozhilov, Y. K., and Schnittler, M. 2000. Distribution and ecology of Myxomycetes in high-latitude regions of the Northern Hemisphere. J. Biogeogr. 27: 741754. Stephenson, S. L., Moreno, G., and Singer, H. 2007a. Notes on some nivicolous myxomycetes from Australia and New Zealand including the description of a new species of Lamproderma. Oesterreichische Zeit. Pilz. 16: 11-23. Stephenson, S. L., Laursen, G. A., and Seppelt, R. D. 2007b. Myxomycetes of subAntarctic Macquarie Island. Austral. J. Bot. 55: 439-449. Stojanowska, W. and Panek, E. 2004. Myxomycetes of the nature reserve near Wałbrzych (SW Poland). II. Dependence on the substrate and seasonality. Acta Mycol. 39: 147-159. Takahashi, K. 2015. Distribution of Myxomycetes on varied leaf litter types in a mixed forest in warm-temperate western Japan. Open J. Forest. 5: 686-696. Tamayama, M. and Keller, H. W. 2013. Aquatic Myxomycetes. Fungi 6(3): 18-25. Yatsiuk, I. I., Leontyev, D. V., and Shlakhter, M. L. 2018. Myxomycetes of the National Nature Park Slobozhanskiy (Ukraine): Biodiversity and noteworthy species. Nord. J. Bot. 36(1-2), njb-01605.

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Glime, J. M. 2017. Invertebrates: Introduction. Chapt. 4-1. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. Ebook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 18 July 2020 and available at .

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CHAPTER 4-1 INVERTEBRATES: INTRODUCTION TABLE OF CONTENTS The Invertebrate Fauna........................................................................................................................................ 4-1-2 Sampling ............................................................................................................................................................. 4-1-9 Preservation of Specimens ................................................................................................................................ 4-1-11 Community Patterns .......................................................................................................................................... 4-1-12 Terrestrial/Limnoterrestrial ........................................................................................................................ 4-1-12 Lobules as Habitat...................................................................................................................................... 4-1-14 Aquatic ....................................................................................................................................................... 4-1-14 Altitudinal Gradients .................................................................................................................................. 4-1-16 Food Webs ........................................................................................................................................................ 4-1-16 Pollution ............................................................................................................................................................ 4-1-18 Harvesting Dangers ........................................................................................................................................... 4-1-18 Summary ........................................................................................................................................................... 4-1-19 Acknowledgments ............................................................................................................................................. 4-1-19 Literature Cited ................................................................................................................................................. 4-1-19

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Chapter 4-1: Invertebrates: Introduction

CHAPTER 4-1 INVERTEBRATES: INTRODUCTION

Figure 1. Marchantia polymorpha that has been nibbled by an unknown organism. Note holes in the thallus. Photo by C. R. Stevenson, with permission.

The Invertebrate Fauna Einstein is credited with saying that the most incomprehensible fact about nature is that it is comprehensible (Miller 1992). The invertebrate community associated with bryophytes, especially in terrestrial habitats, needs still to be comprehended. Dendy (1895) coined the term cryptozoic fauna to describe "the assemblage of small terrestrial animals found dwelling in darkness beneath stones, rotten logs, the bark of trees, and in other similar situations." Although not specifically mentioned, bryophytes surely belong among the "other similar situations," as evidenced by the browsed patches on the liverwort in Figure 1. A comparable term for such bryophyte dwellers in the aquatic realm is meiofauna, defined as "benthic (living on the bottom of a body of water) animals that can fit a mesh size of 1 mm and be retained on a mesh size of 42 µm" (Brave New Biosphere 1999). Although living among bryophytes directly contradicts being on the bottom, the bryophytes do occupy the bottom, and one might think of the habitat they create as simply an extension of that bottom.

For many of the invertebrates, the bryophytes represent a moist island among the drier sites. Invertebrates living there because they are able to survive in interstial collections of water droplets are considered limnoterrestrial, and this limnoterrestrial habitat houses many organisms better known in aquatic habitats, such as copepods, gastrotrichs, rotifers, and tardigrades (Thorp & Covich 2010). The invertebrate fauna are likely to play an important role in nutrient cycling within the bryophyte community, thus facilitating return of detrital matter to ecosystem level nutrient cycling. Merrifield and Ingham (1998) suggested that the diversity of feeding strategies found in moss invertebrate communities provides evidence of withinbryophyte-community nutrient cycling. Studies by Davis (1981) seem to support this suggestion. He found that the moss turf community and the moss carpet community in the maritime Antarctic on Signy Island showed similar levels of productivity, trophic structure, and efficiencies of organic matter transfer, but they differed in Collembola

Chapter 4-1: Invertebrates: Introduction

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(springtails) and Acari (mites) standing crops, turnover of mosses, and accumulation of dead organic matter. Both communities [turf of Polytrichum strictum (= P. alpestre; Figure 2-Figure 3) and Chorisodontium aciphyllum (Figure 4-Figure 5) and carpet of Calliergon sarmentosum (Figure 6), Calliergidium austro-stramineum (Figure 7), Sanionia uncinata (Figure 8), and Cephaloziella varians – a liverwort (Figure 9)] had fauna of Protozoa, Rotifera, Tardigrada, Nematoda, Acari, and Collembola. Despite the diverse fauna, Davis found no evidence that the mosses would have been eaten. However, he based this on known feeding groups of the organisms and not on direct evidence. Nevertheless, it is likely that detrital matter and predation were primary food pathways, permitting nutrient cycling.

Figure 4. Chorisodontium aciphyllum in Antarctica, home of Protozoa, Rotifera, Tardigrada, Nematoda, Acari, and Collembola. Photo from Polar Institute, through Creative Commons.

Figure 2. Polytrichum strictum cushions in Alaska, home for Protozoa, Rotifera, Tardigrada, Nematoda, Acari, and Collembola in the Antarctic. Photo courtesy of Andres Baron Lopez.

Figure 3. Polytrichum strictum, home for Protozoa, Rotifera, Tardigrada, Nematoda, Acari, and Collembola in the Antarctic. Photo by Jan-Peter Frahm, with permission.

Figure 5. Chorisodontium aciphyllum, home of Protozoa, Rotifera, Tardigrada, Nematoda, Acari, and Collembola. Photo by Jan-Peter Frahm, with permission.

Figure 6. Calliergon sarmentosum, home for Protozoa, Rotifera, Tardigrada, Nematoda, Acari, and Collembola in the Antarctic. Photo by David T. Holyoak, with permission.

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Chapter 4-1: Invertebrates: Introduction

Nelson and Hauser (2012) examined what would seem to be a very different habitat from that of the Antarctic samples of Davis (1981) – epiphytic mosses and liverworts of the Pacific Northwest, USA. Despite that seeming difference in climate, the same six groups were dominant: Acari, Tardigrada, Collembola, Nematoda, and Rotifera, in that order. Protozoa were also abundant, but they did not quantify those. They found no differences in major groups between mosses and liverworts, but suggested that there may have been differences between species.

Figure 7. Calliergidium austro-stramineum, home for Protozoa, Rotifera, Tardigrada, Nematoda, Acari, and Collembola in the Antarctic. Photo by Bill Malcolm, with permission.

Figure 10. Mean percent and standard deviation of organisms for each of the five dominant taxa groups in epiphytic mosses and liverworts at Tryon Creek State Natural Area, 1, 7, and 17 October 2011, calculated for all samples together. Redrawn from Neslon & Hauser 2012.

Figure 8. Sanionia uncinata, home for Protozoa, Rotifera, Tardigrada, Nematoda, Acari, and Collembola in the Antarctic. Photo by Michael Lüth, with permission.

Figure 9. Cephaloziella varians (among mosses), home for Protozoa, Rotifera, Tardigrada, Nematoda, Acari, and Collembola in the Antarctic. Photo by Kristian Peters, with permission.

In the Czech Republic, Božanić et al. (2013) attempted to illucidate the factors that determined which invertebrates inhabited bryophyte clumps. They examined the fauna on 15 bryophyte species (61 total samples) and identified 45 invertebrate species in 13 higher taxonomic groups. They found that the two most important factors determining the invertebrate fauna were the size of the moss clump (Figure 12) and the height above ground (Figure 13). The moss genus Brachythecium housed the most invertebrate taxa, with the species Brachythecium curtum (Figure 11) on rotten trees housing the most.

Figure 11. Brachythecium curtum on rotten wood, home for the most invertebrate taxa in a Czech Republic study. Photo by Janice Glime.

Chapter 4-1: Invertebrates: Introduction

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Millipedes preferred bryophyte habitats higher above ground, with Nemasoma varicorne (Figure 14) being the most abundant (Božanić et al. 2013). Mites (Acarina), pseudoscorpions (Pseudoscorpiones), and ants (Formicidae) were only in the lower levels. Interestingly, tree diameter also played a role in locations, with the isopods Trichoniscus pusillus (Figure 15) and Porcellium collicola (Figure 16) occupying mosses on smaller trees, whereas the isopod Trachelipus rathkii (Figure 17) and centipedes Lithobius mutabilis and juveniles of other Lithobius species preferred larger trees.

Figure 14. Nemasoma varicorne female, an abundant above ground millipede that can be found among bryophytes. Photo by Walter Pfliegler, with permission.

Figure 12. Relative numbers of invertebrate groups on bryophytes vs moss sample area. Redrawn from Božanić et al. 2013.

Figure 15. Trichoniscus pusillus, a species among mosses on smaller trees. Photo by Andy Murray, through Creative Commons.

Figure 13. Relative numbers of invertebrate groups on bryophytes vs height above ground. Redrawn from Božanić et al. 2013.

Figure 16. Porcellium collicola, a species among mosses on smaller trees. Photo by Dragisa Savic, with permission.

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Chapter 4-1: Invertebrates: Introduction

Figure 17. Trachelipus rathkii, a species among mosses on larger trees. Photo by Joerg Spelda, SNSB, Zoologische Staatssammlung Muenchen, through Creative commons.

Figure 19. Alona, a bryophyte dweller that is most common among them in the drift. Photo by Yuuji Tsukkii, with permission.

Figure 18. Lithobius mutabilis, a species among mosses on larger trees. Photo by Joerg Spelda, SNSB, Zoologische Staatssammlung Muenchen, through Creative Commons.

Dražina et al. (2011) examined the mieofauna of bryophytes in Europe. These included Turbellaria (flatworms), Rotifera (rotifers), Nematoda (nematodes), Gastrotricha, Oligochaeta (segmented worms), Tardigrada (tardigrades), and Crustacea, as well as small, immature insects. They found more than 100 taxa, with rotifers dominating (52 taxa) and nematodes second (27 taxa). In fast water, rotifers averaged an abundance of 219 individuals cm-3. Velocity accounted for much of the variation in locations, with rotifers being most abundant in high velocity and gastrotrichs, tardigrades, and microturbellarians having a negative relationship to flow velocity. Perić et al. (2014) studied the invertebrate drift and found that the meiofauna formed a "considerable" portion of it among moss-rich areas in a karst stream. They found 60 invertebrate taxa in the drift. Only six taxa, all in the annelid and arthropod meiofauna, comprised 35% of the total drift density. Most of the Macroinvertebrates were immature insects. The Cladocera (Alona spp.; Figure 19) comprised 26,7%, Riolus spp. (Coleoptera: Elmidae; Figure 20) comprised 13.2%, Simulium spp. (Diptera: Simuliidae; Figure 21) 12.2%, Enchytraeidae (Annelida; Figure 22) 10.4%, Hydrachnidia (mites; Figure 23) 6.3%, Orthocladiinae (Diptera: Chironomidae; Figure 24) 3.9%, and Naididae (Annelida; Figure 25) 3.6%.

Figure 20. Riolus subviolaceus adult, a genus that is common in mosses and common in stream drift. Photo from Naturalis Biodiversity Center, through Creative Commons.

Figure 21. Simulium larvae, bryophyte dwellers that are common in the drift. Photo from USDA, through Public Domain.

Chapter 4-1: Invertebrates: Introduction

Figure 22. Enchytraeidae, a family with bryophyte dwellers that are common in the drift. Photo by Aina Maerk Aspaas, NTNU University Museum, Department of Natural History, through Creative Commons.

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Figure 25. Naididae, a family with bryophyte dwellers that are common in the drift. Photo by BIO Photography Group, Biodiversity Institute of Ontario, through Creative Commons.

Drozd et al. (2009) conducted studies in bryophyte fauna in the forests of the submountain and mountain areas of the Czech Republic. They concluded that moisture, bryophyte presence, and surprisingly, bryophyte species were the important characteristics determining total abundance. Their study area bryophytes included the mosses Polytrichum commune (Figure 26), Polytrichastrum formosum (Figure 27), Sphagnum teres (Figure 28), Sphagnum girgensohnii (Figure 29, Sphagnum fallax (Figure 30), Pleurozium schreberi (Figure 31-Figure 32), Eurhynchium angustirete (Figure 33), Oligotrichum hercynicum (Figure 34), and the leafy liverwort Bazzania trilobata (Figure 35-Figure 36).

Figure 23. Hydrachnidia, a mite group with bryophyte dwellers that are common in the drift. Photo by Mnolf, through Creative Commons.

Figure 24. Synorthocladius larva, a member of Orthocladiinae; members of this subfamily are common among stream mosses and stream drift. Photo from Stroud Water Research Center, through Creative Commons.

Figure 26. Polytrichum commune habitat, a species of the submountain and mountain areas of the Czech Republic. Photo by Sten Porse, through Creative Commons.

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Chapter 4-1: Invertebrates: Introduction

Figure 27. Polytrichastrum formosum, a species of the submountain and mountain areas of the Czech Republic. Photo by David T. Holyoak, with permission.

Figure 28. Sphagnum teres, a species of the submountain and mountain areas of the Czech Republic. Photo by J. C. Schou, with permission.

Figure 29. Sphagnum girgensohnii, a species of the submountain and mountain areas of the Czech Republic. Photo by Martin Hutten, with permission.

Figure 30. Sphagnum fallax, a species of the submountain and mountain areas of the Czech Republic. Photo from , with permission.

Figure 31. Pleurozium schreberi, a species of the submountain and mountain areas of the Czech Republic. Photo by Bob Klips, with permission.

Figure 32. Pleurozium schreberi, a species of the submountain and mountain areas of the Czech Republic. Photo by Michael Lüth, with permission.

Chapter 4-1: Invertebrates: Introduction

Figure 33. Eurhynchium angustirete, a species of the submountain and mountain areas of the Czech Republic. Photo by Hermann Schachner, through Creative Commons.

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Figure 36. Bazzania trilobata, a species of the submountain and mountain areas of the Czech Republic. Photo by Barry Stewart, with permission.

Sampling

Figure 34. Oligotrichum hercynicum, a species of the submountain and mountain areas of the Czech Republic. Photo by David T. Holyoak, with permission.

Figure 35. Bazzania trilobata, a species of the submountain and mountain areas of the Czech Republic. Photo by Michael Lüth, with permission.

Drozd et al. (2009) lamented the paucity of comprehensive studies, citing many studies that included only one taxonomic group. They studied the bryophyte fauna using 66 traps in three mountain ranges in the Czech Republic. These traps collected more than 55,000 individuals in 5 sites with a mean of 850 individuals per trap. Litter saples had higher arthropod abundance than did moss cushions. They suggested this was probably influenced by the behavior of the detritivorous arthropods that do not have to move about in search of food. They also suggested that the arthropods might use the bryophytes only as a temporary shelter against predators and desiccation. Quantitative field sampling of bryophytes is a challenge, and what works for one species may not work for another. Hynes (1961) collected mosses by hand and stuffed them into a 180 cc jar until it reached capacity, a sample of ca 300 cm2. But this may not work well for some large growths of Fontinalis spp and produces a large sample to be sorted. Furthermore, adding material from other locations in the clump or different clumps diminishes the ability to detect variability and prevents examining subtle effects of stream location. Pulling the moss from the water generally loses few animals because they are adapted to clinging within the moss mat, but pulling the moss apart to make a smaller sample to fit into 180 cc will dislodge even some of the best adapted. Cutting the moss into smaller segments would be less disruptive, but if no bases are samples, some organisms with preferences for bases may be missed. And increasing the sample size of all collections to one suitable for large clumps of Fontinalis (Figure 37) would create a prohibitive sorting size. I found that collecting a handful, preferably to fit into a baby food jar, worked well (Glime1994). The samples were quantified on the basis of moss dry weight after sorting by hand. Frost (1942) used 200 g wet weight for her moss sample size. Since many of the invertebrates disintegrate quickly, 90-95% ethanol should be added immediately. Lower concentrations become too dilute. This method worked well for insects, but may not be suitable for all the non-chitonous invertebrates. These methods will be discussed with the various groups.

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Chapter 4-1: Invertebrates: Introduction

Figure 37. Fontinalis antipyretica, a large aquatic moss that is difficult to sort through. Photo by Bernd Haynold, through Creative Commons.

Hynes (1961) solved the sorting problem by floating the organisms with a saturated solution of calcium chloride. Even with repeated stirring, those organisms with spines and clinging legs may remain in the mosses, as will those nestled at the bases of leaves that curl around them, creating a bias in the sampling. Determining the faunal composition and community structure of these microhabitats is not an easy task. The most obvious method of sampling invertebrates is sorting them from the bryophytes under the dissecting microscope. But this method is tedious, very time-consuming, and often misses the smaller organisms (personal experience!). The method of wringing and squeezing is much less tedious and faster, a method used by Morgan (1977), but certainly many get left behind, and attached organisms are likely to be preferentially left behind, not to mention damage to larger organisms. To help in this time-consuming task, Paul Davison (pers. comm. 21 June 2006) modified the Baermann funnel (Figure 38) for extracting turbellarians (as well as nematodes, copepods, and tardigrades) from bryophytes. A piece of cheese cloth, muslin, or tissue paper is placed in a funnel to hold a sample (Tylka Nematology Lab 2005). This is usually supported by a piece of screening (Figure 38). Then water is run through the sample with rubber tubing clamped at the end of the funnel. After the sample sits overnight or longer, the water is released from the funnel and collected. The first few drops will have a concentration of nematodes, which are heavier than water. Another method is use of the Berlese funnel, which does not have water, using a light and/or temperature gradient that separates mobile organisms such as arthropods and annelids, but that method leaves the nonmobile ones behind, and doesn't work for nematodes (EDSTEEP). If it is too hot, organisms die before they can drop.

Figure 38. Baermann funnel using moss sample. Water can be replaced with air for non-aquatic organisms, thus making it similar to the Berlese funnel. Modified from Briones 2006.

Nelson and Hauser (2012) discovered that the Berlese funnel and soaking in water gave very different results. For the water extraction, they placed the bryophytes in 200 mL water and allowed to settle for at least two hours, following the protocol for tardigrades described by Thorpe and Covich (2010). The sample was taken by sucking up sediment with a dropper and placing two drops on a depression slide. The Berlese funnel method has a strong bias toward arthropods, in this case mites (Acari), whereas the water method found at least 6 types of tardigrades and many algae and protozoa. They found "almost no taxa overlap" between the two extraction methods! Kreutz and Foissner (2006) likewise used liquid extraction. They placed mud on a slide, but for bryophytes it is necessary to wash the bryophytes into water in something like a Petri plate. Detritus and unattached organisms will be dislodged if the bryophytes are stirred into the water. The precipidated detritus can be placed on a slide and separated using the slide-on-slide method described in Chapter 2-6, Protozoa Ecology. Jennings (1979) used the Baerman funnel to extract invertebrates from mosses on Signy Island in the Antarctic. Fairchild et al. (1987) have taken advantage of the behavior of these invertebrates to develop an extraction method. By creating a vertical temperature and oxygen gradient in samples of Sphagnum (Figure 28-Figure 30), they were able to obtain an 85% efficiency. Merrifield and Ingham (1998) compared several methods of extracting invertebrates. In a study of Eurhynchium oreganum (Figure 39) in the Oregon Coast Range, USA, Merrifield and Ingham first verified extraction efficiency for nematodes and other invertebrates using the Baermann funnel. First, invertebrates were collected from the funnel

Chapter 4-1: Invertebrates: Introduction

apparatus, then more were collected from the mosses on subsequent days, and finally more were collected by squeezing and agitation of the moss. More than 90% of cumulative final counts of the nematodes Monhystera spp. (Figure 40) and Prionchulus muscorum (Figure 41) were extracted by the Baermann funnel technique by day 4 of extraction. Tardigrade extraction was even more efficient, reaching 95% by day 4. Rotifers, however, were less efficiently extracted, with only 42% by day 4 and 55% by day 7.

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Andrew and Rodgerson (1999) investigated diversity gradients of invertebrates on bryophytes on two mountains in Tasmania. they compared two extraction techniques for their effectiveness in representing diversity – Tullgren funnels and sugar flotation – with a new technique using kerosene phase separation. When using two samples bulked together, they found that the kerosene phase separation extracted more total individuals and more Acari (mites) and Collembola (springtails). When they compared single samples (1.5 cm x 2.5 cm), the abundance results were the same, but only three of the nine taxa found in the bulked samples were extracted from the single samples. They therefore recommended that two samples be taken and used as replicates (not bulked).

Preservation of Specimens

Figure 39. Eurhynchium oreganum, home for nematodes. Photo by Matt Goff, with permission.

Figure 40. Monhystera sp., a nematode that is extracted effectively from bryophytes by a Baerman funnel. Photo by Peter Mullin, with permission.

Figure 41. Prionchulus muscorum, a nematode that is extracted effectively from bryophytes by a Baerman funnel. Photo by Peter Mullin, with permission.

Ecologists take note. Simply identifying and counting the faunal organisms and getting someone to identify the bryophytes isn't enough! Whereas you may be confident that your expert has identified everything correctly, it is likely that the expert is less confident and has provided you with the "best" determination possible with the material provided. But ecological specimens typically lack reproductive organs, are not well preserved, and may not even be the whole organism. Systematists always pay careful attention to keeping specimens and publishing their location. Ecologists and physiologists should also. Both the bryophytes and the fauna should be preserved and their locations in permanent, reputable herbaria and museums should be part of any publication based on the data. Furthermore, the specimens should be clearly labelled as voucher specimens, referencing the study. Species concepts change; often physiological and ecological properties are not uniform among members of the earlier species concept. In the absence of a specimen, the data become useless. Yet, in 1950, Fosberg examined 270 ecological publications with discussions of species. Locations of preserved specimens were provided in only five of these publications! I decided to see if the situation had improved by using a much smaller sample size of three recent ecological journals and three recent bryological journals. In the 15 papers I examined from ecological journals, there was no mention of preserving or keeping specimens. In the three bryological journals, all 15 papers dealing with systematics or checklists provided the herbaria locations. However, even among this group of biologists who share the same journals, none of the six ecological papers in the same issues mentioned any preservation of specimens from the species included in the study. This practice of providing no preserved reference material defies the concept that scientific data must be verifiable. I disagree with Fosberg (1950) when he pokes fun at stating the source of the nomenclature. Unlike his concept that this is presented to "verify" the identity of the organism, the source of nomenclature demonstrates the species concept used and provides a link to a source where a description may be found. Thus, if one uses Drepanocladus from Crum 1973, we know that a broad concept of the genus is used and that Sanionia, Warnstorfia, or other genus might now apply instead.

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Chapter 4-1: Invertebrates: Introduction

Community Patterns When only aquatic vs terrestrial are considered, we find a difference in groups dominating the bryophytes. In terrestrial habitats, arthropods dominate (Kinchin 1992). Nevertheless, few arthropods spend their entire life cycle among mosses (Kinchin 1990a). The aquatic fauna, Kinchin (1992) contends, is dominated by nematodes, tardigrades, and rotifers. It is not clear if he includes the peatlands in this aquatic grouping, but I have examined the preserved fauna of stream bryophytes, where I have found insects to be the dominant organisms (Glime 1994). I must admit, however, that my bias was to describe the insect communities. A particularly good reference for the identification of species in Sphagnum pools (Figure 42), particularly in Germany, is that of Kreutz and Foissner (2006). However, those on mosses are not distinguished from those in open water.

Figure 43. Bryum argenteum showing its compact habit. Photo by Dick Haaksma, with permission.

Figure 44. Mat of Hypnum cupressiforme. Photo by Dick Haaksma, with permission.

Figure 42. Sphagnum cuspidatum and S. denticulatum with bog pools. Photo by Jonathan Sleath, with permission

Slow drying, as you will soon see, is a prerequisite for survival in many of these faunal organisms. Supporting his argument, Kinchin found that the Bryum argenteum (Figure 43) fauna was much richer than that of Hypnum cupressiforme (Figure 44). Interestingly, he found that mosses such as Tortula muralis (Figure 45) and Grimmia pulvinata (Figure 46) with long hair points have particularly rich fauna, which might again result from a mechanism for slow drying.

Terrestrial/Limnoterrestrial Kinchin (1992) reviewed the invertebrate fauna among bryophytes in the British Isles and provided us with a summary of the "moss" habitat. He found that acrocarpous cushions support a richer fauna than the more loosely packed pleurocarpous mosses, attributing this to the greater ability of acrocarpous cushions to hold water. He demonstrated this ability experimentally, showing that at 100% saturation a cushion of the acrocarpous Bryum argenteum (Figure 43) held 277% of its "dry" weight in water. The pleurocarpous moss Hypnum cupressiforme (Figure 44), on the other hand, held 1496%. Bryum argenteum held 85% of its dry weight as soil trapped among the rhizoids, whereas H. cupressiforme has less than 1%. But perhaps most importantly, B. argenteum required 180 hours to reach steady dryness, whereas H. cupressiforme required only 132, and this was in a moss starting with more than 5X as much water!

Figure 45. Tortula muralis in a rock crevice. Photo by Michael Lüth, with permission.

Chapter 4-1: Invertebrates: Introduction

Figure 46. Grimmia pulvinata on boulder. Michael Lüth, with permission.

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Photo by

The wonderful fauna of bryophytes led Gadsby (1926) to publish his paper, "Meanderings 'mong mosses." Even after a fire bryophytes such as Funaria hygrometrica (Figure 47) and Ceratodon purpureus (Figure 48) accumulate organic matter and dust, permitting invertebrates to colonize (Clément & Touffet 1981). Others are quick to colonize areas of harvested peat (Curry et al. 1989). Even glacial land in the Antarctic (Schwarz et al. 1993) and geothermal areas of Iceland (Elmarsdottir 2003) and Ireland (Fahy 1974) sport their own bryophyte invertebrate fauna, most likely facilitated by the ameliorating effect of the microclimate within the bryophyte clone. In the Antarctic, Sohlenius et al. (2004) found highest invertebrate densities where there were moss communities. In addition to the protozoa already discussed, these leaves are home to large numbers of rotifers, nematodes, and oribatid mites, and the associated bacteria, fungi, and algae provide their sustenance. Some of the species, particularly Sphagnum (Figure 41) inhabitants, are not found elsewhere. Many live as epiphytes on the leaf, but some live as endophytes, gaining entrance to the cells through pores in Sphagnum leaf and stem cells. These specialists are often elusive by standard sampling techniques. Nevertheless, Hingley showed that 50% of the taxa were present in a single drop of water!

Figure 47. Funaria hygrometrica, a common colonizer after fires that collects organic matter, permitting invertebrates to colonize. Photo by Michael Lüth, with permission.

Figure 48. Ceratodon purpureus, a common colonizer after fire, accumulates organic matter, permitting invertebrate fauna to develop. Photo by Michael Lüth, with permission.

Jones et al. (1994) described mosses as ecosystem engineers that provide living spaces by providing a suitable physical structure. Although Sphagnum (Figure 42) is the most cosmopolitan engineer, bryophytes create habitats for invertebrates in many ecosystems. Sayre and Brunson (1971) compared the moss inhabitants in a variety of habitats to determine what faunal taxa were most common (Figure 49). One of the primary determinants of faunal inhabitants is the film of water surrounding moss leaves, especially Sphagnum (Hingley 1999). Bryophyte habitats generally influence the faunal community structure based on their moisture availability. Five classes can be recognized (Hofmann 1987; Hofmann & Eichelberg 1987): I II III IV V

Submerged mosses Mosses that are permanently moist Mosses that are only rarely dry Mosses that are frequently dry Exposed mosses that are often dry for long periods

In desert cryptogamic crusts, bryophytes seem to be important to the soil fauna (Brantley & Shepherd 2004). Among these invertebrates are arachnids, mites, nematodes, springtails, tardigrades, and other small arthropods. Mixed lichen and moss patches supported 27 taxa at sites in New Mexico, whereas mosses had 29 taxa. Abundance and diversity were higher in winter than in summer, most likely due to a lower water stress. Even the moss Syntrichia ruralis var. pseudodesertorum (Figure 50) may have its own invertebrate community (Kaplin & Ovezova 1986; Ovezova 1989). In Vaccinium heaths, the moss litter is difficult to break down (Frak & Ponge 2002). The invertebrate fauna process the litter, convert it to animal feces, and transform the soil to mor.

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Chapter 4-1: Invertebrates: Introduction

Figure 49. Mean population numbers of faunal groups from 3 2.5-cm diameter cores per moss sample, plotted on a logarithmic scale. Samples represent a variety of habitats from 26 locations in Maryland and Virginia, USA. Redrawn from Sayre & Brunson 1971.

Figure 51. Calliergon sarmentosum, a common component of the moss-invertebrate community in the Antarctic. Photo by Michael Lüth, with permission.

The invertebrate representation can be more limited in the Antarctic than in many other parts of the world. Schwarz et al. (1993) found that the moss-dominated flushes near the Canada Glacier supported a community where Protozoa, rotifers, worms, and tardigrades dominated, with all but the Protozoa occurring at 5-10.83 mm depth in the moss. Following melt, more of the organisms were found in the upper 5 mm of the moss habitat. Mites occurred in lesser quantities and Collembola were nearly absent. On the other hand, a catenulid flatworm in that habitat was a rare find; microturbellarians are quite rare in Antarctica. Bryophytic epiphytes are important habitats for invertebrates. Kellar (1999) and Milne and Short (1999) demonstrated this for Dicranoloma in the cool temperate rainforest of Victoria, Australia. Nadkarni and Longino (1990) have demonstrated this for the neotropics. Lobules as Habitat

Figure 50. Syntrichia ruralis var. pseudodesertorum may have its own invertebrate community. Photo by Paul Slichter, with permission.

In the Antarctic, the structure of the mosses [Calliergon sarmentosum (Figure 51), Drepanocladus sp. (possibly Sanionia uncinata)] provides a complex community where epiphytic algae and invertebrates form a higher diversity than the surrounding algal community (Priddle & Dartnall 1978). For example, Calliergon sarmentosum provides the site of most abundant algae in leaf axils. Six stem zones result from deterioration of basal portions. Benthic invertebrates move actively among these mosses. Six species of rotifers are common in the middle stem zones where there is the greatest abundance of epiphytes. Of these, two colonize the bare underside of leaves whereas four live mostly in leaf axils. Windinduced mixing in the summer provides transportation for at least some of the epiphytes from the shallow portions of the lake. Rotifers settle there as larvae.

As discussed in the chapters on micro-organisms and rotifers, the water-holding lobules of some leafy liverworts may house a variety of invertebrates. In fact, these invertebrates seem in some cases to be attracted to the plants and readily enter the lobules (Hess et al. 2005). In the leafy liverwort Pleurozia purpurea (Figure 52-Figure 53), the fauna include Ciliata, Rhizopoda (protozoans), flatworms, nematodes, annelids, rotifers, tardigrades, and copepods. A detailed discussion of the "trapping" mechanism of the lobules is in sub-Chapter 2-6 on protozoa. Whether these invertebrates are truly trapped and consumed by the liverworts remains unknown. Decaying inhabitants provide food for other members of the community and provide a proximal source of nutrients for the liverwort leaves. These organisms form a unique faunal community where organisms live, consume, die, and decay. Aquatic Bryophytes can offer communities that mimic those of riffles, or house very different communities. In her study of the River Liffey, Ireland, Frost (1942) found that the

Chapter 4-1: Invertebrates: Introduction

numbers of organisms found in 23 bryophyte samples differed little between an acid (ca 282,000 organisms) and an alkaline (ca 306,900 organisms) stream, but the composition of the organism differed. On the other hand, Elgmork and Sæther (1970) found that at least some species exhibited larger numbers of individuals at locations with moss cover on the stones than those without mosses, suggesting that the mosses could accommodate a much larger number of invertebrates.

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Minckley suggested that those animals that were relatively scarce in the moss beds but much more abundant in the rubble of smaller riffles may have been driven there by the preference of crustaceans for the mosses. Inhabiting the riffles permitted the smaller invertebrates to avoid being dinner for the crustaceans.

Figure 54. Fissidens fontanus, an aquatic moss that creates a quiet refuge in the middle of riffles. Photo by Tan Sze Wei Aquamoss website , with permission. Figure 52. The leafy liverwort Pleurozia purpurea, showing the protective nature of the curved leaves. The lobules are underneath. Photo by Sebastian Hess, with permission.

Figure 53. Left: Worm, probably an oligochaete, from the lobule of the leafy liverwort Pleurozia purpurea. Right: Lobule of the liverwort, Pleurozia purpurea. Photos by Sebastian Hess, with permission.

In a study of Doe Run, Meade County, Kentucky, USA, Minckley (1963) found that the invertebrate abundance in beds of the moss Fissidens fontanus (Figure 54) "strongly reflected the fauna of unvegetated riffles." This seems to be almost a contradiction since the same study demonstrated that the closely matted F. fontanus created a "pool environment in the midst of riffles."

Kinchin (1992) considered the faunal inhabitants to grade from unspecialized among the submerged mosses to more specialized, drought-resistant or drought-tolerant toward the dry end. Carpenter and Lodge (1986) found that submerged plants, including bryophytes, affect the physical environment through light extinction, temperature modulation, hydrodynamics, and substrate. They alter the chemistry by providing oxygen, altering inorganic and organic carbon, and sequestering nutrients. Nevertheless, some habitats, while appearing suitable, are not colonized by any species. Aquatic bryophytes in streams generally house the largest and probably the most diverse fauna among the various stream communities (see e.g. Percival & Whitehead 1929; Frost 1942; Badcock 1953; Hynes 1961; Minckley 1963; Thorum 1966; Stern & Stern 1969; Michaelis 1977; Cowie & Winterbourn 1979; Carpenter & Lodge 1986; Suren 1988, 1991a, b; Vlčková et al. 2001/2002; Paavola 2003). Amos (1999) described the torrent among the Fontinalis branches (Figure 55) in a poetic fashion: "All was quiet at the bottom of the torrent moss world, despite the storm of rushing water overhead." Here one could find zones of algae – diatoms, desmids, and filamentous species. Inhabitants included round and segmented worms, rotifers, gastrotrichs, water fleas, copepods, scuds, and a variety of larval insects as well as adults of tiny species. The mountain midge larva anchors there with suction cups that are even better than those of the squid and octopus. Yet Kinchin (1990b, 1992) paints a different picture of the waterfalls in Ein Gedi Nature Reserve, Israel, where the fauna is relatively poor.

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Chapter 4-1: Invertebrates: Introduction

Figure 55. Fontinalis antipyretica houses a wide range of invertebrates in streams and lakes, giving them a refuge from rapid flow and predators. Photo by Michael Lüth, with permission.

Specificity for particular bryophytes may be more a result of the habitat where each bryophyte lives. Paavola 2003) attempted to show the relationship between bryophytes, macroinvertebrates, and fish, with a goal to show concordance and usefulness in predictive power. Bryophytes and macroinvertebrates showed a weak congruence with weak predictive power, but neither had a good congruence with fish. Cowie and Winterbourn (1979) found distinct preferences for certain bryophyte species among the invertebrates in a New Zealand stream, but these differences also reflected habitat differences such as position in sream. Fissidens rigidulus occurred in the torrential water in mid channel. Pterygophyllu quadrifarium occurred where it was water saturated by the inner spray zone of a waterfall. Cratoneuropsis relaxa grew in the outer spray zone. Cowie and Winterbourn suggested that the invertebrates responded to differences in water saturation, flow rates, and detritus-trapping ability by the mosses, the latter also relating to flow rate but including aspects of the moss morphology. In aquatic habitats, bryophytes are particularly important in contributing to faunal diversity (Priddle & Dartnall 1978; Suren & Winterbourn 1992a). In the Antarctic, these faunal groups are dominated by Protozoa, Rotifera, Nematoda, Turbellaria, Tardigrada, Oligochaeta, and Acari (Ingole & Parulekar 1990). In alpine streams of New Zealand, bryophytes provide shelter with reduced flow (Suren 1991b) and catchment for algae and detritus, thus creating a habitat with both shelter and food (Suren 1992), and in some cases materials for constructing larval cases (Suren 1987). Among 23 invertebrate taxa, 14 were found with bryophyte fragments in their gut, but their presence in the gut was only common in several of the aquatic insects (Suren & Winterbourn 1991). Bryophytes contained more indigestible compounds than did other plants, making them less nutritious. Rather, it appears that detritus and periphyton were the primary food sources (Suren & Winterbourn 1992b). In these New Zealand streams, the bryophyte faunal communities were greater in streams above the treeline (Suren 1993). Greater invertebrate density occurred within bryophyte communities with periphyton than those with detritus (Suren 1993). Bryophyte communities were dominated by aquatic insects and Nematoda, oribatid mites, Hydracarina, Copepoda, and Ostracoda (Suren

1988). When artificial mosses were used in place of real ones, similar invertebrate communities developed, but some, e.g. Nematoda, Acarina, Tardigrada, Ostracoda, seemed to suffer from loss of the food supply (Suren 1991a). Linhart et al. (2002) examined the fauna of Fontinalis antipyretica (Figure 55) growing on rocks used to stabilize a side channel of the Morava River in the Czech Republic. The means of moss-dwelling meiofauna were 253,917 ± 178,335 (± SD) per 10 g dry weight of moss and 7,160,461 ± 5,029,047 per 1 m2 of the bottom area during October 1999-November 2000. Bdelloidea (rotifers) formed the dominant group (76%), followed by Monogononta (rotifers) (11.23%), Nematoda (6.38%), Chironomidae (midges) (4.08%), and Oligochaeta (worms) (1.06%). Linhart and coworkers (2002) considered that fine particulate matter trapped by the mosses would serve as both a habitat and a food source. They found that about 4% of the trapped matter was coarse matter (500-1000 µm), 14% medium (10-500 µm), and 82% fine (30-300 µm). Only 10% of the trapped matter is organic. The size and content of the trapped matter were significantly correlated (P30 m) may account for differences in nematode densities. At locality 1, mosses trapped 19 times as much FPOM as the gravel and 3 times as much as the moss at locality 2. Likewise, nematodes at locality 2 comprised only 11% of the meiofauna. Everybody has to eat! Even aquatic habitats dry out from time to time. Aquatic moss-dwelling nematodes are among the dominant

Figure 73. Nematode from the terrestrial moss Sanionia uncinata on the Barton Peninsula of King George Island, Antarctica. Photo by Takeshi Ueno, with permission.

Spaull (1973) found 30 species in 19 genera among mosses on Signy Island, with summer population densities of 0.48 x 106/m2 in the upper 6 cm of Chorisodontium (Figure 38)-Polytrichum (Figure 74) turf compared to 7.47 x 104/m2 in soil beneath the grass Deschampsia antarctica. Nevertheless, in alpine areas in Schistidium apocarpum (as S. grande; Figure 75), Hoschitz (2003) and in the Antarctic (Figure 76; Caldwell 1981a, b), bryophytes and lichens provide a protected shelter in which nematodes may survive. In the Austrian Alps, Plectus sp. (Figure 3) and Eudorylaimus sp. (Figure 70) survive the extreme conditions of the Alps. Plectus murrayi (Figure 77) is likewise a moss inhabitant at Victoria Land in the Antarctic (Melianie Raymond, pers. comm. 2008). Teratocephalus tilbrooki and Plectus antarcticus coexist in the shelter of moss cushions and mats (Pickup 1990b) and were the most abundant taxa on Signy Island in the Antarctic (Spaull 1973). However, on Signy Island Plectus (Figure 3)

Chapter 4-3: Invertebrates: Nematodes

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reaches its greatest abundance in moss carpets and Teratocephalus (Figure 17) in moss turf, suggesting that moss form plays a role, most likely in moisture relations, but possibly also in temperature relations.

Figure 77. Two individuals of Plectus murrayi, an Antarctic endemic that is often found in moss beds. Photo by Melianie Raymond, with permission. Figure 74. Polytrichum strictum in Alaska, a moss where nematodes are known to live in the upper 6 cm in the Antarctic. Photo by Andres Baron Lopez, with permission.

Figure 75. Schistidium apocarpum, a moss that provides a survival refuge in the Antarctic and alpine areas. Photo by David T. Holyoak, with permission.

Figure 76. Moss (reddish) and lichens. This photo shows a typical habitat for Plectus murrayi and occasionally Panagrolaimus davidi and Eudorylaimus antarcticus. The photo was taken near Gondwana Station, Terra Nova Bay, Victoria Land. Photo by Melianie Raymond, with permission.

The common presence of Teratocephalus (Figure 17) seems to be unique to the Antarctic, where it is abundant in the moss turf (Spaull 1973). It survives the frigid cold by a fast dehydration strategy that reduces damage by ice crystals (Wharton 2003). It would be interesting to determine how this fast dehydration relates to its choices of moss species/form. Ditylenchus sp. B occurs in more exposed aerial thalli of lichens (Spaull 1973). The latter species exhibits supercooling ability, whereas the mossdwelling species both have bimodal supercooling point distributions. The high group supercools to ~-7°C and the other at ~-22°C. Pickup (1990b) suggests that field temperatures are likely to reach even lower levels than that. Spaull (1973) found Teratocephalus, Plectus (Figure 3), and Eudorylaimus (Figure 70) in all the bryophyte sampling locations on Signy Island, with the former two accounting for more than 50% of the nematodes among mosses. Cushion-formers such as Andreaea (Figure 78) and Grimmia, on the other hand, had a nematode community where Plectus comprised less than 3%. A similar small percentage of Teratocephalus occurred in Bryum. Eudorylaimus is more abundant in moss carpets and cushions than elsewhere. Eudorylaimus sp. C, in particular, seems to prefer cushions of Andreaea (Figure 78), Grimmia, and Tortula, where it comprises 45% of the individuals in that genus, but it is rare elsewhere (Spaull 1973). Antarctenchus hooperi is less restricted, being common in cushions of Andreaea and Tortula and in carpet-forming Calliergon (Figure 37)-Calliergidium (probably Warnstorfia austrostraminea), but it is likewise rare or absent elsewhere. The tylenchids [Antarctenchus, Aphelenchoides, Ditylenchus, Tylenchus (Figure 18)] are more abundant in moss turf than elsewhere, whereas the monhysterids [Monhystera (Figure 16), Prismatolaimus] are less numerous in moss turf than in other bryophyte formations.

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Chapter 4-3: Invertebrates: Nematodes

possible that these may reflect differences in temperature that would affect rate of development. Competition with tardigrades that share their food sources seems also to be a limiting factor within a cushion.

Figure 78. Andreaea gainii (blackish) in Antarctica, showing cushion growth where nematodes may lurk. Photo from Polar Institute through Creative Commons.

The genus Eudorylaimus is particularly common in the Antarctic. Melianie Raymond (pers. comm. 2008) found Eudorylaimus antarcticus (Figure 79) among mosses in the Antarctic. In the McMurdo Dry Valleys, Eudorylaimus species are unaffected by vegetation type, including bryophytes (Simmons et al. 2009). Plectus (Figure 3) species, although bryophyte dwellers, are more abundant in algae. Its abundance above ground and below ground were significantly correlated in both the microbial mats and mosses. That is, the above ground abundance was a good indicator of below-ground abundance. The ability of Plectus species to migrate vertically is likely to benefit it in this changeable and extreme climate (Overgaard-Nielsen 1948; Kinchin 1989). Kito et al. (1996) found a new species of Eudorylaimus (E. shirasei), bringing the Antarctic total in that genus to seven. Some of the specimens for this new species were collected from moss clumps at Cape Ryugu on the Prince Olav Coast, East Antarctica. It is odd among the members of Eudorylaimus (Figure 70) in having multinucleate intestinal cells, a factor that could simply have been overlooked elsewhere, but that raises questions about the possible effects of the severe Antarctic climate in causing or selecting for this multinucleate state. New species of moss nematodes will most likely continue to be described, particularly in the Antarctic. Sohlenius and Boström (2006) found that 64% of 91 moss cushion samples from nunataks in East Antarctica had nematodes in them. In this harsh environment, 8% of the samples had no microfauna (nematodes, rotifers, or tardigrades) at all. The researchers considered the patchy distribution of nematodes and other organisms among the mosses to be a product of patch dynamics where stochastic processes determined colonization. They further supported this notion with the fact that nematodes in different cushions had different developmental stages, but it is

Figure 79. Eudorylaimus antarcticus, a common nematode among Antarctic mosses. Photo by Melianie Raymond, with permission.

In nunataks of Vestfjella, Heimefrontfjella, and Schimacher Oasis in East Antarctica, the faunal communities associated with mosses lacked organization and represented early stages of succession (Sohlenius et al. 2004). In these exposed nunatak moss habitats, species of Plectus (Figure 3) and Panagrolaimus (Figure 20) were the most frequent of the nematodes, occurring in 26% and 5% of the samples, respectively.

Dangers Lurking among Bryophytes Fungal Interactions Who would think that fungal treachery looms amid the mosses! Although nematode-trapping fungi are known worldwide, they were unknown in the Antarctic until 1973. In their examination of Signy Island mosses, Duddington et al. (1973) found nematode-trapping fungi on a number of moss species: Brachythecium austrosalebrosum, Calliergon sarmentosum (Figure 37), Sanionia uncinata (Figure 80) (all hydrophytic), and Andreaea depressinervis (mesophytic-xerophytic). These fungi sport rings (Figure 81) that are able to constrict around nematodes that wander through them, thus ensnaring them. Several specimens of the predatory Thyronectria antarctica var. hyperantarctica had indeed trapped nematodes within their mossy home. Spaull (in Duddington et al. 1973) also noted fungi with such loops in a sample of the leafy liverwort Cephaloziella sp. (Figure 82) mixed with the lichen Cladonia metacorallifera from Terra Firma Islands in Marguerite Bay (latitude 68º42'S).

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frequent of the endozoic taxa was Harposporium anguillulae (Figure 83). These fungi seemed to have some bryological preferences, with M. ellipsosporum preferring calcicolous mosses. In fact, it appears that acidic habitats might provide a safe haven - the nematophagous fungi were absent from permanently saturated moss carpets and the strongly acidic turf-forming mosses of Polytrichaceae.

Figure 80. Sanionia uncinata, common home of nematodes and nematode-trapping fungi. Photo by Michael Lüth, with permission.

Figure 83. Harposporium anguillulae, fungal parasite with conidiophores and conidia, on a dead nematode. Photo by George Barron, with permission.

Figure 81. Nematode-trapping fungus, possibly Monacrosporium cionopagum, isolated from the moss Calliergidium cf. austro-stramineum on Signy Island in the Antarctic. Redrawn from Duddington et al. 1973.

Figure 82. Leafy liverwort Cephaloziella turneri, member of a genus that is home to nematode-trapping fungi. Photo by Michael Lüth, with permission.

The Antarctic sports at least 18 taxa that either trap nematodes or become endozoic parasites of members of this phylum (Gray et al. 1982). Many of these have been found among the mosses. Among the Hyphomycetes that snare nematodes, Monacrosporium ellipsosporum and M. cionopagum were the most widely distributed. The most

These ensnaring fungi are not restricted to the Antarctic. Duddington (1951) considered the abundance of such fungi among mosses to result from the large amount of water among the shoots and leaves, making the environment favorable for both nematodes and fungi. In the Antarctic, the mosses provide the added benefit of being warmer than the air in summer. Both nematodes and fungi live among Sphagnum (Figure 5). And here we also find nematode ensnaring fungi. In particular, the genus Sporotrichum (Figure 84), known for causing sporotrichosis in those who handle Sphagnum, is able to trap the nematodes that reside there (Dollfus 1946).

Figure 84. The nematode-ensnaring fungus Sporotrichum sp. in action. This is the same genus known so well for causing sporotrichosis in people who work with Sphagnum. Image from Dollfus 1946.

Other fungal treachery looms, although not so dramatically. Several species of nematode-dwelling parasites await. Among these on Signy Island in the Antarctic are Harposporium sp. (Figure 83) and Acrostalagmus sp.

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Chapter 4-3: Invertebrates: Nematodes

The widespread fungus Catenaria anguillulae (Figure 85-Figure 88) parasitizes nematodes (Sayre & Keeley 1969). Its zoospores (swimming spores) are attracted to the nematodes by exudates from the mouth, anus, or other opening of the nematode, including wounds. Once attached, the zoospores encyst, typically in clusters. These eventually germinate and penetrate through the nearby orifice to attack their host, the nematode. Success of the fungus is favored by high temperatures (optimum at 28°C) and moisture, the latter provided by bryophytes.

Figure 88. Zoosporangia of Catenaria anguillulae within a nematode. Red arrows indicate the exit tubes where zoospores escape. Photo by George Barron, with permission.

Safe Site from Predation

Figure 85. Nematode with zoospores of fungus Catenaria anguillulae surrounding its mouth. Photo by George Barron, with permission.

One advantage to living in a habitat with only small chambers is that large organisms don't fit. This affords some protection from predation, but nematodes are definitely not free from it. Some are preyed on by cohabiting tardigrades (Doncaster & Hooper 1961); under experimental conditions, one tardigrade, Macrobiotus richtersi (Figure 89), consumed 61 nematodes per day – no small threat (Sánchez-Moreno et al. 2008). Others must surely fall prey to insects. Even the protozoa may be a threat (Yeates & Foissner 1995). The Testacea (amoebae) can ingest nematodes, attacking mostly from the tail. In New Zealand, it was the protozoa Nebela (Apodera) vas (Figure 90) and Difflugia sp. (Figure 91) that waged the attacks, mostly on Dorylaimus (Figure 7) and Plectus (Figure 3) species among common bryophyte inhabitants.

Figure 86. Nematodes showing infestation by Catenaria anguillulae. Modified from George Barron's image, with permission. Figure 89. Macrobiotus richtersi, a moss-dwelling tardigrade that devours numerous nematodes. Photo through Creative Commons.

Pollution

Figure 87. Zoospore of Catenaria anguillulae. Photo by George Barron, with permission.

Even aquatic organisms can suffer from air pollution. Steiner (1995b) tested responses of several groups of aquatic moss-dwelling invertebrates to SO2 pollution. Nematodes, rotifers, and tardigrades changed their community composition. SO2 at 0.225 ppm for 18 months significantly reduced the numbers of several nematode species. Responses were not so clear at 0.075 ppm, with some species increasing and others decreasing in numbers.

Chapter 4-3: Invertebrates: Nematodes

Lead can also considerably alter the moss-dwelling nematode community. Zullini and Peretti (1986) found that increased lead content in the moss resulted in a significant decrease in diversity, richness, and biomass, but not the density. The Dorylaimina suborder suffered the most by far.

Figure 90. Nebela (=Aphodera) vas, a protozoan that is a nematode predator. Photo by Edward Mitchell, with permission.

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Members of Plectus are quick driers. Acrocarpous cushions are more favorable habitats than pleurocarpous feather mosses. Slow dehydration is important to their survival in a state of anhydrobiosis; some achieve this by coiling. Water is also necessary for their motility, where they can swim, crawl, inch, or bend to move. Some survive by living and reproducing inside the hyaline cells of Sphagnum. Eggs likewise have a long survival and can even survive lack of oxygen. Food strategies are mostly bacteriovores and predators. Some are mycophagous or saprophytic. Woodland mosses often feed on the detritus. They seem to do best in habitats with a low C:N ratio in the food source. Stream mosses serve as nutrient traps that favor nematodes. Bryophytes can provide a safe site against wouldbe predators. However nematode-trapping fungi and fungal parasites may loom there. Bryophytes can also make a safe site by buffering the temperature both in the bryophyte and in the soil beneath. Even antheridia can serve as habitat, and in other cases the nematodes nestle among archegonia to make nematode galls. Galls seem to occur on many species of bryophytes and house nematodes that are often less than 1 mm long. Numbers usually are highest in summer and lowest in winter, with some species migrating to greater depths in winter. Some species among Panagrolaimus can freeze and recover. Others, such as one Aphelenchoides, can tolerate temperatures ranging from meltwater to 61.3ºC. Trehalose can protect some from freezing damage as well as from dehydration damage, most likely by stabilizing membranes.

Acknowledgments Figure 91. Difflugia bacillifera, a moss-dwelling protozoan that preys on nematodes. Photo by Edward Mitchell, with permission.

Summary Among the most common bryophyte-dwelling nematodes are members of the genera Plectus and Eudorylaimus. These nematodes are usually less than 1 cm in length and many are much smaller. Although bryophyte-inhabiting nematodes are rarely studied, they are common there and can reach 480 individuals in just 1 g of moss. Many nematodes adhere to the mosses with an adhesive organ. Water is their most limiting factor. They can migrate vertically among the bryophytes to adjust their moisture level. Some migrate from rhizoids to canopy when the moss is too wet, some move from the rhizoids to the stems when the moss is saturated, and some never leave the rhizoids. The most specialized nematodes, such as Plectus rhizophilus, live in the bryophytes that experience the most events of desiccation, such as the epiphytes.

Jan-Peter Frahm helped me obtain the photographs of the nematode and Pleurozia locules. Aldo Zullini gave me a valuable critique of an early version, provided images, and suggested some older literature I would probably not have found otherwise. George Barron helped me sort our the fungal stories. Tom Powers provided me with additional sources of images, helped with nomenclature, and gave me permission to use the images on the website. Helen Jolley provided the story of nematode galls on Stonea. Melianie Raymond provided me with images and information to tell the Antarctic story. Bryonetters have been wonderful in making their photographs available to me and seeking photographs from others.

Literature Cited Allgén, C. A. 1951. On some species of freshwater and terrestrial nematode genera, found inhabiting southern marine waters. K. fvsiogr. Sällsk. Lund Förh. 21(19): 177184. Asthana, G. and Srivastava, S. C. 1993. Nematode galls on Cheilolejeunea cf. giraldiana (Mass.) Mizut. (Lejeuneaceae) from Tamil Nadu, India. Lindbergia 18: 94-96. Bamforth, S. S. 2003. Water film fauna of microbiotic crusts of a warm desert. J. Arid. Environ. 56: 413-423.

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Barbuto, M. and Zullini, A. 2006. Moss inhabiting nematodes: influence of the moss substratum and geographical distribution in Europe. Nematology 8: 575-582. Barrett, J. 1982. Metabolic responses to anabiosis in the fourth stage juveniles of Ditylenchus dipsaci (Nematoda). Proc. Royal Soc. London B216: 159-177. Beasley, C. W. 1981. Some Tardigrada from Puerto Rico. Texas J. Sci. 33: 9-12. Block, W. 1985. Ecological and physiological studies of terrestrial arthropods in the Ross Dependency 1984-85. Brit. Antarct. Surv. Bull. 68: 115-122. Block, W. and Christensen, B. 1985. Terrestrial Enchytraeidae from South Georgia and the maritime Antarctic. Brit. Antarct. Surv. Bull. 69: 65-70. Boag, B. and Yeates, G. W. 2004. Population dynamics. In: Gaugler, R. and Bilgrami, A. L. (eds.). Nematode Behaviour. CABI Publishing, Cambridge, MA, pp. 345-370. Brown, I. M., Wharton, D. A., and Millar, R. B. 2004. The influence of temperature on the life history of the Antarctic nematode Panagrolaimus davidi. Nematology 6: 883-890. Brzeski, M. 1962a. The nematodes of the peat-mosses in Kosciehska Valley (the western Tatra). Acta Zool. Cracov. 7(1): 23-37. Brzeski, M. 1962b. Nematodes of peat-mosses of the Bialozieza forest. Acta Zool. Cracov. 7(1): 53-62. Bunt, J. S. 1954. The soil-inhabiting nematodes of Macquarie Island. Austral. J. Zool. 2: 264-274. Caldwell, J. R. 1981a. Biomass and respiration of nematode populations in two moss communities at Signy Island, maritime Antarctic. Oikos 37: 160-166. Caldwell, J. R. 1981b. The Signy Island (South Orkney Islands) terrestrial reference sites: XIII. Population dynamics of the nematode fauna. Bull. Brit. Antarct. Surv. 54: 33-46. Cameron, R. E., King, J., and David, C. N. 1970 Microbiology, ecology and microclimatology of soil sites in dry valleys of southern Victoria Land, Antarctica. In: Holdgate, M. W. (ed.). Antarctic Ecology. Academic Press, London & NY, pp. 702-716. Clegg, J. S. 1973. Do dried cryptobiotes have a metabolism? In: Crowe, J. H. and Clegg, J. S. (eds.). Anhydrobiosis. Dowden, Hutchinson & Ross Inc. Coleman, D. C. 1971. Numbers and biomass of soil nematodes of two South Carolina old fields. Amer. Midl. Nat. 85: 262265. Crowe, J. H. 1975. The physiology of cryptobiosis in tardigrades. In: Higgins, R. P. (ed.). Proceedings of the First International Symposium on Tardigrades. Memorie dell'Instito Italino di Idrobiologi 32 Suppl: 37-59. Crowe, J. H. and Madin, K. A. 1974. Anhydrobiosis in tardigrades and nematodes. Trans. Amer. Microsc. Soc. 93: 513-524. Crowe, J. H., Crowe, L., and Chapman, D. 1984. Preservation of membranes in anhydrobiotic organisms: The role of trehalose. Science 233: 701-703. Deguchi, H. 1977. Gall formation by nematodes on Racomitrium lanuginosum (Hedw.) Brid. and R. heterostichum var. diminutum. Hikobia 8: 179. Demeure, Y., Freckman, D. W., and Gundy, S. D. van. 1979. Anhydrobiotic coiling of nematodes in soil. J. Nematol. 11: 189-195. Dixon, H. N. 1905. Nematode galls on mosses. J. Bot. London 43: 251-252. Dixon, H. N. 1908. Nematode galls on mosses. Bryologist 11: 31.

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Wharton, D. A., Young, S. R. and Barrett, J. 1984. Cold tolerance in nematodes. J. Compar. Physiol. B 154: 73-77. Winslow, R. D. 1964. Soil nematode population studies. 1. The migratory root Tylenchida and other nematodes of the Rothamsted and Woburn six-course rotations. Pedobiologia 4: 65-76. Womersley, C. 1987. A reconsideration of diversity of adaptation in nematode anhydrobioses in relation to their environments. In: Vistas on Nematology. A publication celebrating the 25th anniversary of the American Society of Nematologists, E. O. Painter Publ., pp. 165-173. Wood, F. H. 1973. Nematode feeding relationships: Feeding relationships of soil-dwelling nematodes. Soil Biol. Biochem. 5: 593-601. Wright, J. C. 1991. The significance of four xeric parameters in the ecology of terrestrial Tardigrada. J. Zool. 224: 59-77. Yeates, G. W. 1967. Studies on nematodes from dune sands. 9. Quantitative comparisons of the nematode fauna of six localities. N. Zeal. J. Sci. 10: 927-948. Yeates, G. W. 1970. Two terrestrial nematodes from the McMurdo Sound region Antarctica, with a note on

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Glime, J. M. 2017. Invertebrates: Annelids. Chapt. 4-4. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. Ebook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 18 July 2020 and available at .

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TABLE OF CONTENTS Annelida – Segmented Worms ............................................................................................................................ 4-4-2 Water Relations ............................................................................................................................................ 4-4-3 Temperature Tolerance ................................................................................................................................ 4-4-3 Reproduction ................................................................................................................................................ 4-4-4 Food Relations ............................................................................................................................................. 4-4-4 Sampling ............................................................................................................................................................. 4-4-5 Habitats ............................................................................................................................................................... 4-4-5 Aquatic ......................................................................................................................................................... 4-4-5 Peatlands ...................................................................................................................................................... 4-4-5 Prairie Worms .............................................................................................................................................. 4-4-6 Antarctic....................................................................................................................................................... 4-4-6 Dispersal Agents?................................................................................................................................................ 4-4-6 Earthworm Culture .............................................................................................................................................. 4-4-7 Polychaetes.......................................................................................................................................................... 4-4-7 Summary ............................................................................................................................................................. 4-4-8 Acknowledgments ............................................................................................................................................... 4-4-8 Literature Cited ................................................................................................................................................... 4-4-8

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Chapter 4-4: Invertebrates: Annelids

CHAPTER 4-4 INVERTEBRATES: ANNELIDS

Figure 1. Aeolsoma, an aquatic annelid that sometimes inhabits mosses such as Fontinalis. Photo by Yuuji Tsukii, with permission.

Annelida – Segmented Worms Among the bryophyte-dwelling Annelida are worms that qualify as mesofauna (Figure 1). These are organisms, also including mites (Acari) and springtails (Collembola), that can occupy pore spaces that have a diameter of less than 2 mm (Briones 2006). In other words, these are small annelids, primarily in the subclass Oligochaeta. Among the annelids, the family Enchytraeidae is a worldwide but little known family that can be found among the bryophytes. They reach their greatest abundance in the moist temperate soils (Block & Christensen 1985). Unlike the large, pink-red earthworms, these worms are usually grey-white (Briones 2006). Their identification is based primarily on internal characters, hence making them unknowns to the casual observer. And they must be live to be identified because preservatives make them opaque. Enchytraeids are important consumers in the Arctic tundra sedge-moss meadow habitat (Ryan 1977). Although annelids are not as common as some other invertebrates in bryophytic habitats, there are at least some notable exceptions. Fontinalis (Figure 2) has been known to house 67 oligochaetes and 5 leeches (Hirudinea) in a square meter (Berg & Peterson in Macan 1966). Moss balls of Drepanocladus (Figure 3) and Fontinalis also house these annelids. In New Zealand Suren (1993) found oligochaetes to occupy 12.3% of the bryophyte fauna. Three of the most common Enchytraeids in peatlands are Cognettia sphagnetorum, Marionina clavata, and Achaeta eiseni (Figure 4; Briones et al. 1997; Briones pers. comm. 17 March 2009). Nevertheless, Standen and Latter (1977) demonstrated that the common C. sphagnetorum is less common among Sphagnum than it is among

Eriophorum or Calluna in a blanket bog at Moor House in Cumbria. Marionina clavata is aided in its survival by laying two types of eggs, one taking ~112 days and another taking ~271 days for the worms to reach maturity at 10ºC, thus potentially providing them with two different sets of conditions (Springett 1970). A tolerance for low pH levels in C. sphagnetorum and M. clavata (2.9-4) suggests their suitability for peatland habitation (Graefe & Beylich 2003).

Figure 2. Brook moss, Fontinalis duriaei, where annelids can be common. Photo by Janice Glime.

In a Dutch Scots pine forest these three had a vertical zonation pattern in the same order, with Cognettia sphagnetorum (Figure 5) being the first to colonize new needle litter (Didden & de Fluiter 1998).

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numbers ten times as great after 60 years. Because of a proportionally larger increase in Collembola, the proportion of Enchytraeidae in the fauna dropped slightly. More than 60% of the enchytraeids occurred in the top 4 cm of the peat. Within two years after water was returned to a drained peatland, the numbers dropped abruptly to levels near that of pre-drainage.

Figure 3. Moss ball of Drepanocladus from Lake Kucharo, Japan. Photo by Janice Glime.

Figure 5. SEM image of Cognettia sphagnetorum. Photo © María Jesús Iglesias Briones, with permission.

Figure 4. SEM image of Achaeta sp. Photo by María Jesús Iglesias Briones, with permission.

Water Relations Very small annelids (Enchytraeidae) occur among Sphagnum plants. Springett (1970) found six species associated with peat. The moisture changes can result in diurnal vertical migrations (upwards at night), at least in Cognettia sphagnetorum (Springett et al. 1970; Hingley 1993; Briones et al. 1997), a widespread species known from aquatic habitats, Sphagnum peatlands, and on South Georgia in the Antarctic from Polytrichum (Figure 6) clumps (Block & Christensen 1985). Cognettia sphagnetorum (Figure 5) has no cocoon stage, thus permitting it to take full advantage of the growing season in cold, wet climates of places like the Antarctic (Hingley 1993). Several species of Achaeta (Figure 4) are morphologically adapted to drought by having a thicker cuticle. However, it appears that physiological adaptations to drought in the enchytraeids may be limited. On the other hand, they seem also to be intolerant of too much water. In a study on the effects of drainage on the mesofauna of peatlands in Finland, Silvan et al. (2000) found that water-level drawdown resulting from peatland drainage caused an increase in the numbers of all the mesofauna studied, including the Enchytraeidae, with

Figure 6. Clump of Polytrichum that could house annelids. Photo by Michael Lüth, with permission.

Temperature Tolerance In peatlands and elsewhere, the Enchytraeidae are sensitive to temperature, which seems to be a major differentiating factor for population size. Cognettia sphagnetorum increases its reproductive rate, most likely through its capability of fragmentation as a reproductive strategy, in response to warmer temperatures (Briones et al. 1997). Warming seems to result in greater numbers without a concomitant vertical migration. Despite this advantage, Briones et al. (2007) considered that an increase in temperature to a maximum mean annual threshold of 16ºC could cause total loss of this species from some regions. Achaeta eiseni, also a peatland species, is resistant to higher temperatures, increasing in numbers as temperatures increase, whereas numbers of Cernosvitoviella atrata (Figure 7) are greatly reduced by higher temperatures (Briones 2006, pers. comm. 17 March 2009). The latter species is inhibited by its inability to avoid dry conditions, resulting in death at high temperatures (Briones et al. 1997).

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Figure 7. SEM image of Cernosvitoviella atrata. Photo by María Jesús Iglesias Briones, with permission.

Cognettia sphagnetorum and C. glandulosa (known from moss banks and elsewhere; Block & Christensen 1985) are also prepared for the seasonal inundation of the peatlands. They are able to produce red blood under very wet conditions (Healy & Bolger 1984) to survive the low oxygen conditions that arise. Healy and Bolger showed that 35% of the Irish taxa of enchytraeids preferred habitats that were submerged or frequently flooded. Reproduction Any successful inhabitant of mosses must have a life cycle that is coordinated with the moss habitat. One advantage to some Oligochaetes is their ability to reproduce by fragmentation. Christensen (1959) pointed out that the Enchytraeidae contrast with other Oligochaeta in their inability to reproduce by fragmentation. At the same time, he reported on asexual reproduction in three species among the 78 Dutch Enchytraeidae studied by that time. In fact, one species apparently had only asexual reproduction, by fragmentation. Honda et al. (2003) described fragmentation in Enchytraeus japonensis. This worm uses stem cells to accomplish its regeneration. Segments form as organs regenerate. They showed that cells with newly synthesized DNA appeared first as a ring in the tail area. The labelling then migrated, suggesting that the formation of segments occurs before organ regeneration. This regeneration cycle can take as few as ten days (Myohara et al. 1999; Nakamura 2004), and both ends of the worm can regenerate (Nakamura 2004). Nakamura (2004), in a six-and-a-half-year study, determined that the average fragmentation cycle length for the species was 20.4 days. The maximum number of fragmentation events in the life of the worm was 122, with an average of 35.3. The number of fragments in one event was 6.3. The cycle can repeat until the worm is starved or the population density is low, at which time it will differentiate gonads and reproduce once sexually (Honda et al. 2003). At this time I don't know how the number of annelid species using fragmentation relates to bryophytes as a habitat. Food Relations Springett and Latter (1977) experimented with various fungal diets on agar and found they could not keep many Cognettia sphagnetorum alive on the combinations they

tried. Exudates from the mycelia of Basidiomycetes proved most harmful, resulting in 100% mortality in 20 days. They concluded that micro-organisms did not form any part of the natural diet of moorland Enchytraeidae. Hingley (1993) considered peat to be a poor source for food (Hingley 1993), with the moss itself seemingly of poor quality for annelids; only stem material of Sphagnum has been found in gut analyses (Figure 8; Standen & Latter 1977). Nevertheless, these worms feed on items that are generally unpalatable to other animals (Hingley 1993). After these are processed by the annelids, the feces are colonized by fungi and bacteria, which are in turn ingested by Protozoa, rotifers, and nematodes. Hence a food web emerges and peat is processed. Briones (pers. comm.) challenged the suggestion that peatlands offered poor food quality, stating that enchytraeids are known to consume bacteria and dead organic matter, both of which are associated with the peatlands. Briones et al. (2004) used 14C to match the gut contents with the substrate and found that most of the assimilated food came from sediment that is 5-10 years old. Their vertical movements in response to changing moisture did not affect their food source, but at higher temperatures it seemed that they had altered their carbon source since there was a lower 14C enrichment with depth.

Figure 8. Stem section of Sphagnum contortum, like those found in an annelid gut. Photo by Michael Lüth, with permission.

Guts from worms in substrata of Sphagnum, Calluna, and Eriophorum at Moorhouse, Great Britain, all contained mixed decomposing litter, including cellulosic or humified plant material, amorphous humus, and associated fungal mycelia, again suggesting equal nutritional availability in the peatlands (Standen & Latter 1977). The Sphagnum stem material extracted from the gut of Cognettia sphagnetorum (Figure 5) causes one to question if these stems provide nutrition or merely serve to help in grinding other foods, much like the role of sand. In any case, the very high numbers of worms reached in peatlands provides witness that these are not bad systems for enchytraeids (Briones pers. comm.). In the blanket bog at Moor House, Great Britain, the numbers of Cognettia sphagnetorum were significantly less in Sphagnum than they were in Calluna and Eriophorum, suggesting that Sphagnum was not an ideal habitat. However, when these were converted to numbers

Chapter 4-4: Invertebrates: Annelids

per gram dry weight of substrate, there were no significant differences among substrata. The species was in greatest numbers in association with older decomposing litter of Eriophorum and Calluna and with surface layers of Sphagnum. The numbers of worms correlated weakly with unstained fungi, cocci, and moisture.

Sampling Annelids are generally extracted from core samples. Researchers typically use some modification of a Berlese funnel (Didden et al. 1997; See Chapter 4-1). For annelids, a wet funnel is the most common, as suggested by O'Connor (1955) and Overgaard-Nielsen (1948, 1949). The moss samples are placed in a water-filled funnel and the temperature is gradually increased to about 40ºC (~3 hours). The high temperature causes the worms to vacate the mosses and drop down to the funnel. In organic soils, the efficiency is often 95% or more (Healy 1987), but can be less than 50% in some samples (Willard 1972 in Didden et al. 1997). Variations on this include soil cores in an earthenware cylinder suspended over a heated water bath (O'Connor 1955). The worms are driven upward to a layer of cool sand on top of the soil core. The worms are recovered by washing them from the sand. An alternative method is to squeeze water from the mosses onto a microscope slide or into a Petri dish (Hingley 1993). Repeated extraction can be accomplished by soaking the moss in water and squeezing again, repeating this for a standard number of times. A paint brush or strip of filter paper can be used to transfer them to a drop of water on a slide. The sample could be transferred to a test tube, then centrifuged. A concentrated sample can then be removed from the bottom of the test tube with a long pipette. Andrew and Rodgerson (1999) tested three methods of extracting invertebrates from Tasmanian bryophytes: Tullgren funnels, sugar flotation, and kerosene phase separation. When two samples were combined, the kerosene phase separation method extracted more total individuals, more mites, and more Collembola. Nevertheless, only three of the nine taxa were found in the single samples, suggesting that replicate samples are needed. Andrew and Rodgerson attributed this to differences caused by spatial scales. They further found that there is site scale variation at 2 km or less that may be more important that altitudinal variation.

Habitats Aquatic Aquatic bryophytes can serve as annelid (subclass Oligochaeta) habitat, especially for Naididae, reaching as much as 33% of the invertebrate fauna (1968 per dm2) in thick moss vegetation of streams in the West Riding of Yorkshire, UK (Percival & Whitehead 1929). Their numbers were exceeded only by the Chironomidae (midges). This is a sharp contrast to their apparent absence on Potamogeton in those streams. Brusven et al. (1990) found that annelids were the most common non-insect invertebrate in the South Fork of the Salmon River, Idaho, USA. In Brazil, Gorni and da Gama Alves (2007) collected Fissidens and Philonotis (Figure 9) in winter and spring. Bryophytes adhering to rocks in the rapids of the Jacaré

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Pepira River, Brotas, São Paulo, Brazil, and to a vertical rock wall of a waterfall near the river provided a home for 191 Naididae individuals of Nais communis, Pristinella jenkinae, and P. menoni. Among the identifiable species, P. jenkinae was dominant, representing 96.8% of all individuals. This species occupied both the submerged mosses of stream beds and the rock wall mosses with little water. But often the annelids are not very common. In Fontinalis antipyretica in the Czech Republic, Vlčková et al. (2001/2002) found that only about 1.1% of the fauna were annelids in one stream and about 1.4% in another.

Figure 9. Philonotis fontana, representing a genus where Nais communis, Pristinella jenkinae, and P. menoni dwell in Brazil. Photo by Michael Lüth, with permission.

Naididae occupancy of mosses may provide several benefits to these worms. Mosses provide a safe site where the current is reduced in fast water (Vlčková et al. 2001/2002; Habdija et al. 2004). This is important for a group of organism that lack any adaptations for clinging or anchoring. Abundance and diversity are likely to increase with an increase in moss biomass, and more biomass makes available more periphyton and detritus (Egglishaw 1969; Suren 1993; Vlčková et al. 2001/2002; Linhart et al. 2002a, b). Like Thienemann (1912), I rarely found oligochaetes among the bryophytes in Appalachian Mountain, USA, streams (Glime 1968). But Percival and Whitehead (1929) found that Eiseniella teträedra was a frequent inhabitant among the mosses in shallow water (3-4 cm). Nevertheless, even in thick moss beds, it reached a density of only 6 per dm2. The Naididae (Nais elinguis), on the other hand, reached as many as 12,000 per dm2 among the thick moss beds. Thickness of moss growth, as well as time of year and recent history of river conditions, influenced the density of oligochaetes. Percival and Whitehead suggest that the much smaller numbers of these naidids in the loose moss mats may be due to "feeble" setae and no ability to attach to the moss. Hynes (1961) compared the oligochaetes, including Eiseniella teträedra, on mosses and silk in a Welsh mountain stream and found little difference in the percentage of organisms, suggesting that the moss need not be a living organism and might only provide a substrate, perhaps with trapped detritus as a food source. Peatlands Unlike many other kinds of animals, the annelids are not very diverse in peatlands. Hingley (1993) reported that

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only three families of Oligochaeta occur in peatlands, with the most common being the Enchytraeidae. Duinen et al. (2006) found that in Estonia and The Netherlands, only Cognettia sphagnetorum occurred in ombrotrophic raised bogs, i.e., in the most nutrient-poor situations. In Estonia, Nais variabilis (Figure 10), Lumbriculus (=Lumbricus) variegatus (Figure 11), and species with sexual reproduction occur only in more minerotrophic water bodies with a higher decomposition rate and consequent higher nutrient content. The lagg zone (marginal area around the bog where nutrients are often higher) fares somewhat better, having ten species of oligochaetes. This zone is absent in The Netherlands due to agriculture.

under moss mats when looking for moss-feeding beetles in the Byrrhidae. In drier times it can burrow down as much as 5 m.

Figure 12. The giant Palouse earthworm (Driloleirus americanus), an endangered worm that seems to seek moisture under mosses in the Palouse Prairie. Photo by Yaniria Sanchez-de Leon, with permission.

Figure 10. Nais variabilis, a moss-dwelling annelid. Photo by Yuuji Tsukii, with permission.

Antarctic As in the peatlands, the Enchytraeidae are common in the Antarctic bryophytes. Block and Christensen (1985) found Cognettia sphagnetorum in Polytrichum clumps and C. glandulosa in moss banks. On South Georgia and Signy Island, they found seven taxa in soil and peat, but suspected that five of those had been introduced by human activity on the islands.

Dispersal Agents? The presence of bryophyte diaspores in earthworm castings suggests a possible dispersal mechanism (During et al. 1987). Van Tooren and During (1988) found various spores and vegetative diaspores in the guts of terrestrial earthworms [Allolobophora caliginosa, A. chlorotica, and Lumbricus terrestris (Figure 13-Figure 14)] in The Netherlands. Especially rhizoid tubers and spores occurred. However, it is not clear that these provided any nutritional value to the worms because some remained viable and grew new plants, suggesting digestion was not possible. Rather, they most likely were simply mixed in with the soil that was being consumed.

Figure 11. Lumbriculus (=Lumbricus) variegatus, an annelid that is used to feed pets and that lives in minerotrophic peatlands. Photo from Wikimedia Commons.

Prairie Worms It is possible that mosses may provide refugia for one rare species. The giant Palouse earthworm (Driloleirus americanus; Figure 12), named because it can reach nearly a meter in length, is the subject of a petition to declare it an endangered species and afford it protection (Palouse Prairie Foundation 2007). Few recent reports of its presence exist. In one such report, however, near Moscow, Idaho, USA, two researchers found it in a somewhat mesic area under forest canopy. The area had abundant mosses and these researchers found several of the worms near the surface

Figure 13. Lumbricus terrestris, the common earthworm, is able to transport various diaspores, thus being a potential dispersal agent for bryophytes. Photo by Michael Linnenbach through GNU Free Documentation.

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they can also be a nuisance. One person complained that the earthworms were the largest deterrent to the establishment of a moss garden. The worms would "plow" up the surface and detach the moss from the soil. It appeared that they also chewed up the moss, but there seems to be only circumstantial evidence of that.

Figure 14. Lumbricus terrestris wending its way in a clump of the moss Rhynchostegium confertum. Photo by Serhat Ursavas, with permission.

From a bryological point of view, it thus appears that the worms might serve as dispersal agents, although it was spores, not the more easily established tubers, that remained viable after traversing the earthworm gut (Van Tooren & During 1988). Tubers seemed unable to survive the journey through the gut. Twenty-five species of mosses germinated from diaspores from gut contents, with Pottia/Phascum (Figure 15) being the most common. This compares to the presence of only eight species of mosses in the samples of earthworms, indicating transport from other locations. For buried diaspores, earthworms may facilitate their movement from beneath the surface to the castings above ground where they are exposed to light and able to germinate. On the other hand, Bryum rubens (Figure 16) is not known to produce sporophytes in this area and relies on vegetative diaspores. It is one of the most common species in the area, but is not common above ground. It was also rare in the worm samples, causing Van Tooren and During to suggest that mechanical and chemical processes in the gut cause high mortality of the rhizoidal tubers in this species.

Figure 16. Clump of Bryum rubens, a moss that does not produce sporophytes and relies on dispersal of vegetative diaspores. Photo by Michael Lüth, with permission.

Polychaetes I completely overlooked this mostly marine group when I wrote this chapter (Figure 17). It was only when two people posted pictures on Bryonet of strange organisms they found among bryophytes that I realized there are terrestrial polychaetes that may inhabit bryophytes. These Bryonet organisms were not polychaetes, but they did raise the question. However, I have been unable to find any published documentation that polychaetes ever occur on bryophytes.

Figure 17. Syllid polychaete undergoing epitoky – becoming sexually mature. Photo by Megan McCuller, through Creative Commons. Figure 15. Pottia bryoides, a member of one of the genera that had the highest germination in cultures from earthworm guts. Photo by Michael Lüth, with permission.

Earthworm Culture Peatmoss is recommended as an additive to rich soil for rearing earthworms (Mascio 2006; How to Grow Your Own Earthworms 2009; Oliver 2009) Most farmers seem to consider earthworms to be their friends because they reputedly aerate the soil. However,

Storch and Welsch (1972) described adaptations to air breathing in polychaetes from the mangrove swamps of Sumatra. Their exterior is protected by a cuticle that varies in thickness. The gills have extracellular spaces that have blood lacunae in the epidermis in at least one species. But the terrestrial polychaetes seem to be poorly known. Thank you to Bryonet and its wonderful subscribers! Parergodrilus heideri and Hrabeiella periglandulata are the only terrestrial European flatworms, where they live in forest soils (Dumnicka & Rozen 2002) and would seem to be likely candidates for bryophyte dwelling (Juan Larrain,

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pers. comm. 29 February 2012). But both Larrain and I searched the web for links to bryophytes to no avail. Rather, Schlaghamerský and Šídová (2009) examined the vertical distribution of a population in the Czech Republic of Hrabeiella periglandulata in soil and determined that they avoided the organic layer, which would include bryophytes. Perhaps the minute Parergodrilus heideri (Rota 1997) and Hrabeiella periglandulata (Rota 1998) are hiding among them somewhere with the right moisture conditions. But it is more likely that the temperature of their environment is modified by the presence of bryophytes at the surface.

Summary Many bryophyte-inhabiting annelids (segmented worms) are mesofauna, i.e. able to occupy spaces with a diameter < 2 mm. The Enchytraeidae are among the most common. Bryophyte-dwelling annelids may form zones in the soil and bryophytes and some species may migrate up and down daily in response to changing moisture conditions. Enchytraeids have a wide tolerance to water, but have little adaptation to drought. Some species produce red blood to survive low oxygen conditions. Although most Enchytraeidae cannot reproduce by fragmentation, some enchytraeids can reproduce by this method in a cycle of ~20.4 days. Cognettia sphagnetorum increases its reproductive rate when temperatures get warmer, but an annual mean above 16ºC could cause annihilation. Some species thrive in higher temperatures, whereas others are seriously affected. Neither mosses nor fungi seem to serve as food for the annelids, although Sphagnum stems have been found in guts. In peatlands, 5-10-year old sediments seem to be an important food source. Bryophytes in streams can provide safe sites where reduced current provides more debris for food. Despite their apparent distaste for bryophytes, annelids may disperse vegetative diaspores by eating them and depositing them elsewhere unharmed, indicating at least some are not digested.. Worms can be extracted from bryophyte samples using funnel systems. Smaller taxa can be extracted by squeezing water onto a microscope slide. The Palouse earthworm (Driloleirus americanus) is a rare species that occurs under moss mats in the prairie.

Acknowledgments María Jesús Iglesias Briones provided invaluable help in obtaining images, literature, and a critique of an early draft on the annelids. Bryonetters have been wonderful in making their photographs available to me and seeking photographs from others.

Literature Cited Andrew, N. and Rodgerson, L. 1999. Practical conservation. Extracting invertebrates from bryophytes. J. Insect Conserv. 3(1): 53-55. Block, W. and Christensen, B. 1985. Terrestrial Enchytraeidae from South Georgia and the maritime Antarctic. Brit. Antarct. Surv. Bull. 69: 65-70. Briones, M. J. I. 2006. Enchytraeidae. In: Lal, R. (ed.). Encyclopedia of Soil Science, 2nd ed., Vol. 1. CRC Press, pp. 514-518. Briones, M. J. I., Ineson, P., and Heinemeyer, A. 2007. Predicting potential impacts of climate change on the geographical distribution of enchytraeids: a meta-analysis approach. Global Change Biol. 13: 2252-2269. Briones, M. J. I., Ineson, P., and Piearce, T. G. 1997. Effects of climate change on soil fauna; responses to enchytraeids, Diptera larvae and tardigrades in a transplant experiment. Appl. Soil Ecol. 6: 117-134. Brusven, M. A., Meehan, W. R., and Biggam, R. C. 1990. The role of aquatic moss on the community composition and drift of fish-food organisms. Hydrobiologia 196: 39-50. Christensen, B. 1959. Asexual reproduction in the Enchytraeidae (Olig.). Nature 184: 1159-1160. Didden, W. A. M. and Fluiter, R. de. 1998. Dynamics and stratification of Enchytraeidae in the organic layer of a Scots pine forest. Biol. Fertil. Soils 26: 305-312. Didden, W. A M., Fründ, H.-C., and Graefe, U. 1997. Enchytraeids. In: Benckiser, G. (ed.). Fauna in Soil Ecosystems. Recycling Processes, Nutrient Fluxes, and Agricultural Production. CRC Press, pp. 135-172. Duinen, G. A., Timm, T., Smolders, A. J. P., Brock, A. M. T., Verberk, W. C. E. P., and Esselink, H. 2006. Differential response of aquatic oligochaete species to increased nutrient availability - a comparative study between Estonian and Dutch raised bogs. In: Verdonschot, P. F. M., Wang, H., Pinder, A., and Nijboer, R. (eds.). Developments in Hydrobiology 186. Aquatic Oligochaete Biology IX. Selected papers from the 9th Symposium on Aquatic Oligochaeta, 6-10 October 2003, convened at the Conference Centre De Wageningse Berg, Wageningen, The Netherlands. Hydrobiologia 564: 143-155. Dumnicka, E. and Rozen, A. 2002. The first record of the terrestrial polychaete Hrabeiella periglandulata Pil et Chalupsk, 1984, in Poland, with a note on anatomy and ecology. Fragmenta Faunistica 45: 1-7. During, H. A., Brugues, M., Cros, R. M. and Lloret, F. 1987. The diaspore bank of bryophytes and ferns in the soil in some contrasting habitats around Barcelona (Spain). Lindbergia 13: 137-149. Egglishaw, H. J. 1969. The distribution of benthic invertebrates on substrata in fast flowing streams. J. Anim. Ecol. 38: 1933. 2160 Glime, J. M. 1968. Aquatic Insect Communities Among Appalachian Stream Bryophytes. Ph.D. Dissertation, Michigan State University, East Lansing, MI. Gorni, G. R. and da Gama Alves, R. 2007. Naididae (Annelida, Oligochaeta) associated with briophytes (sic) in Brotas, State of São Paulo, Brazil. Rev. Bras. Zool. 24 . Graefe, U. and Beylich, A. 2003. Critical values of soil acidification for annelid species and the decomposer community. Newslett. Enchytraeidae 8: 51–55.

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Habdija, I., Habdija, B. P., Matonickin, R., Kuciníc, M., Radanovíc, I., Miliša, M., and Mihaljevíc, Z. 2004. Current velocity and food supply as factors affecting the composition of macroinvertebrates in bryophyte habitats in karst running water. Biologia, Bratislava 59: 577-593. Healy, B. 1987. The depth distribution of Oligochaeta in an Irish quaking marsh. Hydrobiologia 155: 235-247. Healy, B. and Bolger, T. 1984. The occurrence of species of semi-aquatic Enchytraeidae (Oligochaeta) in Ireland. Hydrobiologia 115: 159-170. Hingley, M. 1993. Microscopic Life in Sphagnum. Illustrated by Hayward, P. and Herrett, D. Naturalists' Handbook 20. [iiv]. Richmond Publishing Co. Ltd., Slough, England, 64 pp.. 58 fig. 8 pl. (unpaginated). Honda, M., Suzuki, T., Matsumoto, S., and Gamou, S. 2003. Segment formation of Enchytraeus japonensis (Oligochaeta: Enchytraeidae): The 7th international symposium on earthworm ecology, Cardiff, Wales, 2002. Pedobiologia 47: 522-525. How to Grow Your Own Earthworms. 2009. Accessed on 12 March 2009 at . Hynes, H. B. N. 1961. The invertebrate fauna of a Welsh mountain stream. Arch. Hydrobiol. 57: 344-388. Linhart, J., Uvíra, V., and Vlčková, Š. 2002a. Permanent and temporary meiofauna of an aquatic moss Fontinalis antipyretica Hedw. Acta Univers. Palack. Olom. Biol. 3940: 131-140. Linhart, J., Vlčková, Š., and Uvíra, V. 2002b. Bryophytes as a special mesohabitat for meiofauna in a rip-rapped channel. River Res. Appls. 18: 321-330. Macan, T. T. 1966. Freshwater Ecology. Wiley & Sons Inc., N. Y. Mascio, Mike. 2006. Square Foot/Intensive Gardening – earthworms. Accessed on 12 March at . Myohara, M., Yoshida-Noro, C., Kobari, F. and Tochinai, S. 1999. Fragmenting oligochaete Enchytraeus japonensis: A new material for regeneration study. Development Growth Differentiation 41: 549-555. Nakamura, Y. 2004. The relation of fragmentation frequency to fragment number in Enchytraeus japonensis Nakamura, 1993 (Oligochaeta, Enchytraeidae) cultured several years under laboratory conditions. Mem. Fac. Agr. Ehime Univ. 49: 1926. O'Connor, F. B. 1955. Extraction of enchytraeid worms from a coniferous forest soil. Nature 175: 815-816. Oliver, George S. 2009. Friend Earthworm: Lesson 4. Housing the Earthworm Stock. Accessed on 12 March 2009 at . Overgaard-Nielsen, C. 1948. Studies on the soil microfauna. I. The moss inhabiting nematodes and rotifers. Naturvidenskabelige Skrifter Laerde Selsk Skrifter, Äarhus 1948(1): 1-98.

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Overgaard-Nielsen, C. 1949. Studies on the microfauna. II. The soil inhabiting nematodes. Natura Jutlandica 2: 1-132. Palouse Prairie Foundation. 2007. Giant Palouse Earthworm (Driloleirus americanus). Accessed on 12 March 2009 at . Percival, E. and Whitehead, H. 1929. A quantitative study of the fauna of some types of stream-bed. J. Ecol. 17: 283-314. Rota, E. 1997. First Italian record of the terrestrial polychaete Parergodrilus heideri Reisinger, with anatomical and ecological notes. Italian J. Zool. 64: 91-96. Rota, E. 1998. Morphology and adaptations of Parergodrilus Reisinger and Hrabeiella Pizl & Chalupský, two enigmatic soil-dwelling annelids. Italian J. Zool. 65: 75-84. Ryan, J. K. 1977. Synthesis of energy flows and population dynamics of Truelove Lowland invertebrates. In: Bliss, L.C. (ed.). Truelove Lowland, Devon Island, Canada: A High Arctic Ecosystem. University of Alberta Press, pp. 325-346. Schlaghamerský, J. and Šídová, A. 2009. Dynamics and vertical distribution of a Hrabeiella periglandulata (Annelida) population in South Moravia, Czech Republic. Pesquisa Agropecuária Brasileira 44: 917-921. Silvan, N., Laiho, R., and Vasander, H. 2000. Changes in mesofauna abundance in peat soils drained for forestry. Forest Ecol. Mgmt. 133: 127-133. Springett, J. A. 1970. The distribution and life histories of some moorland Enchytraeidae (Oligochaeta). J. Anim. Ecol. 39: 725-737. Springett, J. A. and Latter, P. M. 1977. Studies on the microfauna of blanket bog with particular reference to Enchytraeidae. 1. Field and laboratory tests of microorganisms as food. J. Anim. Ecol. 46: 959-974. Springett, J. A., Brittain, J. E., and Springett, B. P. 1970. Vertical movement of Enchytraeidae (Oligochaeta) in moorland soils. Oikos 21: 16-21. Standen, V. and Latter, P. M. 1977. Distribution of a population of Cognettia sphagnetorum (Enchytraeidae) in relation to micro-habitats in a blanket bog. J. Anim. Ecol. 46: 213-229. Storch, V. and Welsch, U. 1972. Ultrastructure and histochemistry of the integument of air-breathing polychaetes from mangrove swamps of Sumatra. Marine Biol. 17: 137-144. Suren, A. M. 1993. Bryophytes and associated invertebrates in first-order alpine streams of Arthur's Pass, New Zealand. N. Z. J. Marine Freshwat. Res. 27: 479-494. Thienemann, A. 1912. Der Bergbach des Sauerlandes. Internat. Revue D. Ges. Hydrobiol. Hydrographie. Biol. Suppl. 4. Ser. pp. 22-71. Tooren, B. F. van and During, H. J. 1988. Viable plant diaspores in the guts of earthworms. Acta Bot. Neerl. 37: 181-185. Vlčková, S., Linhart, J., and Uvíra, V. 2001/2002. Permanent and temporary meiofauna of an aquatic moss Fontinalis antipyretica Hedw. Acta Universitatis Palackianae Olomucensis 39-40: 131-140. Willard, J. R. 1972. Soil invertebrates: I. Methods of sampling and extraction. Canadian IBP Technical Rept. 7: 1-40.

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Glime, J. M. 2017. Invertebrates: Rotifers. Chapt. 4-5. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. Ebook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 18 July 2020 and available at .

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CHAPTER 4-5 INVERTEBRATES: ROTIFERS TABLE OF CONTENTS Rotifera – Rotifers................................................................................................................................................................... 4-5-2 Reproduction........................................................................................................................................................................... 4-5-2 Bdelloidea ....................................................................................................................................................................... 4-5-4 Monogononta .................................................................................................................................................................. 4-5-5 Bryophytes as Habitat ............................................................................................................................................................. 4-5-6 Habitat Characteristics .................................................................................................................................................... 4-5-6 Abundance ...................................................................................................................................................................... 4-5-9 Sampling .............................................................................................................................................................................. 4-5-10 Extraction Techniques .......................................................................................................................................................... 4-5-11 Adaptations ........................................................................................................................................................................... 4-5-11 Particle Feeders ............................................................................................................................................................. 4-5-14 Spines ............................................................................................................................................................................ 4-5-14 Small Size ..................................................................................................................................................................... 4-5-14 Mobility vs Attachment? ............................................................................................................................................... 4-5-15 Protection ...................................................................................................................................................................... 4-5-15 Dormant States .............................................................................................................................................................. 4-5-17 Physiological Adaptations..................................................................................................................................................... 4-5-17 Anhydrobiosis ............................................................................................................................................................... 4-5-17 Changes During Anhydrobiosis ............................................................................................................................ 4-5-17 Longevity during Anhydrobiosis........................................................................................................................... 4-5-19 Age Differences .................................................................................................................................................... 4-5-20 Size Differences – Aquatic vs Terrestrial .............................................................................................................. 4-5-20 Reproductive Effects ............................................................................................................................................. 4-5-20 Temperature Protection ......................................................................................................................................... 4-5-20 Recovery Rate ....................................................................................................................................................... 4-5-20 The Bryophyte Connection ................................................................................................................................... 4-5-20 Other Protections during Anhydrobiosis ............................................................................................................... 4-5-21 Surviving Fungi ............................................................................................................................................................ 4-5-21 Food ...................................................................................................................................................................................... 4-5-21 Role in the Food Web ........................................................................................................................................................... 4-5-24 Specific Habitats ................................................................................................................................................................... 4-5-25 Lobule Dwellers ............................................................................................................................................................ 4-5-25 Retort Cells ................................................................................................................................................................... 4-5-28 Roofs ............................................................................................................................................................................. 4-5-28 Arctic and High Arctic .................................................................................................................................................. 4-5-30 Antarctic........................................................................................................................................................................ 4-5-31 Nunataks ............................................................................................................................................................... 4-5-34 Bog and Fen Habitats .................................................................................................................................................... 4-5-35 Species Richness ................................................................................................................................................... 4-5-36 Abiotic Factors ...................................................................................................................................................... 4-5-40 Acidity .................................................................................................................................................................. 4-5-40 Surface Configuration ........................................................................................................................................... 4-5-41 Pitcher Plants ........................................................................................................................................................ 4-5-43 Aquatic Bryophytes....................................................................................................................................................... 4-5-43 Streams.................................................................................................................................................................. 4-5-43 Waterfalls .............................................................................................................................................................. 4-5-46 Krakatau ........................................................................................................................................................................ 4-5-47 Seasons ................................................................................................................................................................................. 4-5-48 Danger amidst the Bryophytes .............................................................................................................................................. 4-5-48 Ozone Hole and Pollution Dangers? ..................................................................................................................................... 4-5-50 Summary .............................................................................................................................................................................. 4-5-51 Acknowledgments ................................................................................................................................................................ 4-5-51 Literature Cited ..................................................................................................................................................................... 4-5-52

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Chapter 4-5: Invertebrates: Rotifers

CHAPTER 4-5 INVERTEBRATES: ROTIFERS

Figure 1. Two bdelloid rotifers that commonly inhabit bryophytes. Photo by Paul Davison, with permission.

Rotifera – Rotifers Rotifers, also known as wheel animals, are so-named because of the ciliated corona on the head. The corona creates a circular movement that is used to direct food to the mouth. Rotifers have up to five simple eyes (Figure 2) that are light-sensitive and often are red. This sensitivity to light permits some species to be phototactic (moving toward or away from light). Rotifers are natural partners for organisms like bryophytes that often experience extended periods of drought. Pourriot (1979) considered the number of species that inhabit mosses to be over 200. The number is surely larger now. Anthony von Leeuwenhoek discovered in 1702 that rotifers could tolerate months in a state of desiccation, hence marking the earliest studies on cryptobiosis, or life in a dormant state without water (Alpert 2000). This desiccation tolerance is particularly common in the class Bdelloidea. In this dry state, they are easily dispersed along with fragments of the mosses they inhabit. Not much bigger than some protozoa (mostly 0.10.5 mm long, but up to 2 mm), they form a phylum of their own, the Rotifera, with at least 2000 species (Howey

1999). They are multicellular and even possess a primitive brain, at least in females (Hingley 1993).

Figure 2. Brachionus quadridentatus (Monogononta) showing red eyespot. Photo by Frank Fox, through Creative Commons.

Chapter 4-5: Invertebrates: Rotifers

Rotifers have a variety of means of protection. Some are encased in a lorica (rigid case or shell; Figure 3, Figure 13-Figure 14). Others build tubes or cases (Figure 53, Figure 82). Some have sharp spines (Figure 13). And some simply hide, many of which use bryophytes for hiding.

Figure 3. Colurella adriatica, showing location of the mastax and other prominent features. This one is sitting on the green alga Spirogyra sp., but it sometimes occurs among mosses. Photo by Michel Verolet, with permission.

Moss-dwelling rotifers have been around for a long time. Waggoner and Poinar (1993) reported on fossil habrotrochid rotifers from Dominican amber. These revealed microfossils from the bracts of a moss from the Eocene-Oligocene (circa 34 million years ago) in the northern Dominican Republic. It is interesting that these match the thecae (sheaths) of living moss dwellers in Habrotrocha, being almost identical with H. angusticollis (Figure 4). These parthenogenetic (producing unfertilized eggs) bdelloid rotifers seem to have a well-adapted body plan that has persisted for 35 million years.

Figure 4. Habrotrocha angusticollis, a moss inhabitant. Photo by Yuuji Tsukii, with permission.

It is likely that many species of rotifers remain to be described. The most likely habitat for these discoveries is

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that of bryophytes. The bryophyte dwellers are often very small, rarely swim, and go dormant (see below) as a tun (Figure 61) or a resting egg, all characteristics that make them less likely to be noticed and more difficult to identify. Shiel and Green (1996) remarked that considerably more rotifers in New Zealand and the Australasian region remain undescribed. At that time the region had 388 valid species in 66 genera. Yet less than 5% of these were endemic to the Australasian region. With the potential differences in physiology and biochemistry, it is also likely that DNA analysis will reveal many microspecies and perhaps even different species that are not recognizable based on morphology alone. Kaya et al. (2009) compared "DNA species" with morphological species of bdelloid rotifers from mosses in Turkey and the United Kingdom. They found that traditional identification methods underestimate rotifer diversity by factors of 2 at the local level and 2.5 at a regional level. Each moss sample had 3-9 morphospecies, but the DNA species ranged 8-12 per moss sample. These DNA species numbers indicated greater differences in diversity among locations (gamma diversity) than within samples (alpha diversity). Rotifer biologists consider that the number of cryptic species that can be revealed by DNA taxonomy may be overwhelming (Suatoni et al. 2006; Fontaneto et al. 2008). This knowledge that the Rotifera include many cryptic species (species that look alike but can't interbreed), as demonstrated by DNA, is supported by a diversity of narrow ecological niches (see, for example, Fontaneto et al. 2011). This allows for physiological/biochemical differences that permit the species to survive in a wide range of cosmopolitan habitats. This diversity and cosmopolitan distribution has led to superfluous names in many of the rotifer genera. This chapter follows the nomenclature of Segers (2007); for species described after that publication it follows EOL .

Reproduction The lifespan of many rotifers is as much as 30-40 days, not counting their time in dormant states (Ricci 2001). But Wikipedia (2016) considers it to be much shorter for Monogononta, ranging 2 days to 3 weeks for females. And species of these animals can often be found in active or dormant states on both aquatic/wetland (Priddle & Dartnall 1978; Bateman & Davis 1980; Ricci 1983; Ricci et al. 1989; Linhart et al. 2002a) and terrestrial mosses (Bartos 1949; Ramazotti 1958; Overgaard-Nielsen 1967; Kukhta et al. 1990). Several species are even known from the harsh environment of mosses growing on roofs (Hirschfelder et al. 1993). Rotifers (depending on the taxon) have three types of individuals: mictic (mixing) females, amictic females (not reproducing sexually), and males. Rotifer eggs may be attached to a substrate (Figure 5-Figure 6) or remain attached to the parent (Figure 7) (EOL 2016). The female rotifers themselves live only a few days to a few weeks. The males have no digestive tract, are often sexually mature at birth, and are short-lived, as you might expect when they don't eat. Hence, it is also understandable that males are much smaller than females (Figure 8).

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Chapter 4-5: Invertebrates: Rotifers

Figure 5. Bdelloid rotifer eggs on alga. Photo by Michel Verolet, with permission.

Figure 8. Cephalodella gibba in copulation, male on left. Photo by Michael Plewka , with permission.

Figure 6. Egg of rotifer on an algal filament. Photo by Michel Verolet, with permission.

Figure 9. Asplanchna girodi vitellarium. Photo by Michael Plewka , with permission.

Bdelloidea

Figure 7. Brachionus with 3 eggs. Photo by Jean-Marie Cavanihac, with permission.

The female reproductive system of rotifers consists of one (Monogononta) or two (Bdelloidea) ovaries. Each ovary has a vitellarium gland (Figure 9) that supplies the eggs with yolk.

Bdelloid rotifers (class Bdelloidea; Figure 10-Figure 11), known as moss rotifers, are less species rich (over 450 described species) than the Monogononta (ca 1500 species). The Bdelloidea are the most common rotifers in peatlands (bogs and fens; Bielańska-Grajner et al. 2011) and other mosses (Sayre & Brunson 1971; Ricci et al. 2003b; Gilbert & Mitchell 2006). All known taxa are parthenogenetic, i.e., they have only females that reproduce asexually, giving rise to more females (Hingley 1993). However, Danchin et al. (2011) analyzed the genome of one of these, Adineta vaga (Figure 12), a moss dweller, and found four genotype modifications that suggested rare events of sexual reproduction may have occurred.

Chapter 4-5: Invertebrates: Rotifers

Figure 10. Bdelloid rotifer taken from bryophytes. Photo courtesy of Dan Spitale.

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Brachionus (Wikipedia 2011; Figure 2, Figure 7, Figure 13-Figure 14). In this genus, with some members occurring among bryophytes, increases in population density can induce sexual reproduction. The sexually produced eggs can become resting eggs that survive unfavorable conditions (Plewka 2014). It appears that at least in Brachionus calyciflorus (Figure 13) only one allele is needed to turn off sexual reproduction and force all reproduction to be parthenogenetic. Brachionus urceolaris (Figure 14) sometimes lives among bryophytes (Figure 7; Hingley 1993), but it is primarily a cosmopolitan planktonic species like the other Brachionus species (EOL 2016). It is mostly parthenogenetic, but it occasionally produces males.

Figure 11. Examples of bdelloid rotifers and trophi, the hardened part of the mastax. Photos by Diego Fontaneto, through Creative Commons Figure 13. Brachionus calyciflorus, a species that needs only one allele to turn off sexual reproduction. Academy of Natural Sciences in Philadelphia, through Creative Commons.

Figure 12. Adineta vaga, a moss dweller that is 0.2-0.3 mm when extended. Photo by Michael Plewka , with permission.

Monogononta The Monogononta is the second major class of rotifers, and by far the largest (ca 1500 species) (Wikipedia 2012a). Among these are members that have both sexual and asexual reproduction. The short-lived, uncommon males, however, serve only for reproduction and thus are much smaller than females. Some males are so reduced that they have little more than a bladder and a penis! One such monogonont is the mostly planktonic genus

Figure 14. Brachionus urceolaris, a bryophyte dweller. Photo courtesy of Emily Toscana Guerra from Rotifer World Catalog, through Creative Commons.

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Chapter 4-5: Invertebrates: Rotifers

In the Monogononta, two types of reproduction occur. In one type, females produce unfertilized eggs that develop into females, just as in the bdelloids (Hingley 1993). But in the second type, sexual females appear only when environmental conditions are unfavorable, such as drought or cold. These females produce a sexual egg that forms a thick-walled resting "egg" when fertilized (Figure 15). That resting egg develops into a female. If the egg is not fertilized, it develops into a male.

(drifting in open water) species. Bryophytes are among these macrophytic (referring to plants that are visible without a microscope) substrates that support the periphyton, but Duggan did not include them in his study, considering bryophytes to be a separate habitat. Periphytic rotifers seem to have preferences among macrophyte species based on differences in physical structure or complexity, food concentration or composition, chemical factors, macrophyte age, and differences in protection from predation they provide (Duggan 2001). The same factors are likely to control bryophyte choices as well. Terrestrial and wetland rotifers crawl through the spaces among leaves and branches of bryophytes, living in the water film surrounding the plant (Hingley 1993). In her website on rotifers, Jean-Marie Cavanihac (2016) considers Rotaria rotatoria (formerly Rotifer vulgaris; Figure 17) to be one of the most frequent rotifers on mosses, and as a free-living (unattached) rotifer, it moves like a caterpillar.

Figure 15. Euchlanis triquetra with expelled resting egg. Photo by Michael Plewka , with permission.

Bryophytes as Habitat Moss-dwelling rotifers have attracted the attention of rotifer specialists for some time (Burger 1948). The family Habrotrochidae (see Lobule Dwellers below) seems to occur mostly on mosses but is also benthic (living on the bottom of a water body) (Wallace & Snell 1991). There are two species in the genus Elosa (Figure 16) that are common on Sphagnum (Figure 25-Figure 27, Figure 109Figure 112), and these are considered bog specialists (Pejler & Bērziņš 1993b).

Figure 16. Elosa worrallii, a Sphagnum dweller. Photo by Jersabek et al. 2003 from Rotifer World Catalog, through Creative Commons.

Rotifers occur with bryophytes in both aquatic and terrestrial habitats, with bryophytes often providing a water space in the latter. Duggan (2001) points out that the periphytic (living on plant surfaces) species of rotifers have received little attention compared to the planktonic

Figure 17. Rotaria rotatoria, a bdelloid rotifer from moss. Photo by Christian D. Jersabek, through Creative Commons.

The bryophyte dwellers feed on the bacterial and protozoan inhabitants, swim among the leaves, or nestle between the leaves and branches where they gain more protection against their predators (Hingley 1993). The same is true for those living in terrestrial habitats as well as in ponds, lakes, and waterways. Habitat Characteristics Although not restricted to these habitats, rotifers are common on mosses in alpine Sphagnum (Figure 25-Figure 27, Figure 109-Figure 112) bogs and in wetlands. Bryophytes may be particularly useful to stream and other aquatic rotifers as a substrate. Pejler and Bērziņš (1989) contend that rather than any chemical attraction for a substrate, some substrates might be avoided, perhaps due to lack of periphyton. The genus Lecane (Figure 122) is a very large, widespread genus that has little preference for any particular substrate (Pejler & Bērziņš 1994). In fact, it furthermore seems to have good dispersal, as indicated by its rapid ease of colonization on an artificial substrate of cotton. Fontaneto and Ricci (2006) consider that rotifers are probably best dispersed in their dormant state (allowing them to be dispersed along with their bryophytic substrate). The species on various macrophytes differ, even when a different species of macrophyte is growing in close proximity (Pontin & Shiel 1995; Duggin et al. 2001). Likewise, bryophyte species composition explains most of the variation in monogonont rotifers in springs and fens

Chapter 4-5: Invertebrates: Rotifers

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(Hájková et al. 2011). Bryophytes form four functional groups, supporting the importance of plant form in their selection of the bryophyte substrate. Species composition of monogonont rotifers differs significantly (P moss 0%:corn 100% > moss 100%:corn 0% > moss 75%:corn 25%. The highest weight gain of 15 g peaked at the fourth day in L. aurora fed with moss 50%:corn 50%. Furthermore, the snails exhibited a strong positive weight gain correlation with increasing days of feeding with 25% moss to 75% corn.

Figure 50. Hyophila involuta, with a snail; the snail Limicolaria aurora can thrive on this moss. Photo by Li Zhang, with permission.

Figure 49. Orthotrichum urnigerum, member of a genus known to be grazed by snails. Photo by Michael Lüth, with permission.

Low Nutritional Quality? That rasping tongue is not always enough to accomplish the task of obtaining nutrients from mosses. Oyesiku and Ogunkolade (2006) experimented with snails and the moss Hyophila involuta (Figure 50). In their laboratory experiments, the snails (Limicolaria aurora; Figure 51) gained the most weight when fed with Hyophila involuta paste. The snails that had only unground moss actually lost weight. Those in the field experiment (restricted to Hyophila involuta) either lost weight or remained the same. Fecal matter of the field snails had fragments of moss that had lost the chlorophyll from their cells as well as that of abundant algae and Cyanobacteria. The presence of these snails on the moss was seasonal from April until October, when the moisture and lower temperature of the moss may have provided a favorable habitat. This experiment suggests that in this case the snail was unable to penetrate the cells of the moss, making it an unlikely food source in nature. Rather, the researchers suggest that the snails most likely use the moss as a moist and cool habitat. Oyesiku and Bello (2012) experimented further with the effect of the moss Hyophila involuta (Figure 50) as a food for the snail Limicolaria aurora (Figure 51). The study was based on an interest in including mosses as feed when breeding snails. The moss was mixed in various ratios with corn pap powders (Zea mays). Overall, there was a significant correlation with the feed ratios of decreasing order of moss 50%:corn 50% > moss 25%:corn

Figure 51. Shell of Limicolaria aurora. Photo by David G. Robinson, USDA APHIS PPQ at Bugwood.org, through public domain.

Chemical Deterrents to Herbivory Longton (pers. comm. 1996) has speculated that phenolic compounds that protect the leafy gametophytes deter herbivory, especially on perennials. This could account for greater herbivory on the annual Funaria hygrometrica (Figure 52) than on perennial Brachythecium rutabulum (Figure 66) or Mnium hornum (Figure 80). The phenolic compounds in the latter two species were released only after severe hydrolysis, leading Davidson et al. (1990) to suspect that the phenolic acids might be tightly bound to cellulose in the cell wall. The greater palatability of the F. hygrometrica supports the general theory that perennials invest more resources in defense against herbivory than do annuals such as F. hygrometrica.

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Chapter 4-8: Invertebrates: Molluscs

Figure 52. Young sporophytes of Funaria hygrometrica before spores form. Photo by Michael Lüth, with permission.

Food for Some

Algae growing on mosses, especially in the aquatic habitat, could be a prominent source of food for gastropods. In the Negev Desert, adult desert snails (Sphincterochila zonata) fed exclusively on algae on the soil surface, creating an algal turnover of 142 kg hectare-1, despite being active for only 8-27 days in winter during the rainy period (Shachak & Steinberger 1980). Other Negev Desert snails feed on the mosses themselves. Sphincterochila boissieri (Figure 55) feeds on shrubs there, but its feces indicate that it also feeds on the moss Tortula atrovirens (=Desmatodon convolutus; Figure 56) (Yom-Tov & Galun 1971). This is a snail that has color morphs of brown and white, but they apparently don't affect its temperature (Yom-Tov 1971; Slottow et al. 1993). However, their rodent predators choose more brown than white snails, enough to exhibit significant differences in their choices (Slottow et al. 1993).

Clearly for some slugs and snails there are bryophytes that do indeed seem palatable. Ochi (1960) reported that the thallose liverwort Conocephalum conicum (Figure 53) served as food for a slug. Merrifield (2000) found evidence of heavy grazing on epiphytic bryophytes, particularly the moss Syntrichia laevipila (Figure 54), of Oregon white oaks (Quercus garryana) in the Willamette Valley, Oregon, USA, and considered that either springtails or slugs were likely responsible. She considered that the abundance of gemmae on S. laevipila may be a response to this grazing. Figure 55. Sphincterochila boissieri, a species that is known to eat Tortula atrovirens in the Negev desert. Photo by Mark A. Wilson, through Creative Commons.

Figure 53. Conocephalum conicum showing feeding damage upper middle) by something, perhaps a slug. Photo by John Hribljan, with permission.

Figure 56. Tortula atrovirens, a moss that is eaten by the Negev Desert snail, Trochoidea seetzeni. Photo by Des Callaghan, with permission.

Figure 54. Syntrichia laevipila on bark. Photo by Jonathan Sleath, with permission.

Szlavecz (1986) examined feeding preferences in 31 individuals of the snail Monadenia hillebrandi mariposa (Figure 27). Collections of field feces indicated that they consumed the mosses Rhytidiadelphus sp. (Figure 57) and Grimmia trichophylla (Figure 58) in nature, among other things. In the lab, they preferred shrub and bay litter over mosses, but preferred mosses and lichens over grasses and pine litter. More green moss than brown occurred in the feces, whereas brown material was more common from consumed tracheophytes (Figure 59).

Chapter 4-8: Invertebrates: Molluscs

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Grime and Blythe (1969) found bryophytes in the feces of four species of snails out of the six examined from Winnats Pass, Derbyshire, England, on 13 October. But then, tracheophyte foods are often less nutritious as the plants prepare for winter. Studies by Chatfield (1973), Williamson & Cameron (1976), and Richter (1976) indicate that at least juvenile snails might do best on a mixed diet. But for Cepaea nemoralis (Figure 60-Figure 61), it appears that even though mosses are part of their habitat, they are seldom part of the diet (Williamson & Cameron 1976).

Figure 57. Rhytidiadelphus squarrosus, a member of a genus that has been found in feces of the snail Monadenia hillebrandi mariposa. Photo by Michael Lüth, with permission.

Figure 60. Cepaea nemoralis, banded snail juvenile at Old Sulehay Forest, UK, a species that lives in a mossy habitat but apparently does not eat them. Photo by Brian Eversham, with permission. Figure 58. Grimmia trichophylla showing awns. Photo by Michael Lüth, with permission.

Figure 61. Cepaea nemoralis, a species that lives in a mossy habitat but apparently does not eat them. Photo by Stefan Haller, with permission.

Figure 59. Comparison of green and brown portions of plant material eaten by the snail Monadenia hillebrandi mariposa. Modified from Szlavecz 1986.

In the tropical montane rainforest of Brazil, those small, flattened snails in the Charopidae (Figure 62) eat bryophytes (Maciel-Silva & dos Santos 2011). Both Canalohypopterygium tamariscinum (syn. = Hypopterygium tamarisci; Figure 63) and Lopidium concinnum (Figure 64) had evidence of leaf herbivory, mostly in the beginning of the rainy season (September to December). A species of snail in the Charopidae and a moth larva in the Geometridae were the culprits. Using an index of damage (ID) in 2007, 2008, Maciel-Silva and dos Santos found that C. tamariscinum had higher damage (68%, 35%) than L. concinnum (38%, 23%) in these two years (Figure 65). These rates were lower than those for

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Chapter 4-8: Invertebrates: Molluscs

tracheophytes. They found no correlation with phenols, proteins, or the ratio between them (Figure 65).

Figure 65. Charopidae and Geometridae damage to mosses in 10 colonies of plants. Image from Adaises MacielSilva and Nivea Dias dos Santos.

An Avoidance of Gametophores? Figure 62. Charopidae feeding on Lopidium concinnum from an Atlantic Forest, Brazil. Photo by Adaises Maciel-Silva and Nivea Dias dos Santos, with permission.

Figure 63. Canalohypopterygium tamariscinum, a food source for Charopidae. Photo by Niels Klazenga, with permission.

Figure 64. Evidence of Charopidae herbivory on Lopidium concinnum from an Atlantic Forest, Brazil. Photo by Adaises Maciel-Silva and Nivea Dias dos Santos, with permission.

Davidson and Longton (1985, 1987; Davidson 1988, 1989) reported that several species of generalist slugs consumed bryophytes. In some cases, the protonema (threadlike stage that develops from moss spore) is readily consumed (Grime 1979). In Great Britain, capsules and protonemata of several mosses [Brachythecium rutabulum (Figure 66), Mnium hornum (Figure 67-Figure 68), and Funaria hygrometrica (Figure 69)] were eaten preferentially to leafy gametophores by slug species in the genus Arion (Figure 70) (Davidson & Longton 1987; Davidson et al. 1990). Cambs (2012) found that the slug Limax maculatus (Figure 40) likewise would eat capsules, but the leafy parts seemed to serve only as an emergency food. It appears that some may even eat calyptrae (covering over capsule; Figure 71). Ferulic acid, present in shoots but absent in young capsules of Mnium hornum, is a phenolic compound that is only released after severe hydrolysis. Its antibiotic role as an antifungal agent (Sarma & Singh 2003) and in antiherbivory (Seigler 1983; Smith 2011) may contribute to this preference for capsules, as discussed below. Davidson and coworkers found that older capsules with spores were less preferred than the green ones (Figure 72; Davidson & Longton 1987; Davidson et al. 1990).

Figure 66. Slug eating capsules of Brachythecium. Note the number of setae that are missing capsules. Photo by Janice Glime.

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Figure 71. Slug on moss calyptra, apparently finding something to eat. Photo courtesy of Sarah Lloyd.. Figure 67. Young, green capsules of Mnium hornum that are preferred by Arion slugs. Photo by Michael Lüth, with permission.

Figure 68. Mature capsules of Mnium hornum. Photo by Janice Glime.

Figure 72. Relative damage by slugs (Arion spp.) of sporophyte stages of two species of bryophytes. n=300-500 at day 0. LCI = late calyptra stage; EOI = early operculum intact; LOI = late operculum intact; OF = operculum fallen; EF = empty and fresh. Redrawn from Davidson et al. 1990.

Davidson (1989) found that slugs consumed only trivial amounts of Brachythecium rutabulum shoots (Figure 66). Mnium hornum (Figure 80) was also ignored, but after 5-7 days of starvation Arion rufus (10-15 cm long; Figure 73) and A. subfuscus (5-7 cm long; Figure 74) ate significant quantities of shoots of this species. The garden slug Arion hortensis (Figure 75) still ignored the moss even after 7 days of starvation. Figure 69. Capsules of Funaria hygrometrica – potential snail food. Photo by Michael Lüth, with permission.

Figure 70. Arion rufus on mosses in a woodland above Poole's Cavern, Buxton, UK. Photo by Brian Eversham, with permission.

Figure 73. Arion rufus on a bed of mosses. Photo by Jean Bisetti, with permission.

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Chapter 4-8: Invertebrates: Molluscs

Figure 74. Arion subfuscus, a slug known to consume Mnium hornum. Photo by Gary Bernon, USDA APHIS at Bugwood.org, through public domain.

Figure 75. Arion hortensis s.s. at Bridge House, Swavesey, UK. Photo by Brian Eversham, with permission.

Given the choice of capsules or vegetative material, both Arion rufus (Figure 3, Figure 70, Figure 73) and A. subfuscus (Figure 76) preferred immature capsules (see Figure 77 with a slug on immature capsules of Leucolepis acanthoneuron) of all three mosses, with Mnium hornum (Figure 80) being top choice (Davidson 1989). Setae were generally ignored, but A. subfuscus did occasionally eat M. hornum and Brachythecium rutabulum (Figure 66) setae. All three slugs also ate protonemata in the laboratory, and for B. rutabulum and Funaria hygrometrica (Figure 52) the protonemata were eaten just as much by A. rufus and A. subfuscus as were immature capsules. In fact, dry weight consumption exceeded that of immature capsules. Young shoots were also eaten, but less readily.

Figure 76. Arion subfuscus, a slug that prefers immature capsules. Photo by Sanja 565658, through Creative Commons.

Figure 77. Slug browsing on immature capsule of the moss Leucolepis acanthoneuron. Photo from UBC website, with permission.

Davidson and Longton (1987) suggested that Arion hortensis (Figure 75) was restricted by the physical structure of the capsule to consuming developing spores from broken capsules in Polytrichum commune (Figure 78); no spores were eaten from unbroken capsules. When approaching Mnium hornum (Figure 80), the slugs would withdraw their tentacles, then retreat, suggesting some sort of chemical deterrent; they behaved similarly in the presence of extracts from the capsule. It is likely that hydroxycinnamic and phenolic acids in this species and in Brachythecium rutabulum (Figure 66) provided this chemical protection against herbivory (Davidson et al. 1989). Stems of both species were apparently protected by ferulic and possibly m- and p-coumaric acids bound in the cell walls of the shoots (Davidson et al. 1989), explaining the preference of the slugs for capsules. On the other hand, when moss extracts were placed on communion wafers, the slugs ate them more readily, suggesting that chemistry alone was not the likely deterrent (Anonymous 1987; Davidson et al. 1990). Rather, some physical feature of the mosses, perhaps the cell wall, deterred these slugs.

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of Buxbaumia viridis (Figure 84). They found three types of damage on the sporophytes. In one type the seta and lower part of the capsule remain. For this type, they actually observed slugs feeding on the capsules; the same kind of slug was also feeding on young green capsules of Herzogiella seligeri (Figure 85). In a second type, the entire capsule is gone, but the seta remains. This could have been slugs as well, but they were unable to observe them and considered that ants or birds might also feed on them. The capsules are grazed in spring before they mature, thus likely being unable to accomplish a successful dispersal. The third type was destruction by a fungus, causing abortion of the capsule development.

Figure 78. Polytrichum commune capsules showing the persistent hairy calyptra and waxy capsule that is only eaten by snails when the capsule is broken. Photo by Michael Lüth, with permission.

Presence of moss cells of Brachythecium rutabulum (Figure 79) and Mnium hornum (Figure 80-Figure 81) in the feces of previously starved Arion suggest that the leafy mosses are not digested well (Davidson et al. 1990). On the other hand, all three species of slugs named above readily consumed Funaria hygrometrica (0.4-6.5 mg wet weight per slug; Figure 69) in overnight feeding trials. The importance of mosses as food may rest with the organisms living on the mosses – fungi, bacteria, protozoa, rotifers, etc., making indigestibility of the mosses inconsequential.

Figure 80. Mnium hornum shoots – a species that was ignored in experiments until the slugs were starved. Photo by Janice Glime.

Figure 79. Brachythecium rutabulum cells as they might be seen in feces. Photo by Tom Thekathyil, with permission.

Mostly indirect evidence suggests that slugs and snails graze capsules of Buxbaumia viridis (Figure 84) (Gordon Rothero, Birds feeding on moss capsules, Bryonet-l, 10 April 2003; Figure 84). Michael Lüth (Bryonet 23 September 2017) observed and photographed a slug grazing on the capsule of Buxbaumia viridis (Figure 83). Dave Kofranek reports tasting it – to him it tastes like cucumbers (Bryonet 24 September 2017). Infante Sánchez and Heras Pérez (2015) exlored the herbivory on capsules

Figure 81. Mnium hornum leaf tip cells, what one might see in feces. Photo by Bob Klips, with permission.

It is perhaps not surprising that snails eat the capsules of Splachnum (Figure 82). This genus has odors that attract flies, so they may serve as attractants to gastropods as well. However, no studies have attempted to test this hypothesis with snails.

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Figure 82. Snail on setae of Splachnum capsules in Alaska, eating capsules. Photo courtesy of Blanka Shaw.

Figure 85. Herzogiella seligeri with capsules, a species in which young capsules are eaten by slugs. Photo by Hermann Schachner, through Creative Commons.

Guy Brassard reported to me that Stéphane Leclerc has taken a picture of a slug in Quebec, Canada, eating a Buxbaumia aphylla (Figure 86-Figure 88) capsule!

Figure 83. Buxbaumia viridis with slug eating capsule. Photo by Michael Lüth, with permission.

Figure 86. Buxbaumia aphylla that are immature and have not been eaten. Photo by Štĕpán Koval, with permission.

Figure 84. Buxbaumia viridis capsules. Note that the leafy part belongs to another species of moss. Photo by Adolf Ceska, with permission.

Stark (1860) relayed a story of the ill fate of collected specimens of Buxbaumia aphylla (bug-on-a-stick moss; Figure 87) on their journey from Scotland to England. A slug had inadvertently been included in the package and it managed to destroy their prized specimens. On the other hand, B. aphylla can fool you. After repeated observations with my graduate student, Chang-Liang Liao, we have discovered in the field that what appeared to me to be grazing on capsules of Buxbaumia aphylla is really only the splitting of the capsule top as it dries (Figure 87), and that this occurs on nearly every capsule.

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Digestibility

Figure 87. Buxbaumia aphylla showing exposed green spores in the capsule that has split open. Photo by Janice Glime.

So what did the slugs derive from the consumed mosses? When they consume preferred foods such as lettuce leaf or carrot root, the resulting feces contain macerated, partially pigmented tissue (Davidson 1989). When they consumed bryophytes, on the other hand, large pieces of leaf, whole leaves, and even stem pieces remained intact. Most cells still contained green chloroplasts. Evidently the moss did little more than fill the gut. Even the preferred capsules were poorly digested, with capsule wall fragments, opercula, and peristome teeth remaining. Mature spores seemed unharmed, but immature spores seemed to have experienced some digestion, appearing broken, colorless, and shrivelled. Likewise, the protonemata seemed to be digestible, resembling the lettuce and carrots in being macerated and colorless or brown. Caution must be used in conducting laboratory experiments with food choices. Jennings and Barkham (1975) found that bryophytes all gave low palatability scores when six species of slugs, including the three in the Davidson (1989) study, had a choice of foods. The wider range of choices in the field may permit them to avoid the less palatable bryophytes. Role in Bryophyte Competition with Lichens Rosso and McCune (2003) found that molluscs on shrubs in the Pacific Northwest, USA, exhibited significant herbivore activity on the lichens. Bryophytes, on the other hand, had little change in cover between stems in exclusions and those available for herbivory. It appears that the mollusc herbivory on lichens (Boch et al. 2011) may benefit the bryophytes by contributing to the successful competition of the bryophytes over the lichens in the understory of these forests. Palatable Gametophytes

Figure 88. Buxbaumia aphylla that may have been damaged by a herbivore. Photo by Janice Glime.

Slugs also eat hornworts (Anthocerotophyta; Figure 89). Bisang (1996) reported that they especially eat the green sporophytes.

Figure 89. Phaeoceros carolinianus, a hornwort with mostly green sporophytes, a food source for slugs. Photo by Michael Lüth, with permission.

Des Callaghan (Bryonet 10 June 2011) reports slugs feasting on the gametophytes of Hookeria lucens (Figure 90) near a stream. In only six days they completely removed all the plants by dining on them, leaving behind only a stump and a slime trail (Figure 91). This was a research station, so Callaghan needed to find a way to discourage the slugs. Suggestions from Bryonetters included sprinkling ground glass around the study area (Michael Richardson, Bryonet 10 June 2011); putting out cups of beer to attract and drown the slugs or putting curry powder or other hot substance around the mosses (Janice Glime, Bryonet 10 June 2011); copper rings that are effective in gardens and could be made with a coil of wire (David Bell, Bryonet 10 June 2011).

Figure 90. Hookeria lucens in healthy condition. Photo by Des Callaghan, with permission.

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Chapter 4-8: Invertebrates: Molluscs

Figure 91. Temperature/humidity data logger with Hookeria lucens eaten by slugs. Photo by Des Callaghan, with permission.

Annie Martin (Bryonet 11 June 2011) is a professional gardener and described her experience in trying to eliminate slugs. She suggested putting salt on the head (if put on the tail the slug continues to live and eat). Her experience with beer is that it just keeps on attracting snails night after night, even though many of them drown, so it is an ineffective waste of money. Brown mulch seems to provide a favorable habitat, so she eliminated it, a technique that worked, but isn't relevant for discouraging snails on mossy rocks. Aquatic Grazing Grazing by gastropods (slugs and snails) can be so severe as to define distribution of a bryophyte species. Lohammar (1954) found that in northern Europe Fissidens fontanus (Figure 92) was absent in lakes where Fontinalis antipyretica (Figure 93) was also absent. Gerson (1982) suggested that scarcity of Fissidens in some places is due to snail grazing. In the presence of Fontinalis, this smaller moss lives among the Fontinalis fronds where it is presumably protected from snail grazing by the inedible forest of Fontinalis surrounding it and the density of the Fontinalis stems.

Figure 93. Fontinalis antipyretica, a moss that apparently protects the smaller Fissidens from grazing by snails. Photo by Bernd Haynold, through Wikimedia Commons.

It may be that in the aquatic habitat the snail effect on some bryophytes is much greater than in the terrestrial habitat. But it is not necessarily all bad. Steinman (1994) opined that snail grazing could account for the apparent unresponsiveness of epiphytes following phosphorus enrichment in a woodland stream in Tennessee, USA, where bryophytes were prominent. And some bryophytes seem prepared to fight back. The thallose liverwort Ricciocarpos natans (Figure 94) exhibits molluscicidal properties that are active against the snail carrier of schistosomiasis (Wurzel et al. 1990).

Figure 94. Ricciocarpos natans, a species with molluscicidal properties, floating on the water surface. Photo by Janice Glime.

Bryophyte Antifeedants

Figure 92. Fissidens fontanus, a moss that seems to be vulnerable to snail grazing except where it is protected by Fontinalis species. Photo by Michael Lüth, modified by Janice Glime, with permission.

Based on the foregoing discussion, it appears that at least some bryophytes are able to discourage browsing by slugs (Frahm & Kirchhoff 2002). Alcohol extracts of the moss Neckera crispa (Figure 95) and leafy liverwort Porella obtusata (Figure 96) have antifeedant activity against the slug Arion lusitanicus (Figure 97). Extracts of 0.5% dry weight of the moss had low activity, whereas

Chapter 4-8: Invertebrates: Molluscs

those from the liverwort exhibited moderate activity at only 0.05%. At 0.25% the antifeedant activity of Porella obtusata was complete. It is likely that this activity is not specific for slugs and may discourage insects, bacteria, and fungi as well.

Figure 95. Neckera crispa, a moss that has antifeedant activity against the slug Arion lusitanicus. Photo by Michael Lüth, with permission.

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Marchantia polymorpha (Figure 98) (Nils Cronberg, Bryonet 7 April 2016). Cronberg has observed this species feeding on Marchantia and has noticed that as the slug had invaded the wetland, Marchantia polymorpha had disappeared in parallel with the invasion.

Figure 98. Marchantia polymorpha showing a nibbled thallus on the upper left, about 1/3 down and 1/3 over from the corner. It also has a tear that is not likely the result of herbivory. Photo by James K. Lindsey, with permission.

Dispersal Agents It appears that slugs are not all bad in the bryophyte world and may instead be a necessary vector for some propaguliferous taxa (Stolzenburg 1995). Slugs and snails (Figure 99) leave a trail of mucous as they go, and as you well know if you have handled these molluscs, this secretion can be sticky. It is therefore no surprise that these animals have dispersal abilities.

Figure 96. Porella obtusata. Photo by Jan-Peter Frahm, with permission.

Figure 99. Snails such as this one traversing epiphytic mosses in Japan may be effective dispersal agents. Photo by Janice Glime.

Figure 97. Arion lusitanicus, a slug that traverses mosses, but finds Neckera crispa and Porella obtusata unpalatable. Photo by Mogens Engelund, through Wikipedia Commons.

On the other hand, Arion lusitanicus (Figure 97), also known as the murder slug, easily eats the thallose liverwort

Slugs are able to disperse the brood branches of Dicranum flagellare (Figure 100) (Kimmerer & Young 1995). These tiny branches become entrapped in the secretions and are deposited in the ensuing slime trail. Kimmerer and Young found that these can be transported at least 23 cm from the colony, although the mean distance in their study was only 3.7 cm.

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Figure 100. Dicranum flagellare showing the tight flagellate branches that can be dispersed by slugs. Photo by Janice Glime.

And it appears that the secretion increases the ability of the propagule to adhere to its substrate without affecting the germination rate. In fact, experiments by Davidson (1989) suggest that passage of spores through the slug's digestive system may enhance germination success. All plates containing mature spores from slug (Arion spp.; Figure 97) fecal pellets produced shoots, whereas only 80% of the plates with uneaten mature Mnium hornum (Figure 67-Figure 68) spores and 70% of those with uneaten Brachythecium rutabulum (Figure 101) spores produced shoots.

Figure 101. Brachythecium rutabulum, for which the spores germinate better if they have passed through the gut of a slug (Arion). Photo by Michael Lüth, with permission.

For those snails and slugs that nibble on spores, one might assume that not all spores end up inside them. Unless they have perfect aim with that huge foot, their somewhat clumsy feeding method is undoubtedly going to render some spores as passengers in the mucous on the foot. Sooner or later, these will be deposited in a new location. The ability of snails and slugs to glide across bryophytes and to climb setae to capsules suggests that these animals may be important as dispersal agents. But how widespread are herbivory and dispersal among bryophytes that temporarily host these slow-moving animals?

Although we know that bryophyte spores reach the mollusc gut, experiments are needed to see if spores expelled in feces are able to colonize successfully. Davidson (1989) found that Brachythecium rutabulum (Figure 101) and Mnium hornum (Figure 80) spores eaten by Arion species actually germinated better than controls. Manfred Türke sent me images of mosses in the feces of the slug Arion vulgaris (Figure 102). I was amazed at the size of the fragment of moss in the feces (Figure 103Figure 104). This is a potential means for dispersal, but the various species of bryophytes must be tested for viability. Digestive enzymes and extreme pH could damage the moss cells. On the other hand, the pathogenic fungi Phytophora spp. (Figure 105) survive as both oospores and filaments and are viable after passing through the digestive system of this slug species (Telfer et al. 2015). This was demonstrated by culturing the feces on agar.

Figure 102. Arion vulgaris, a slug that eats mosses, potentially dispersing them. Photo by Dilian Georgiev through Creative Commons.

Figure 103. Arion vulgaris feces with bryophytes and other material in it. Photo courtesy of Manfred Türke.

Figure 104. Arion vulgaris bryophyte from slug feces. Photo courtesy of Manfred Türke.

Chapter 4-8: Invertebrates: Molluscs

Figure 105. Phytophthora parasitica zoosporangia, a genus that survives passage through the gut of Arion vulgaris. Photo by Tashkoskip, through Creative Commons.

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Figure 107. Leptobryum pyriforme with capsules. Spores are able to pass through the guts of at least some slugs and remain viable. Photo by Michael Lüth, with permission.

To provide additional information on the potential dispersal ability of slug feces, Boch et al. (2013) fed capsules of four bryophyte species [Bryum pallescens (Figure 106), Funaria hygrometrica (Figure 69), Leptobryum pyriforme (Figure 107), Pellia endiviifolia (Figure 108)] to three slug species [Arion vulgaris (Figure 102), A. rufus; Figure 3, Figure 70, Figure 73), Limax cinereoniger (Figure 109)]. Among the 117 bryophyte samples, 51.3 % of the spore cultures had germination following gut passage.

Figure 108. Pellia endiviifolia with sporophytes. The spores of this species are able to pass through the gut of several slug species and remain viable. Photo by Janice Glime.

Figure 109. Limax cinereoniger on a mat of moss. Photo by Michal Maňas through Creative Commons. Figure 106. Bryum pallescens with capsules. Spores of this species pass through the guts of several slugs and retain their viability. Photo by David T. Holyoak, with permission.

Boch et al. (2013) found that germination rates did not differ among the bryophyte species, but the species of slug

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had strong effects. Among these three slugs, Limax cinereoniger (Figure 109) ate the lowest percentage of the bryophytes provided, and even correcting for that, they had the lowest percentage of feces samples (12.9%) producing protonemata. On the other hand, 76% of those of Arion vulgaris (Figure 102) and 74% of those of Arion rufus (Figure 3, Figure 70, Figure 73) produced protonemata (Figure 110).

Figure 110. Comparison of spore germination from bryophytes cultured from the feces of three species of slugs. White bars = Arion rufus; light grey bar = Arion vulgaris, dark grey bar = Limax cinereoniger. Redrawn from Boch et al. (2013).

Türke et al. (2013) provide evidence that slugs do indeed disperse fragments of mosses by consuming spores and fragments. For tracheophyte seeds, they suggested an average of 5 m dispersal distance, exceeding the typical less than 1 m in dispersal by ants. In some slugs, the seeds are destroyed in the digestive tract, but in other cases they remain viable propagules. Boch et al. (2015) discussed several ways that slugs benefit bryophytes. Their herbivory on tracheophytes (lignified vascular plants) permits more light to reach the low-growing bryophytes. But they also crawl across bryophytes and some eat the bryophytes. This puts them in the position to disperse spores, fragments, and other propagules. Nevertheless, documentation of the effect of the slugs on the bryophyte community is meager. Boch and coworkers (2015) designed a factorial common garden experiment to determine some of the effects of slugs on the bryophyte vegetation. They collected sporophytes of 11 native and 1 invasive bryophyte species [Barbula convoluta (Figure 111), Brachythecium rutabulum (Figure 101), Brachythecium velutinum (Figure 112), Bryum sp. (Figure 106), Campylopus introflexus (Figure 113), Ceratodon purpureus (Figure 114), Funaria hygrometrica (Figure 69), Leptobryum pyriforme (Figure 115), Marchantia polymorpha (Figure 98), Phascum cuspidatum (Figure 116), Plagiomnium affine agg. (Figure 117), Pohlia sp. (Figure 118)], representing 8 families. They used three enclosure treatments: slugs previously fed with bryophyte sporophytes, slugs that had not been fed sporophytes, no slugs. The researchers demonstrated that bryophyte cover increased in 21 days from 1.4% to 3.9% in plots where slugs had been fed, an increase that was 2.8 times higher than in the other two treatments. After eight months, the species richness was 2.6X higher (5.8 vs 2.2) than in the other treatments. The researchers concluded that the slugs contributed to

increasing bryophyte cover and diversity by reducing the dominance of tracheophytes. The early increase in cover in the enclosures with slugs fed sporophytes suggests that they also accomplish dispersal.

Figure 111. Barbula convoluta with capsules. Photo by Kristian Peters, with permission.

Figure 112. Brachythecium velutinum with unopened capsules. Photo by Michael Lüth, with permission.

Figure 113. Campylopus introflexus with capsules. Photo by Michael Lüth, with permission.

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Figure 117. Plagiomnium affine with developing capsules. Photo by Jan-Peter Frahm, with permission.

Figure 114. Ceratodon purpureus with young capsules, showing the normal proliferation. Photo by Michael Lüth, with permission.

Figure 118. Pohlia nutans with immature capsules. Photo by Michael Lüth, with permission.

Figure 115. Leptobryum pyriforme with numerous immature capsules. Photo by Michael Lüth, with permission.

Figure 116. Phascum cuspidatum with unopened capsules. Photo by Michael Lüth, with permission.

When the question of bryophyte dispersal by slugs arose on Bryonet, Scott Redhead (Bryonet 26 August 2016) suggested that this might even occur in the Splachnaceae. To that suggestion, Michael Lüth posted an image of Tetraplodon mnioides (Figure 119) showing one uneaten capsule and one that had been removed by an animal, possibly a slug, documenting his own observations of capsule herbivory. Christian Schröck (Bryonet 26 August 2016) likewise observed grazed capsules in Voitia and Tetraplodon. However, we need observations of feeding to determine the identity of the herbivores.

Figure 119. Tetraplodon mnioides with one capsule eaten by an unidentified herbivore. Photo by Michael Lüth, with permission.

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Lüth (2010) suggested that the pre-dispersal stage of the capsules on Splachnaceae are likely to attract herbivores that differ from the flies that spread the spores. At this earlier stage, the capsules have a different odor from that during the dispersal stage. This odor lasts for only a short time and is therefore often missed by field biologists. On Bryonet (26 August 2016), Lüth explained that Splachnum ampullaceum smells like Vaccinium oxycoccos and occurs in the same habitats, often blending with these cranberries. And Tetraplodon mnioides (Figure 119) smells like Vaccinium myrtillus. Although not all evolutionary successes are linked to adaptation, it makes one wonder if these early odors are adaptive to facilitate a longer dispersal and subsequent deposition in dung, although one might assume that would require a larger mammal, not a slug.

Figure 121. Zosterops japonicus, a bird that passes intact snails through the gut. Photo by Dick Daniels, through Creative Commons.

Figure 120. Splachnum ampullaceum sporophytes with a cranberry of similar color to the right. Photo by Michael Lüth, with permission.

I think most people would consider dispersal by snails and slugs to be distance-limited. But perhaps, with the help of birds, this is not so limited. Kawakami et al. (2003) demonstrated that the Japanese White-eyes (Zosterops japonicus; Figure 121) and the Brown-eared Bulbuls (Hypsipetes amaurotis; Figure 122) are birds that eat snails. In fact, five species of snails are able to remain in their shells and appear in the feces. If these snails had eaten moss spores, those spores might be transported a considerable distance, yet be viable in the gut of the snail. It is probably a rare event. Lots of questions remain in this relationship, but the scenario brings up interesting hypotheses. Malone (1965) discovered another possibility, exemplified by the Killdeer (Charadrius vociferus; Figure 123). Malone found two species of freshwater snails attached to the feet of the Killdeer. These were able to remain attached and viable long enough to effect dispersal. The snail Galba obrussa was able to survive 14 hours on Killdeer feet out of water. But the likelihood that an aquatic snail is carrying bryophyte spores is small due the rarity of capsules. Nevertheless, if a wetland snail has similar behavior, it has a better chance of having consumed spores from wetland mosses.

Figure 122. Hypsipetes amaurotis, a bird that passes intact snails through the gut. Photo by Nubobo, through Creative Commons.

Figure 123. Charadrius vociferus, a species that disperses snails on its feet. Photo by Andrew C, through Creative Commons.

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One additional factor determining the suitability of a slug for spore (or fragment) dispersal is the habitat where feces are likely to be deposited. Researchers have made the first steps in understanding the role of slugs in bryophyte dispersal, but much remains to be explored.

Bryophytes as Home Because of their small movement space, bryophytes can serve as safe sites for smaller snails. Birds can be significant consumers of snails, particularly during migration (Shachak & Steinberger 1980), and bryophytes can make the snails less conspicuous, if not hiding them completely. In terrestrial habitats, arachnids such as spiders and daddy-long-legs (Opiliones) are also predators on snails (Nyffeler & Symondson 2001). While some spiders can probably navigate the spaces within the moss mat, it seems unlikely that most mature daddy-long-legs could manage without getting caught. In addition to the arachnids, carabid beetles prey on terrestrial gastropods (Symondson 2004). Some of these beetles use a pump mechanism to extract the gastropod remains from its shell. Even snails are predators on slugs. The shell of the snail makes navigation among the bryophyte branches more difficult, potentially making the bryophytes a refuge for the smaller of vulnerable slugs. In a study of bryophyte inhabitants in the Bükk Mountains of Hungary, Varga (2008) found the tiny gastropods Punctum pygmaeum (Figure 124) and Pupilla muscorum (Figure 154) among the terrestrial mosses Plagiobryum zieri (Figure 125), Hypnum cupressiforme (Figure 126), and Tortella tortuosa (Figure 127). Standen (1898) found Punctum pygmaeum from moss shakings. From my own observations, it appears that snails and slugs are common on and even in bryophyte clumps, but finding documentation on the use of bryophytes by these small species evades even the aggressive Google search.

Figure 125. Plagiobryum zieri, a moss that supports the gastropods Punctum pygmaeum and Pupilla muscorum. Photo by Michael Lüth, with permission.

Figure 126. Slug on Hypnum. Photo by Janice Glime.

Figure 127. Tortella tortuosa in Europe. Photo by Michael Lüth, with permission.

Figure 124. The tiny Punctum pygmaeum on Ena montanum, both on a moss. Photo by Stefan Haller, with permission.

The European snails Azeca goodalli (Figure 128), Euconulus fulvus (Figure 129), Columella edentula (Figure 130), Discus (subgen Goniodiscus) rotundatus (Figure 131), Lauria cylindracea (Figure 132-Figure 133, Vertigo pusilla (Figure 134), and Vitrina pellucida (Figure

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135) live among mosses, among other substrata (Cloudsley-Thompson & Sankey 1961). Carychium tridentatum (Figure 136), Discus rotundatus, Cepaea hortensis (Figure 137), Oxychilus navarricus (formerly O. helveticus; Figure 138), and several rare species of Aegopinella (formerly in Retinella) [A. pura (Figure 139), A. nitidula (Figure 140-Figure 141)] are known under mossy brick rubble (Verdcourt 1954). Clausilia bidentata (10-11 mm; Figure 142) is also rare, but can be found under moss. Standen (1898) reported on Clausilia rugosa (Figure 143) swarming on mossy walls in the UK and feeding on mosses and lichens. Standen (1898) found the snail Acme lineata on a patch of the thallose liverwort Marchantia sp. (Figure 98). Figure 131. Discus rotundatus on moss. Christophe Quintin, through Creative Commons.

Photo by

Figure 132. Lauria cylindracea on bark. Christophe Quintin, through Creative Commons.

Photo by

Figure 128. Azeca goodalli shell. Photo by Francisco Welter Schultes, through Creative Commons.

Figure 129. Euconulus fulvus. Photo by Brian Eversham, with permission. Figure 133. Lauria cylindracea, whose small size can be seen in comparison to this seed. Photo by Christophe Quintin, through Creative Commons.

Figure 130. Columella edentula. Photo © Roy Anderson , with permission.

Figure 134. Vertigo pusilla on bark. Photo © Roy Anderson , with permission.

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Figure 138. Oxychilus navarricus on the moss Rhytidiadelphus squarrosus. Photo © Roy Anderson , with permission. Figure 135. Vetrina pellucida on bark. Anderson , with permission.

Photo © Roy

Figure 136. Carychium tridentatum on moss-covered branch. Photo © Roy Anderson , with permission.

Figure 139. Aegopinella pura on leaf litter. Photo © Roy Anderson , with permission.

Figure 137. Cepaea hortensis venturing into one of the Pottiaceae mosses. Photo by Stefan Haller, with permission.

Figure 140. Aegopinella nitidula on moss. Anderson , with permission.

Photo © Roy

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Figure 141. Aegopinella nitidula showing shell coils. Photo by Brian Eversham, with permission. Figure 144. Eucobresia diaphana on a species of the moss Tortula. Photo by Stefan Haller, with permission.

Figure 142. Clausilia bidentata on moss. Christophe Quintin, through Creative Commons.

Photo by

On the South Pacific Kermadec Islands, Iredale (1913) remarked that in dry weather one must look for the snails among the mosses, where they hide from the dryness. He commented that they are quite variable in choice of trees, with one bole producing a dozen or more while the next half dozen adjoining trees disclose none. Not surprisingly, new species still lurk amid the bryophytes. Efford (1998) found a new species of the carnivorous New Zealand endemic genus Rhytida (Figure 145), and reported observations by others of R. patula and R. meesoni perampla crawling on mosses and tree trunks at night. These and other New Zealand snails often fall prey to introduced predators. Wainuia urnula (Figure 146), another night-active snail on mosses, tree trunks, and rocks, was readily eaten by possums, rats, and hedgehogs in captivity. Efford (2000) found that 82% of the 315 W. urnula snails examined had an unusual food in the feces and gut – terrestrial amphipods. Its relative, W. edwardi (Figure 147), did not consume amphipods, and no other gastropod is known to consume them. The adaptation for consuming amphipods appeared to be largely behavioral, although there were some differences in the teeth.

Figure 143. Clausilia rugosa on bark, a species that eats mosses and lichens. Photo by O. Gargominy, through Creative Commons.

Eucobresia diaphana (Figure 144) lives in humid, cool places on mountains and in forests of Europe, where it is likely to encounter mosses, as seen in Figure 144 (Welter Schultes 2012b), but other than this picture, I can't verify what use it might make of them.

Figure 145. Rhytida otagoensis, member of a carnivorous genus that has some moss-dwellers. Image by James Atkinson, with permission.

Chapter 4-8: Invertebrates: Molluscs

Figure 146. Wainuia urnula, a tiny night-active New Zealand endemic snail that traverses mosses, as shown here. Photo by Andrew Spurgeon, with permission.

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Tropical islands, especially Hawaii, are particularly vulnerable to invasive species. With all the visitor traffic and import/export business, hitchhikers easily reach the islands. Snails are among these, and may be one of the causes of the apparent extinction of the bird called Po'ouli (Melamprosops phaeosoma; Figure 149) (Mountainspring et al. 1990). This native Hawaiian bird is especially adapted to feeding on land snails and insects on branches and under mosses, lichens, and bark. Its toes are large and are used for prying up moss and bark to acquire tree snails. The bill is stout, withstanding the force needed for manipulating the snails. Its demise is due largely to increased activity and habitat modification by feral pigs, avian disease, and possible gene pool impoverishment due to low numbers. But it also suffers competition for food by the introduced garlic snail (Oxychilus alliarius; Figure 150), a native of northwestern Europe (Welter Schultes 2012a) that emits a garlic odor when it is disturbed. This species is likewise a moss-dweller of mountain slope forests. It feeds on living and dead plant tissue, but it also consumes small snails and the eggs of other snails and slugs (Oxychilus 2011).

Figure 147. Wainuia edwardi, member of a genus that lives among mosses. Photo by James W. Atkinson, with permission.

Epiphytic Wiesenborn (2003) observed snails in the Riverside Mountains of California and found that the active snails preferred epiphytic mosses (Figure 148) and lichens compared to plant detritus and four sizes of rocks as habitat. They suggested that the epiphytes could provide these snails with food or moisture. Tree bark soon becomes a desert after the rain dries up, but mosses remain moist much longer, permitting the snails to be active longer and to search there for food where other small invertebrates likewise take refuge from desiccation.

Figure 148. Monachoides incarnatus on bark where it often encounters bryophytes. Photo by Stefan Haller, with permission.

Figure 149. Poʻouli (Melamprosops phaeosoma) on a mossy branch. Note the sturdy beak used to pry loose bark or crush snails found under bryophytes. Photo through Wikimedia Commons.

Figure 150. Oxychilus alliarius on moss on bark. Photo © Roy Anderson , with permission.

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The slug Prophysaon vanattae (scarletback taildropper; Figure 151) is one of those slugs that seems to find a safe site under mosses on trees on Vancouver Island, Canada (Kristiina Ovaska, pers. comm. 30 June 2009). But it also hangs on epiphytic moss mats in the moist deciduous forest there and may even lay eggs there (Figure 152).

Calcareous Areas Because of the need for calcium to make the shell, many snails are dependent on limestone habitats to obtain this important resource. Hence, this is a good place to look for snails on mosses growing there. Pupilla muscorum (Figure 154) is named for its occurrence among mosses in Great Britain, although it also occurs under stones and in leaf litter (Ehrmann 1956). This tiny (3-4 mm high shell) moss snail often prefers calciferous ground, but others describe it as indifferent to limestone content (Nordsieck 2012a). These snails are ovoviviparous. The eggs can survive over winter inside the female's body and are laid in the favorable conditions of spring. At that point, it is not the eggs that must survive because the juveniles usually hatch during oviposition. Pupilla triplicata (Figure 155) is likewise a moss dweller in Hungary and elsewhere (Deli et al. 2002).

Figure 151. Prophysaon vanattae, the scarletback taildropper, can be found hiding under mosses. Photo by Kristiina Ovaska, with permission.

Figure 154. Pupilla muscorum. Photo by Malcolm Storey, through Creative Commons.

Figure 152. Prophysaon vanattae with eggs on a moss. Photo by Kristiina Ovaska, with permission.

Pilsbry (1948) suggested that the pupillid snail Bothriopupa variolosa in eastern North America might prefer mossy rocks and trees.

Figure 155. Pupilla triplicata, a European moss dweller. Photo by O. Gargominy, through Creative Commons.

Figure 153. Bothriopupa tenuidens; B. variolosa seems to prefer mossy tree trunks and rocks. Image copyright Gary Rosenberg, www.DiscoverLife.

Another tiny conical snail (2-3 mm) of calcareous areas is Acicula fusca (Figure 156) in moss on chalk cliffs at Ballycastle, and on chalk underlying basalt at Black Head, Antrim, UK (Anderson 1996). And Pomatias elegans (Figure 157) occurs on mosses in limestone areas in the Burren, County Clare, UK (Platts et al. 2003).

Chapter 4-8: Invertebrates: Molluscs

Figure 156. Acicula fusca, a tiny snail that lives among mosses on chalk cliffs. Photo © Roy Anderson , with permission.

Figure 157. Pomatias elegans at Cheddar, Somerset, UK. Photo by Roger S. Key, with permission.

Trochulus (formerly Trichia) plebeia (Figure 158) occurs in wet mossy areas by springs in limestone areas (Gilbert et al. 2005). Trochulus villosus (Figure 159) lives in the German Alps and requires high moisture (Welter Schultes 2010), making bryophytes useful for maintaining that moisture. This strange genus of snails has hairs on its shell that help to hold it against wet surfaces (Gilbert et al. 2005). I don't have any indication that these hairs offer any particular help for living among bryophytes, but if they have any tactile properties, they could help keep it from getting stuck between branches by warning that the passage was getting too narrow.

Figure 158. Trochulus plebeia, a hairy snail, at Sugley Wood, UK. Photo by Brian Eversham, with permission.

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Figure 159. Trochulus villosus on mosses in Germany. Photo by Stefan Haller, with permission.

The European family Clausiliidae, known as door snails, derive their name from the "sliding door" that covers the opening of the shell (Wikipedia 2012a). This calcareous door is known as a clausilium, hence the family name. It permits the snail to retreat into its shell and seal it off against predators. Cochlodina laminata (Figure 160), the plaited door snail, lives "between mosses" as well as leaf litter, but may also be found climbing trees in deciduous forests and montane pine forests (Welter Schultes 2012b). Clausilia dubia (Figure 161) is a calciphilic inhabitant of humid, shady rocks and old walls, but also lives on tree trunks "full of moss." Michael Proctor (pers. comm. 23 April 2016) informed me that this species is very common on Carboniferous limestone in Yorkshire Dales, UK, in the bryophyte and lichen habitats. Macrogastra ventricosa (Figure 162), the ventricose door snail, lives in places with plentiful mosses on the forest floor or on tree trunks, mostly in the mountains (Welter Schultes 2012b). Macrogastra attenuata (Figure 163) lives between moss-covered rocks as well as on stones, rocks, and leaf litter in montane forests.

Figure 160. Cochlodina laminata on bark where it appears to be grazing mosses. Photo by Andrew Dunn, through Creative Commons.

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Bogs and Mires

Figure 161. Clausilia dubia with moss. Gargominy, through Creative Commons.

Photo by O.

Figure 162. Macrogastra ventricosa on moss. Photo by J. C. Schou, Biopix, through Creative Commons.

True bogs are acid, poor fens are acid, intermediate fens have intermediate pH levels, and rich fens are basic. For a snail, that pH range is an important consideration in choice of habitat because of the need for calcium in forming a shell. Because of this relationship, most malacologists have considered Sphagnum (Figure 164) peatlands, heathlands, and pine forests as unsuitable habitats for snails and consequently have poor snail biodiversity (Karlin 1961; Kerney & Cameron 1979; Horsák & Hájek 2003). In fact, Nekola (2010) found that highly and even moderately acidic sites had significantly (P70% cover of bryophytes with those having 70% bryophyte cover, whereas at Laamhel the addition of water plus nutrients was the only treatment that resulted in a large shift to greater survival with high bryophyte cover. Although van Tooren (1990) was unable to demonstrate significant effects of bryophytes in his 1990 study, he and his coworkers did find them on the same slope in the 1981 study (Keizer et al. 1985). Bryophytes under the growing conditions of that year significantly reduced mortality of the tracheophytes Linum catharticum and Carlina vulgaris. Apparently, bryophytes may serve as deterrents to slugs in some years when weather conditions might otherwise encourage herbivory, but provide little support for them in years when nutrients and/or water availability are different. Such interactions between species that change with the weather require further investigation.

Mussels (Bivalve Molluscs)

Figure 171. Planorbarius corneus. Photo © Roy Anderson , with permission.

Mussels are not common bryophyte inhabitants, but can occasionally occur there in aquatic environs. Frost (1942) found Sphaerium corneum (Figure 173) and four species of Pisidium (Figure 174) among the mosses in the

Chapter 4-8: Invertebrates: Molluscs

limestone stream in her River Liffey, UK, survey, but their typical niches were elsewhere in the stream. Some bivalve molluscs and other organisms can actually turn the relationship around and provide a home for the bryophytes. Yes, some of these animals actually have mosses growing on them. Neumann and Vidrine (1978) found Fissidens fontanus (Figure 92) and Leptodictyum riparium (Figure 175) growing on freshwater mussel shells.

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ECHINODERMATA I refuse to create a chapter for this marine phylum, but one observation is interesting enough to note here. Claudio Delgadillo-Moya (pers. comm. 30 March 2016) reported to me that a student who is working on sea urchins has found moss tissue in the gut of one and leafy liverwort fragments in another! There is no bryophyte known to be marine, but some do tolerate sea spray and live near the water. Most likely one of these, no, two of these, fell into the water or washed in from a stream or river. Resourceful urchin!

Summary

Figure 173. Sphaerium corneum on an aquatic plant. Photo © Roy Anderson , with permission.

Figure 174. Pisidium amnicum. Photo © Roy Anderson , with permission.

Figure 175. Leptodictyum riparium, a moss known to grow on freshwater mussels. Photo by Michael Lüth, with permission.

Snails and slugs (gastropods) have often been observed on bryophytes. They are adapted to land with a calcified slime epiphragm to cover the shell opening and respiratory pore in the body. A radula of many teeth permits them to scrape their food. Reproduction is mostly by simultaneous hermaphroditism. This may be facilitated by a love dart that facilitates movement of sperm cells to the sperm pouch by injecting hormones. Larvae develop within the egg in most so that the gastropods are typically oviparous. A few are known to deposit eggs in mosses. The white desert snail, Eremarionta immaculata, is common on bryophytes and seems to prefer them as a habitat. The copse snail, Arianta arbustorum is a night-active inhabitant. More quantitative studies have shown that some slugs and snails prefer bryophytes. More active snails might be found at night, whereas tiny snails might take refuge in the bryophytes during the day. Adaptations include "jumping" (Hemphillia), small size, conical snail, hibernation/estimation, and no shell (slugs). Snails might use them as a safe site to escape spiders, daddy-long-legs, and beetles, whereas other predators may lurk among the bryophytes. In streams, bryophytes may protect them from fish, ducks, shore birds, and amphibians. Bryophyte leafy plants and capsules can serve as food for snails and slugs, but some of these molluscs seem to avoid leaves with awns. Nutritional quality may be poor in some, and some have antiherbivore compounds that interfere with development, digestion, and palatability. In some cases the moss structure is such that the snails actually lose weight, whereas moss paste fosters a weight gain. But the gastropods may gain their nutrition from adhering algae and Cyanobacteria. In some cases protonemata and green capsules are preferred to leafy plants. Fissidens fontanus can be virtually eliminated by snails in lakes where there is no Fontinalis antipyretica to protect it. And some leafy mosses are palatable. But some slugs won't eat the moss even when they have been starved for 7 days. They have even been observed retreating from a moss. Various phenolic compounds seem to be involved in their reluctance to eat some bryophyte species. Ricciocarpos natans has molluscicidal properties that are effective against snail vectors of schistosomiasis.

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The moss may not offer any nutrition. Intact cells of leaves, capsules, and mature spores pass through the gut, and it seems that only young spores and protonemata become pale during their trip through the digestive system. Because of their mucous trail, slugs and snails are able to disperse some bryophytes, including brood branches, spores, and leaf fragments. And it appears that the mucous helps the dispersed fragment to adhere to its new substrate. Spores can even pass through the digestive system and survive, thus adding another form of dispersal. Gastropods can be common among epiphytes, avoid acid habitats, and abound in limestone habitats. Tiny mussels are able to live among bryophytes in aquatic habitats. Fissidens fontanus and Leptodictyum riparium can live on the shells. Echinoderms generally have no association with bryophytes, but if a bryophyte falls into the marine water it may occasionally be eaten.

Acknowledgments Bryonetters have been wonderful in making their photographs available to me and seeking photographs from others. Paul Davison has been helpful in providing suggestions and offering images. And a long time ago Allen Neumann sent me a specimen of a clam shell with Fissidens fontanus growing on it. Numerous photographers and malacologists have been helpful in providing images and information. Michael Lüth's photographs are a valued contribution. I thank all those photographers who have made their images available through the public domain.

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Stolzenburg, W. 1995. Partners in slime. Nature Conservancy Sept/Oct: 7. Symondson, W. O. C. 2004. Coleoptera (Carabidae, Staphylinidae, Lampyridae, Drilidae and Silphidae) as predators of terrestrial gastropods. In: Barker, G. M. (ed.). Natural Enemies of Terrestrial Molluscs. CABI Publishing, pp. 37-84. Szlavecz, K. 1986. Food selection and nocturnal behavior of the land snail Monadenia hillebrandi mariposa A. G. Smith (Pulmonata: Melminthoglyptidae). Veliger 29: 183-190. Telfer, K. H., Brurberg, M. B., Haukeland, S., Stensvand, A., and Talgo, V. 2015. Phytophthora survives the digestive system of the invasive slug Arion vulgaris. Eur. J. Plant Pathol. 142: 125-132. The Great Snail Hunt. 2012. The Open University. Accessed 22 April 2012 at . Tooren, B. van. 1990. Recruitment and establishment of shortlived flowering plant species in Dutch chalk grassland. In: Hillier, S. H., Walton, D. W. H., and Wells, D. A. (eds.). Calcareous Grasslands – Ecology and Management. Proceedings of a joint British Ecological Society/Nature Conservancy Council symposium, 14-16 September 1987 at the University of Sheffield. Bluntisham Books, Bluntisham, Huntingdon. Türke, M., Weisser, W. W., Knop, E., Fischer, C., and Boch, S. 2013. Gastropodochory 2.0: Slugs and snails disperse plant seeds, ferns, mosses and lichens – recent findings of what began in 1934. The Preliminary Program for 98th ESA Annual Meeting (August 4-9, 2013). Turton, W. 1840. A Manual of the Land and Fresh-water Shells of the British Islands. Longman, Orme, Brown, Green, and Longmans, London, p. 188. Varga, J. 2008. Analysis of the bryofauna of some moss species. Науковий вісник Ужгородського університету Серія Біологія, Випуск 23: 264-265. Verdcourt, B. 1954. The ecology of the Bedfordshire Mollusca (cont.). Bedfordshire Nat. 8: 16-17. Welter Schultes, Francisco. 2010. Species summary for Trochulus villosus. Updated 27 August 2010. Accessed 17 April 2012 at . Welter Schultes, Francisco. 2012a. Species summary for Oxychilus alliarius. Updated 30 January 2012. Accessed 15 April 2012 at . Welter Schultes, Francisco. 2012b. Native Door Snails (Clausiliidae). The Native World of Molluscs. Accessed 8 April 2012 at . Wiesenborn, W. D. 2003. White desertsnail, Eremarionta immaculata (Gastropoda: Pulmonata), activity during daylight after winter rainfall. Southw. Nat. 48: 202-207. Wikipedia. 2012a. Clausiliidae. Updated 18 March 2012. Accessed 8 April 2012 at . Wikipedia. 2012b. Reproductive system of gastropods. Updated 1 January 2012. Accessed 17 April 2012 at . Williamson, P. and Cameron, R. A. D. 1976. Natural diets of the landsnail, Cepaea nemoralis. Oikos 17: 493-500.

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Wurzel, G., Becker, H., Eicher, T., and Tiefensee, K. 1990. Molluscicidal properties of constituents from the liverwort Ricciocarpos natans and of synthetic lunularic acid derivatives. Planta Med. 56: 444-445. Yom-Tov, Y. 1971. Body temperature and light reflectance in two desert snails. J. Mollusc. Stud. 39: 319-326.

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Yom-Tov, Y. and Galun, M. 1971. Notes on the feeding habits of the desert snails Sphincterochila boissieri Charpentier and Trochoidea (Xerocrassa) seetzeni Charpentier. Veliger 14: 86-89.

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Chapter 4-8: Invertebrates: Molluscs

Glime, J. M. 2017. Tardigrade Survival. Chapt. 5-1. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. Ebook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 18 July 2020 and available at .

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CHAPTER 5-1 TARDIGRADE SURVIVAL

TABLE OF CONTENTS Tardigrades – Water Bears .................................................................................................................................. 5-1-2 Suitability of Bryophytes as Habitat ................................................................................................................... 5-1-3 Adaptations of Tardigrades ................................................................................................................................. 5-1-5 Survival of Hazardous Conditions ...................................................................................................................... 5-1-6 Physical Adaptations .................................................................................................................................... 5-1-7 Pigments ............................................................................................................................................... 5-1-8 Physiological Adaptations............................................................................................................................ 5-1-8 Light response ....................................................................................................................................... 5-1-8 Cryptobiosis .......................................................................................................................................... 5-1-9 Tun Formation .................................................................................................................................... 5-1-10 Dangers in a Tun ................................................................................................................................. 5-1-13 Effects of Size ..................................................................................................................................... 5-1-13 Longevity ............................................................................................................................................ 5-1-14 Dangers and Protective Mechanisms .................................................................................................. 5-1-15 Anhydrobiosis ..................................................................................................................................... 5-1-16 Osmobiosis ......................................................................................................................................... 5-1-17 Anoxybiosis ........................................................................................................................................ 5-1-17 Cryobiosis ........................................................................................................................................... 5-1-18 Temperature ........................................................................................................................................ 5-1-19 Diapause (Encystment) .............................................................................................................................. 5-1-19 Eggs ........................................................................................................................................................... 5-1-21 Migration? .................................................................................................................................................. 5-1-21 Summary ........................................................................................................................................................... 5-1-22 Acknowledgments ............................................................................................................................................. 5-1-22 Literature Cited ................................................................................................................................................. 5-1-22

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CHAPTER 5-1 TARDIGRADE SURVIVAL

Figure 1. Dactylobiotus sp. on the green alga Spirogyra. Photo by Yuuji Tsukii, with permission.

Tardigrades – Water Bears Tardigrades (tardus = slow, gradus = step, or slow walkers), also known as water bears or moss piglets, are close relatives of the arthropods (Garey et al. 1996, 1999; Giribet et al. 1996). Water bears resemble small bears (0.1-1 mm), complete with claws, but a few too many legs (4 pairs) (Figure 1). They are either armored (Heterotardigrada) or unarmored (Eutardigrada). The aquatic ones are usually a translucent white, whereas the terrestrial ones are often colored. Each of the eight legs has claws, which, when combined with their slow gait, makes them look very much like miniature polar bears with some extra legs. The very common Macrobiotus hufelandi (Figure 2) lumbers along at a maximum of 17.7 cm h-1 (Ramazzotti & Maucci in Mach 2010). Tardigrades are just the right size to move among the bryophyte leaves, they lumber along slowly like bears, and they are downright cute! Tardigrades, comprising about 900 species (Garey et al. 2008), can be found in marine, aquatic, and terrestrial habitats (Goeze 1773; May 1948; Greven 1980; Maucci 1986; Kinchin 1994). On land they frequently live in association with bryophytes (Figure 3; Figure 4) and lichens (Mihelčič 1967; Mehlen 1969; Utsugi 1984; Meininger et al. 1985; Mancardi 1988; Szymanska 1994; Bertolani & Rebecchi 1996; Tarter et al. 1996; Miller

1997; Jerez Jaimes et al. 2002; Boeckner et al. 2006; Bartels et al. 2009; Meyer & Hinton 2009; Rossi et al. 2009; Simmons et al. 2009). In water, algae, as well as bryophytes, provide homes.

Figure 2. Macrobiotus hufelandi, a common tardigrade that is among those inhabiting mosses. Photo by Paul Bartels, with permission.

These terrestrial tardigrades depend on the water drops that adhere to mosses and liverworts (Hingley 1993) and are therefore often termed limnoterrestrial (living in terrestrial habitats, but requiring a water film). Aquatic bryophytes can also house tardigrades (Hallas 1975; Kinchin 1987b, 1988; Steiner 1994a, b), as do the algae. However, of the ~1000 tardigrades reviewed by Guidetti

Chapter 5-1: Tardigrade Survival

and Bertolani (2005) and Garey et al. (2008), only 62 were truly aquatic. The others depend on water associated with the interstitial spaces of terrestrial algae, lichens, bryophytes, soil, and leaf litter. Water bears are found in habitats from hot springs to layers under the ice (in cryoconite holes in glaciers) and occupy every continent of the world.

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that have been studied, altitudinal gradients and microenvironmental variables (including substrate choice among vegetation, bryophytes, and leaf litter) seem to have gotten the bulk of the attention (Guil et al. 2009). Guil and coworkers found a unimodal distribution relative to altitude for species richness, but it was closely tied to habitat variation. The best predictor of the distribution of tardigrades in the Sierra de Guadarrama Mountain Range, Spain, was bioclimatic classification. Soil, climate, vegetation structure, and leaf litter type worked both separately and in combination to determine species richness, explaining nearly 60% of the species richness in micro-scale plots. Abundance, on the other hand, was significantly influenced only by soil composition and leaf litter type. The macro-environmental gradients seemed to be unimportant in determining abundance (e.g. Figure 6).

Figure 3. This tardigrade resided among the leaves of the moss Hypopterygium arbuscula (Figure 4). Photo courtesy of Filipe Osorio.

Figure 5. Echiniscus testudo dormant stage (tun), demonstrating the rigid nature of its armor that prevents it from extensive changes in size. Photo by Power & Syred through Creative Commons. Figure 4. Hypopterygium arbuscula, a known bryophyte habitat for tardigrades in Chile. Photo by Juan Larrain through Creative Commons.

Most of the terrestrial tardigrades are bryophyte inhabitants (Nelson 1991a). These terrestrial bryophyte taxa have a life span ranging 3-4 months (Franceschi et al. 1962-1963), 3-7 months for Macrobiotus hufelandi (Figure 2; Morgan 1977), up to about 3 months for roofmoss-dwelling Echiniscus testudo (Figure 5; Morgan 1977), to about 2 years (Altiero & Rebecchi 2001) of active life (not counting dormant periods). The bryophyteinhabiting taxa are more common in temperate and polar zones than in the tropics (Nelson 1991a). Some, as for example Echiniscus testudo (Figure 5), live almost exclusively on bryophytes (Corbet & Lan 1974). Despite their cosmopolitan distribution (Romano 2003), broad habitat requirements, and relative visibility (compared to protozoa, for example), the tardigrades remain poorly known. As late as 1985, Hidalgo and Coombs reported that 16 states in the USA had no records of tardigrades. Species not previously described are easily discovered by those who know where to look for them. The environmental conditions that affect tardigrades are likewise poorly studied (Guil et al. 2009), despite the extensive studies on a few species that have become travellers of the universe in space. Among those conditions

Figure 6. Echiniscus species (E. testudo occurs almost exclusively on bryophytes) seem to be unresponsive to moisture changes. Photo by Martin Mach, with permission.

Suitability of Bryophytes as Habitat The importance of bryophytes as a tardigrade habitat is evident by the number of publications on "moss" tardigrade fauna: Mihelčič 1967; Hallas 1975; Pilato & Sperlinga 1975; Morgan 1976; Bruegmann 1977; Morgan 1977; Maucci 1978, 1980; Bertolani 1983, 2001; Binda 1984;

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Utsugi 1984; Meininger et al. 1985; Hofmann 1987; Hofmann & Eichelberg 1987; Kinchin 1987a, b, 1988, 1994; Meininger & Spatt 1988; Mancardi 1988; Bertolani et al. 1990; Tarter & Nelson 1990; Kathman & Cross 1991; Nelson 1991a, b; Utsugi & Ohyama 1991; Moon et al. 1994; Szymanska 1994; Miller & Heatwole 1995; Adkins & Nelson 1996; Tarter et al. 1996; Hooie & Davison 2001; Guidetti & Jönsson 2002; Jönsson 2003; Meyer et al. 2003; Hooie 2005), to name a few. It appears that when tardigrade lovers want to collect a lot of them, they collect bryophytes and lichens – or just bryophytes (generally lumped into "mosses"). Unfortunately, the authors rarely name the bryophytes from which their prizes were extracted. However, some evidence suggests that little specificity exists for bryophyte species, and lichens are as suitable as bryophytes, with no apparent differences in tardigrade species (Meyer & Hinton 2007). I have to wonder, however, why reports on tardigrades from liverworts are so scant (Figure 7). Perhaps it is just as suggested to me by Łukasz Kaczmarek, that most zoologists do not understand the differences between mosses and liverworts. (Neither do my students when they begin looking at them.)

tardigrades have a prolonged life span when it is interrupted by such a dormancy period. And bryophytes contain food items, such as algae, protozoa, and nematodes, as well as the bryophytes themselves, sufficient for the tardigrades. Most likely, the small chambers among the bryophyte branches also afford protection from larger would-be predators. And when fragments of bryophytes disperse, they may carry tardigrades with them. It is the interstitial water of bryophytes that provides the suitable habitat for tardigrades (Hallas 1975). This water is typically found in leaf sheaths of bryophytes. Hallas investigated the drying of "cushions" of Hypnum cupressiforme (Figure 8). It required 19 hours for the moss to dry to a stable weight. The water retention relative to the weight of the cushion decreased linearly with the density of the cushion. However, the rate of drying can change with the temperature, saturation of the air, and air movement. He concluded that all compartments dry at the same rate, independent of the initial water content of each pocket, and the small variation in drying time is of no consequence for the tardigrades and other inhabitants. The tardigrades were concentrated in the living, chlorophyllous layer (compared to deeper, senescent layers) where there was more moisture. The water pockets connect vertically from one leaf to the next, but only connect horizontally when it rains. Therefore, horizontal migration of the tardigrades is only possible when the moss becomes saturated during rainfall. In H. cupressiforme (Figure 8), such bridges are formed only when rainfall exceeds 3 mm. Hallas considered that nighttime and morning dew were not sufficient for the most common tardigrade [Macrobiotus hufelandii (Figure 2) – a species that comprised 91% of the 386 tardigrades] to become active, suggesting that it would take ten times that amount of water to bring the moss to saturation levels.

Figure 7. SEM view of tardigrades on the lower sides of leaves of a leafy liverwort. Photo by Łukasz Kaczmarek and Łukasz Michalczyk, with permission.

Ramazzotti and Maucci (1983) considered mosses suitable habitat based on three needs of the limnoterrestrial tardigrades: 1. a structure that allows sufficient oxygen diffusion 2. the ability to undergo alternate periods of wetting and drying resulting from solar radiation and wind 3. a medium that contains sufficient food. Based on these criteria, bryophytes are particularly good habitats for tardigrades in several ways (Ramazzotti & Maucci 1983; Claps & Rossi 1984; Adkins & Nelson 1996). Their structure permits sufficient oxygen diffusion, both in aquatic and terrestrial habitats. Bryophytes experience drying, which most do slowly, permitting the tardigrades likewise to dry slowly, and both have a tolerance to dehydration that permits them to survive adverse conditions (Kinchin 1994). Furthermore, the

Figure 8. Mat of Hypnum cupressiforme. Photo by Dick Haaksma, with permission.

But bryophytes do pose their problems for the tiny tardigrades. These animals are quite light weight, so imagine their struggle to control their movements when they encounter fully hydrated bryophytes with a continuous bath of water surrounding them. Greven and Schüttler (2001) observed these slow-moving creatures [Macrobiotus sp., Echiniscus testudo (Figure 5) on Encalypta streptocarpa [=E. contorta] (Figure 9) when the bryophyte was fully hydrated. The poor bears could barely

Chapter 5-1: Tardigrade Survival

move and had difficulty maintaining the direction of their movements in the water. They could easily become dislodged by rainwater unless they are able to nestle in a leaf axil or other protected niche. And that is often a good place to look for them.

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roads were species adapted to xeric habitats. These species typically fed on fungi and algae, whereas those farther from the road were more likely to be omnivores or carnivores, presumably because they had more freedom to move about in a somewhat more moist environment.

Adaptations of Tardigrades

Figure 9. Encalypta streptocarpa, a tardigrade habitat that can be difficult to navigate when it is fully hydrated. Photo by Michael Lüth, with permission.

On the other hand, Polytrichastrum [=Polytrichum] formosum (Figure 10) did not sustain a continuous water film and the tardigrades seemed also unable to move in this "dry" habitat (Greven & Schüttler 2001). Rather, they seemed confined to the leaf axils, where water collected. As water receded, the animals ceased movement and formed a tun (protective dormant stage of tardigrade that is altered both chemically and physically) right there, permitting it to survive without water for up to 10 years (Jönsson & Bertolani 2001)! Perhaps tardigrades were the inspiration for the Rip Van Winkle story.

One might ask if these bryophyte-dwelling creatures have any special adaptations that permit them to live where they do. Their greatest adaptation is that they live in a habitat that permits them to dry slowly and go into a dormant state, as we will discuss shortly – a kind of behavioral/physiological adaptation. In fact, it appears that limnoterrestrial species actually require a habitat where they have dormant periods. And for many, the stylets permit them to suck the contents out of bryophyte cells, among other things, making them one of the few organisms specially adapted for obtaining nutrients from bryophytes. Like insects, tardigrades have chitin, in this case in the innermost layer of the cuticle. The chitinous armor of some terrestrial tardigrades (heterotardigrades) may slow drying and offer protection from damage while dry. Of course small size is essential for living in the miniature world of bryophytes. And their claws (Figure 11-Figure 13) may permit them to clamber about more easily among the leaves and branches of the bryophytes. But Bertolani and Biserov (1996) consider that the reduction of claws on the fourth pair of legs is an adaptation to moving among the interstitial spaces in the soil. Does this same adaptation pertain to those among bryophytes?

Figure 10. Polytrichastrum formosum, a moss that does not maintain a water film and is thus a poor tardigrade habitat. Photo by Des Callaghan, with permission.

Figure 11. Claws on four of the eight legs of Echiniscoides sigismundi (a tidal zone species). Photo by Martin Mach, with permission.

Moisture seems to be the greatest determinant of species distribution among bryophytes. Richness among epiphytic bryophytes in the Cincinnati, Ohio, USA area was greatest in areas of high humidity (Meininger et al. 1985). Hofmann and Eichelberg (1987) found that the tardigrades lacked correlation with bryophyte species but that their distribution could be predicted by the degree of moisture they prefer. It is therefore not surprising that some bryophytes housed no tardigrades. Tardigrades in association with roads along the Alaska pipeline demonstrate a moisture relationship (Meininger & Spatt 1988). Dust resulting from gravel roads associated with the pipeline alters the habitat for both mosses and tardigrades. Those tardigrades living among mosses near

Figure 12. Claws of a tardigrade that is most likely Cornechiniscus cornutus (a bryophyte-dweller). Photo by Martin Mach, with permission.

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Chapter 5-1: Tardigrade Survival

Figure 15. Cornechiniscus cornutus showing one of its two head horns. Photo by Martin Mach, with permission.

Figure 13. Claws of Echiniscus sp., a genus with many bryophyte-dwelling species. Photo by Martin Mach, with permission.

Their light weight facilitates tardigrade dispersal. Their bodies are flexible, permitting them to nestle in leaf axils or move in small spaces. But most of these as adaptations to the bryophyte habitat are speculation. There have been no tests to determine if any of these traits actually increases their survival among bryophytes compared to other habitats. Some very interesting experiments could be designed. Let's examine one of the bryophyte-dwelling tardigrades as an example of potential adaptations. Martin Mach (The Water Bear) found Cornechiniscus cornutus (Figure 14) among bryophytes on a mountain top in Hungary. This cute little bear has two horns on its head (Figure 15) and a nice salmon color. But it is slow and clumsy, out-classed by the faster-moving and more abundant Ramazzottius (formerly Hypsibius) oberhaeuseri (Figure 25). Do such ornamentations as horns and hairs help to reduce predation in this habitat? Is that an advantage to offset the slower movement? Does the bright color protect the water bear from UV damage, especially while it is dry?

Figure 14. Cornechiniscus cornutus. Mach, with permission.

Aquatic organisms rarely need to be concerned with desiccation. However, if an animal is to survive among terrestrial bryophytes, it must be prepared for drying when the bryophyte dries out, and many of the tardigrade habitats are in dry places, including cryptogamic crusts (assemblages of Cyanobacteria, algae, lichens, & mosses) in the prairie and desert, and among epiphytes on trees. These bring with them the very hazards mentioned above – UV light in the absence of water for protection, and extremes in temperature. And the watery body must be hydrated for oxygen to enter it. To unravel the relative importance of these stressors related to desiccation, Wright (1991) studied fifteen species of tardigrades and their responses to insolation, elevation, standardized desiccation rate, and hydration capacity of the plant substrate. There was considerable variation in ecotype among seven species with xeric associations. Macrobiotus hufelandi (Figure 2) and Hypsibius dujardini (Figure 16), both hygrophilic species, are absent from habitats that desiccate rapidly. On the other hand, the xerophiles Milnesium tardigradum (Figure 51) and Ramazzottius oberhaeuseri (Figure 25) avoid locations with high insolation and rapid desiccation rate, but also avoid poorly drained sites and sites with prolonged hydration. Despite these differences, Milnesium tardigradum often associates with the two Hypsibius species and may use them for food. The lack of association among Macrobiotus hufelandi, Paramacrobiotus (formerly Macrobiotus) richtersi (Figure 17), and Hypsibius prosostomus may be due to competitive exclusion.

Photo by Martin

Survival of Hazardous Conditions The biggest hazard a bryophyte imposes on a tardigrade is intermittent desiccation. But in addition to that desiccation, the organism may be subjected to high or low temperatures, low oxygen conditions, and UV light for prolonged periods. With little ability to move elsewhere, it needs some other type of protection.

Figure 16. Hypsibius dujardini with the alga Chlorococcum in its gut. Photo by Willow Gabriel through EOL Creative Commons.

Chapter 5-1: Tardigrade Survival

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tactile extensions to warn of an environment that is too tight, or chemical sensors to aid in finding food or sensing predators – or sensing a low-oxygen environment. Hypothesis testing is needed!

Figure 17. Paramacrobiotus richtersi. Photo by Science Photo Library through Creative Commons.

Physical Adaptations The soft-bodied tardigrades appear to have few structural adaptations to survive drought. Some, like Echiniscus, have long hairs (Figure 18-Figure 19), but the hairs are so few that one can hardly imagine they are of any help to reduce water loss or protect the dry animal. Hmmm...What might their function be? Or are they a nondetrimental left-over? One interesting idea is that they might serve as straws to draw water into the body – a hypothesis requiring both experimentation and TEM examination. But suppose that is true. It could be a way to cause water to enter without drowning the animal – i.e., permitting maintenance of an air layer on the surface. On the other hand, they could serve as fine hairs to collect dew and fog on their surface and direct it to the skin, thus increasing surface area and collection surface for the water. Or the hairs cold act like whiskers on a cat, facilitating navigation among the mosses. More interesting experiments are needed!

Figure 19. Two of the long hairs of Echiniscus. Photo by Martin Mach, with permission.

Echiniscus viridianus (Figure 20) seems to do well among bryophytes. It was originally described from mosses in Alabama USA (20 individuals) and later collected in New Mexico and the Azores Islands, again among mosses (Pilato et al. 2007). As in other members of the genus, this species bears armor and long hairs.

Figure 20. Echiniscus viridianus. Photo by Paul J. Bartels, with permission.

Figure 18. Echiniscus trisetosus, illustrating the sparse but long hairs and plates of armor. Photo by Łukasz Michalczyk and Łukasz Kaczmarek, with permission.

Oxygen availability can be a problem, and for this reason the tardigrades avoid dense bryophytes and usually remain in the top few centimeters of soil where more oxygen is available (Ramazzotti & Maucci 1983). Hence, another possibility for the long hairs is that they could be

Spines/hairs and body armor may offer a bit more protection. Some bryophyte-dwelling species such as Cornechiniscus cornutus (Figure 21) and some members of the genera Echiniscus (Figure 22-Figure 23) and Ramazzottius (Figure 24-Figure 25) (and others) have "armor" on their bodies that is somewhat leathery. I am aware of no studies that demonstrate the ability of the armor to reduce water loss, but it would appear to be a good possibility. Other possible advantages of this armorlike cuticle may include protection from fungi and other pathogens and some kinds of predators, particularly while in cryptobiosis, and it most likely would afford limited UV protection. How little we know!

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Chapter 5-1: Tardigrade Survival

Figure 21. Cornechiniscus cornutus showing armor. Photo by Martin Mach, with permission.

Figure 25. Ramazzottius oberhaeuseri, a tardigrade with armor. Photo by Martin Mach, with permission.

Pigments

Figure 22. Echiniscus sp. posterior dorsal side showing a type of armor. Photo by Martin Mach, with permission.

Figure 23. Tardigrade sp. showing a type of armor. Photo by Martin Mach, with permission.

Terrestrial tardigrades come in green, brown, yellow, orange, pink, red, purple, or black, whereas aquatic ones are white (Hebert 2008). Bonifacio et al. (2012) pointed out that despite the many studies on tardigrades relative to space travel, little is known about the nature or function of their pigments. They described the carotenoid nature of the pigments and the ability of these pigments to decrease under high oxidative stress. They hypothesized that these pigments had an anti-oxidant function and could possibly protect the animals during extreme environmental conditions. It is possible that the wonderful colors of some tardigrades (Figure 26-Figure 27) are adaptations against UV damage to DNA, especially during prolonged periods in a cryptobiotic state. Such pigmentation advantages have been demonstrated in bryophytes (Martínez Abaigar & Olivera 2007) and copepods (Byron 1982), so it is reasonable to expect them to serve similar functions in tardigrades, particularly in those more open habitats such as cryptogamic crusts. It would be an interesting study to examine the relationship of color with habitat in tardigrades. I am aware of no such study, but Martin Mach (pers. comm. 18 October 2012) pointed me to the publication of Ernst Marcus (1929). Marcus suggested that pigments were a response to UV light. He cited as support the findings that pigmentation varies with winter vs summer UV radiation. Experiments are needed to support this hypothesis, and one must wonder how this relates to those living among bryophytes. Marcus pointed out that insolation does not penetrate well into the moss colony, making bright pigments unnecessary for these tardigrades. Physiological Adaptations Light Response

Figure 24. Armor on Ramazzottius oberhaeuseri. Photo by Martin Mach, with permission.

Tardigrades have a pair of eyes, although at times they may be "ghost eyes" (Figure 28) that cannot be seen

Chapter 5-1: Tardigrade Survival

through ordinary observation (Mach 2012). These eyes respond to light, and at least in Macrobiotus hufelandi (Figure 2) the response changes with size and age (Beasley 2001). The smaller, younger size group had a significantly negative response to light. Beasley hypothesized that this behavior serves to conserve body moisture in small individuals that have a larger surface area to volume ratio than do larger ones. The response is not a phototaxis (directional response to light), but rather was photokinesis (non-directed, random movement), resulting in either an increase in speed or a change in direction when exposed to light. Such behavior would seem to support finding a "safe" place away from light.

Figure 26. Adult Echiniscus sp., demonstrating one of the bright colors found in tardigrades. Photo by Martin Mach, with permission.

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Cryptobiosis Albert Szent-Gyorgyi, a 20th Century Hungarian biochemist, once stated "Water is life's mater and matrix, mother and medium. There is no life without water." In their cryptobiotic state, tardigrades come close to disproving that statement. Anthony van Leuwenhoek first described cryptobiosis in 1702, but it was not until 1959 that Keilin coined the term cryptobiosis (Wright 2001). Cryptobiosis is a reversible ametabolic state that can be induced by dehydration and cooling, and possibly osmotic stress and anoxia. Metazoans such as tardigrades use glycerol and the disaccharide sugars sucrose and trehalose (Wright 2001) as protectants. Before entering the cryptobiotic state, these substances must be synthesized from glycogen reserves, hence requiring a preparatory period. Certain behavioral adaptations may help to delay the desiccation, thus permitting these compounds to be synthesized. One of the factors that contributes to the tolerance of desiccation is the ability to reduce surface area during tun formation (Wright 1991), hence slowing the process. Those that are most desiccation tolerant have the greatest infolding. This means those species with thick dorsal plates (Figure 29) are at a disadvantage. As Wright showed for Echiniscus testudo (Figure 5), there is very little surface area reduction possible.

Figure 29. Echiniscus mauccii showing the plates that make shrinkage during desiccation all but impossible. Photo by Diane Nelson and Paul Bartels, with permission. Figure 27. Cornechiniscus cornutus, a bryophyte-dwelling "horned" species that exhibits brilliant colors that could afford UV protection. Photo by Martin Mach, with permission.

Figure 28. Ghost eyes of Ramazzottius oberhaeuseri. Photo by Martin Mach, with permission.

Moss cushions help to make survival of this cryptobiotic state possible. The small spaces among mosses hold static air that can slow the dispersion of water vapor (Wright 1989). This permits the slow drying that is necessary for survival of the tardigrade in the desiccated state. Mosses in exposed positions may desiccate rapidly. Some mosses [e.g. Polytrichum (Figure 30), Dawsonia spp. (Figure 31)] are able to slow this process by folding their leaves (van Zanten 1974). Wright experimented with tardigrades on mosses in their natural habitat. Eutardigrada species were hydrated at least 24 hours before the experiments. As expected, there is considerable variation among species in their ability to tolerate desiccation. But they also differ in the lethal humidities (53-78%) for initial desiccation. Those species that are best able to tolerate rapid initial drying are also those most able to acquire tolerance to low humidities of 25-31% following drying in high humidity.

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reproduction stops, and metabolism is extremely reduced and may possibly even cease. For the limnoterrestrial (living in water films on land) tardigrade, it appears to be an essential part of survival and life, and it stops the aging clock.

Figure 30. Polytrichum formosum showing open leaves (left & right) and folded leaves (center). Photo by Michael Lüth, with permission.

Figure 32. Head region of Paramacrobiotus [=Macrobiotus] areolatus. The bulbous oval to the right of the three filaments (stylets and buccal tube) is the pharynx. Photo by Martin Mach, with permission.

Despite the apparent absence of structural adaptations, desiccated tardigrades, like their mossy habitats, have great survival capabilities. They have two forms of dormancy: cryptobiosis and encystment (Guidetti et al. 2006). The cryptobiosis of tardigrades is exhibited in several forms:

Figure 31. Dawsonia superba, a moss that seems unfavorable for tardigrade feeding and rolls its leaves when dry. Photo by Jan-Peter Frahm, with permission.

Typically, tardigrades are desiccated in 80% relative humidity (Wright 1991). In this condition, they dehydrate rapidly, then abruptly reduce water loss (the permeability slump). This slump occurs in both live animals prior to tun formation and in extended dead animals, so it is not a physiological phenomenon. This slump permits the animals to retain considerable water in their desiccated state. Crowe (1972) examined the humidity effects on Paramacrobiotus areolatus (Figure 32). He found that at humidities lower than 70% this species became flattened and crumpled. Above this level, dehydrating animals form tuns. This appears to be an active process that is not as effective in anesthetized animals. Tuns of active animals lose water at only 0.3 times the rate of anaesthetized animals. The anaesthetized animals reach moisture equilibration with the environment within one hour, whereas tuns do not reach that equilibrium within 100 hours. In dry air, tardigrades can reach as little as 2-3% water content without dying if they are able to dry properly. Literally meaning "hidden life," cryptobiosis is a state of suspended animation in which the organism is able to survive unfavorable conditions while expending little energy. During that state, the organism does not feed,

• • • •

anhydrobiosis (induced by loss of water) cryobiosis (induced by declining temperatures) anoxybiosis (induced by insufficient oxygen) osmobiosis (induced by loss of water due to higher external salt concentrations) (Bertolani et al. 2004).

To be active, tardigrades must stay in a water film in order to breathe (Bordenstein 2008). But in a cryptobiotic state, as discussed below, tardigrades can survive not only desiccation, but temperatures as low as 0.05K (-272.95ºC) for 20 hours or -200ºC for 20 months (Miller 1997). They have even survived 151ºC for a few minutes (Lindahl & Balser 1999). They become active again after living with 0% hydration (Lindahl & Balser 1999). This desiccated dormant state also permits them to survive pressures of 6000 atmospheres (Seki & Toyoshima 1998), i.e. six times the pressure of the deepest part of the oceans! Yet they can also survive the vacuum and UV radiation of space (Jönsson et al. 2008), a feat not known for any other animal. The ability of tardigrades to undergo cryptobiosis is more widely known than their encystment behavior. True cryptobiotic states are survived as a tun (Figure 33-Figure 43). The tardigrades will be the ones to survive when everything else is deceased. Tun Formation When they undergo desiccation, the tardigrades form a tun (Figure 33-Figure 43) (Lindahl & Balser 1999). The tun is a barrel-shaped, dry, dormant tardigrade. Tuns are

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formed in the process of entering true cryptobiosis, i.e., in anhydrobiosis, osmobiosis, and cryobiosis, but not in anoxybiosis. Although the stimulus differs among these, each ultimately involves the loss of free water.

Figure 36. Tun of Hypsibius sp. Photo by Martin Mach, with permission.

Figure 33. Tun of Ramazzottius oberhaeuseri. Photo by Martin Mach, with permission. Figure 37. Tun of Echiniscus sp. Photo by Martin Mach, with permission.

Figure 34. Tardigrade tun – water bear in a state of anhydrobiosis. Photo by Janice Glime.

Figure 38. Tun of Echiniscus sp. on moss leaf. Photo by Martin Mach, with permission.

Figure 35. Tardigrade tun – water bear in a state of anhydrobiosis. Note the buccal apparatus (resembles a tuning fork on left end). Photo by Janice Glime.

Figure 39. Multiple tuns of Echiniscus sp. on a single moss leaf. Photo by Martin Mach, with permission.

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Figure 40. Tun of Echiniscus sp. on moss leaf. Photo by Martin Mach, with permission.

Figure 41. Tun of Echiniscus sp. on a moss leaf. Photo by Martin Mach, with permission.

Figure 42. Tun of Echiniscus sp. Photo by Martin Mach, with permission.

a resting form in a cryptobiotic state in which the tardigrade appears to be dead (Crowe 1972). During tun formation, loss of free and bound water is greater than 95% (Bertolani et al. 2004). The body folds and the appendages are withdrawn (Lindahl & Balser 1999). Wax is extruded onto the surface and most likely reduces water loss (Wright 1988a, b). Those tardigrades with the most variability in the thickness of this cuticle, making them more pliable, are those able to have the greatest surface area reduction when they form tuns (Wright 1988a, 1989). The thin areas would permit greater infolding. Lipids of the inner cuticle are thickest in the species that are best able to tolerate rapid drying. Crowe (1972) demonstrated that the cuticle of Paramacrobiotus areolatus (Figure 32) is removed from air contact during tun formation and becomes less permeable to water. Crowe likewise suggested that this loss of permeability might be due to a lipid phase change,. Despite the waxy cuticular protection, the water content is reduced to less than 1% (Lindahl & Balser 1999) and the tun becomes shrivelled and wrinkled (Hingley 1993). Echiniscus testudo (Figure 5), an armored tardigrade, has much thicker dorsal (back) plates, apparently compensating for its limited ability to reduce surface area as it is drying (Wright 1988a, 1989). The tardigrade bodies synthesize cell protectants such as trehalose, glycerol, and heat shock proteins that contribute to successful recovery from the tun state (Wright 1989). Trehalose is typically found in high concentrations in animals in a state of cryptobiosis (Crowe & Crowe 1984). Trehalose is able to bond with DPPC and alter the spacing of polar head groups, apparently stabilizing the dry cell membranes. Hengherr et al. (2008b) determined that levels of trehalose varied considerably among species. In fact, they detected no changes in trehalose levels in any Heterotardigrada, and Milnesium tardigradum (Figure 51) apparently had no trehalose at all. They did demonstrate that tardigrade embryos can accumulate high levels of trehalose, seemingly explaining the high level of desiccation tolerance in that life cycle stage. Tun formation is essential to tardigrade survival under desiccating conditions. For Paramacrobiotus areolatus (Figure 44), and probably most tardigrades, if the humidity is low (90-day category) never hatched. They considered this variable hatching time to be a form of bet-hedging.

Figure 73. Racomitrium sudeticum, where Bertolanius volubilis in the Northern Apennines of Italy undergoes diapause, forming spring cysts that differ from winter cysts. Photo by Michael Lüth, with permission.

Figure 74. Racomitrium elongatum, a moss habitat in the Northern Apennines of Italy where Bertolanius volubilis makes different cysts in spring and winter. Photo by Michael Lüth, with permission.

Figure 76. Macrobiotus szeptyckii egg showing the highly decorated surface of eggs laid free from the exuvia. Photo by Łukasz Kaczmarek and Łukasz Michalczyk, with permission.

Eggs

Migration?

Eggs that are laid externally are typically ornamented (Figure 75-Figure 76) (Nelson 1991a). These may be laid singly or in groups.

Anhydrobiosis is not the only strategy available to organisms to escape drying conditions. Some organisms migrate to deeper levels of the moss or soil to escape

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drought. However, it appears that this option might not be available to many of the slow-moving tardigrades. Wright (1991) found that those tardigrades living in the interstices of the moss habitat migrate vertically to the soil C-zone (layer just above bedrock) as a means of escaping or slowing desiccation. The exception to this behavior is Echiniscus testudo (Figure 5). Nelson and Adkins (2001) examined this depth relationship in cushions of the moss Schistidium rivulare (=Grimmia alpicola; Figure 77). They found that among five species, only one (Echiniscus viridissimus) was more frequent in the top layer, regardless of the wet or dry condition of the moss. (Hmmm... Could the green that gives it its name indicate it has a photosynthetic symbiont that requires light, or just a penchant for green food?) Nelson and Adkins (2001) concluded that none of the Schistidium (Figure 77) inhabitants used migration as a means to escape reduction in moisture. They speculated that for tardigrade inhabitants of xeric mosses, there was no advantage to migration. Rather, they stayed put and went into a state of anhydrobiosis in both upper and lower layers.

Figure 77. Schistidium rivulare, a moss where excessive hydration can cause death to its tardigrade inhabitants. Photo by Michael Lüth, with permission.

Summary Tardigrades (water bears) are common in both aquatic and terrestrial bryophytes. The land dwellers require a water film and thus are called limnoterrestrial tardigrades. Despite their worldwide distribution, they are not well known. The bryophyte habitat offers sufficient oxygen, wetting and drying, sufficient food, a dispersal vehicle, and protection. Moisture is probably the most important factor in their distribution. Species of bryophytes do not seem to affect the types of tardigrades species. Tardigrades are adapted to the bryophyte habitat by their small size, stylets that permit sucking contents from bryophyte cells, flexible bodies, and a very responsive life cycle. Colored pigments in some may offer UV protection, especially during dry periods. Tardigrades can encyst or go into a cryptobiotic state as a tun. Cysts may differ between summer and winter. Tardigrades must dry slowly to survive the cryptobiotic

state. While in it, they are resistant to high and low temperature extremes, absence of water, extreme pressure, vacuum, and radiation. Anhydrobiosis is induced by diminishing hydration; cryobiosis is induced by low temperatures near 0ºC; osmobiosis is induced by a change in salinity; anoxybiosis is induced by low oxygen. Tardigrades form trehaloses that protect the cell membranes while dehydrated or at low temperatures. They typically can survive about 10 years in the tun, but one specimen resumed physiological activity after 120 years on a herbarium moss specimen, then died. Nevertheless, DNA damage accumulates during cryptobiosis; survival seems to be based on DNA repair. Furthermore, high temperatures and high humidity destroy trehalose. Another means of long-term survival is by producing resistant eggs. Variable hatching times may provide a form of bet-hedging in some species.

Acknowledgments Like all of my chapters, this one is really the product of the efforts of many biologists. Roberto Bertolani provided an invaluable update to the tardigrade taxonomic names, offered several suggestions on the text to provide clarification or correct errors, and obtained permission to use his published photographs from the Journal of Limnology. Paul Davison and Des Callahan have been helpful in providing suggestions and offering images. Filipe Osorio has sent me images several times, thinking of this project even when I was not soliciting help. Martin Mach and Yuuji Tsukii have given permission to use their many images that illustrate the species and life cycle stages. Martin Mach's website has been invaluable. Łukasz Kaczmarek has provided me with references, images, contact information, and many valuable comments on early stages of the manuscript. Martin Mach was kind enough to send me corrections for typos in the previous online version. Marty Janners and Eileen Dumire provided me with the views of two novices in the readability of the text. Thank you to Michael Lüth for permission to use his many images and to all those who have contributed their images to Wikimedia Commons and other public domain sites for all to use. I fear I have forgotten some who have helped – I have worked on this chapter for too many years!

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Society of America, Savannah, GA, 3-8 August 2003. Accessed on 3 July 2004 at . Mihelčič, F. 1967. Baummoose und Flechten als Lebenstätten fur Tardigraden. Carinthia II 77: 227-236. Miller, W. R. 1997. Tardigrades: Bears of the moss. Kans. School Nat. 43(3): 3-15. Miller, W. R. and Heatwole, H. 1995. Tardigrades of the Australian Antarctic Territories: the Mawson Coast, East Antarctica. Invert. Biol. 114: 27-38. Moon, S. Y., Kim, W., and Bertolani, R. 1994. Doryphoribius koreanus, new species of Tardigrada from Korea. Proc. Biol. Soc. Washington, DC 107: 514-516. Morgan, C. I. 1976. Studies on the British tardigrade fauna. Some zoogeographical and ecological notes. J. Nat. Hist. 10: 607-632. Morgan, C. I. 1977. Population dynamics of two species of Tardigrada, Macrobiotus hufelandii (Schultze) and Echiniscus (Echiniscus) testudo (Doyère), in roof moss from Swansea. J. Anim. Ecol. 46: 263-279. Nelson, D. R. 1991a. Tardigrada. Chapt. 15. In: Thorp, J. H. and Covich, A. P. Ecology and Classification of North American Freshwater Invertebrates. Academic Press, New York, pp. 501-521. Nelson, D. R. 1991b. A new species of Diphascon from New Brunswick, Canada (Tardigrada). Can. J. Zool. 69: 19111915. Nelson, D. R. 2002. Current status of the Tardigrada: Evolution and ecology. Integrat. Compar. Biol. 42: 652-659. Nelson, D. R. and Adkins, R. G. 2001. Distribution of tardigrades within a moss cushion: Do tardigrades migrate in response to changing moisture conditions? Zool. Anz. 240: 493-500. Neumann, S., Reuner, A., Brummer, F. and Schill, R. O. 2009. DNA damage in storage cells of anhydrobiotic tardigrades. Comparative Biochemistry and Physiology, Part A 153: 425429. Pilato, G. 1979. Correlations between cryptobiosis and other biological characteristics in some soil animals. Boll. Zool. 46: 319-332. Pilato, G. and Sperlinga, G. 1975. Tardigradi muscicoli di Sardegna. [Moss-inhabiting Tardigrada from Sardinia.]. Animalia 2(1-3): 79-90. Pilato, G., Fontoura, P., and Lisi, O. 2007. Remarks on the Echiniscus viridis group, with the description of a new species (Tardigrada, Echiniscidae). J. Limnol., 66(Suppl. 1): 33-39. Ramazzotti, G. and Maucci, W. 1983. The phylum Tardigrada – 3rd edition: English translation by C. W. Beasley. Mem. Ist. Ital. Idrobiol. Dott. Marco de Marchi 41: 1B680. Ramløv, H. and Westh, P. 1992. Survival of the cryptobiotic eutardigrade Adorybiotus coronifer during cooling to – 196°C: Effect of cooling rate, trehalose level, and short term acclimation. Cryobiology 29: 125-130. Ramløv, H. and Westh, P. 2001. Cryptobiosis in the eutardigrade Adorybiotus (Richtersius) coronifer: Tolerance to alcohols, temperature and de novo protein synthesis. Zool. Anz. 240: 517-523. Rebecchi, L., Boschini, D., Cesari, M., Lenicioni, V., Bertolani, R., and Guidetti, R. 2009a. Stress response of a boreoalpine species of tardigrade, Borealibius zetlandicus (Eutardigrada, Hypsibiidae). J. Limnol. 68: 64-70.

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Rebecchi, L., Cesari, M., Altiero, T., Frigieri, A., and Guidetti, R. 2009b. Survival and DNA degradation in anhydrobiotic tardigrades. J. Exper. Biol. 212: 4033-4039. Rebecchi, L., Guidetti, R., Borsari, S., Altiero, T., and Bertolani, R. 2008. Dynamics of long-term anhydrobiotic survival of lichen-dwelling tardigrades. Hydrobiologia 558: 23-30.

Reuner, A., Hengherr, S., Brümmer, F., and Schill, R. O. 2010. Comparative studies on storage cells in tardigrades during starvation and anhydrobiosis. Uncorrected proof. Accessed on 14 February 2010 at . Romano, F. A. III. 2003. On water bears. Florida Entomol. 86: 134-137. Rossi, G., Claps, M., and Ardohain, D. 2009. Tardigrades from northwestern Patagonia (Neuquén Province, Argentina) with the description of three new species. Zootaxa 2095: 21-36. Schill, R. O. and Fritz, G. B. 2008. Desiccation tolerance in embryonic stages of the tardigrade. J. Zool. 276: 103-107. Schokraie, E., Hotz-Wagenblatt, A., Warnken, U., Frohme, M., Dandekar, T., Schill, R. O., and Schnölzer, M. 2011. Investigating heat shock proteins of tardigrades in active versus anhydrobiotic state using shotgun proteomics. J. Zool. Syst. Evol. Res. 49 (Suppl 1): 111–119. Seki, K. and Toyoshima, M. 1998. Preserving tardigrades under pressure. Nature 395: 853-854. Simmons, B. L., Wall, D. H., Adams, B. J., Ayres, E., Barrett, J. E., and Virginia, R. A. 2009. Terrestrial mesofauna in above- and below-ground habitats: Taylor Valley, Antarctica. Polar Biol. 32: 1549-1558. Sømme, L. 1996. Anhydrobiosis and cold tolerance in tardigrades. Eur. J. Entomol. 93: 349-357. Steiner, W. 1994a. The influence of air pollution on mossdwelling animals: 1. Methodology and composition of flora and fauna. Rev. Suisse Zool. 101: 533-556. Steiner, W. A. 1994b. The influence of air pollution on mossdwelling animals: 2. Aquatic fauna with emphasis on Nematoda and Tardigrada. Rev. Suisse Zool. 101: 699-724. Szymanska, B. 1994. The Tardigrada from the Axel Heiberg Island and the associate bryophyte species. Folia Entomol. Hung. 55: 359-368. Tarter, D. C. and Nelson, D. R. 1990. An altitudinal comparison of the tardigrade fauna (Phylum: Tardigrada) from mosses on Spruce Mountain, West Virginia. 51st Annual Meeting of the Association of Southeastern Biologists, Baltimore, MD, USA, 18-20 Apr 1990. (World Meeting Number 902 5017). Tarter, D. C., Nelson, D. R., and Midkiff, E. F. 1996. New distributional records, including two state records, of tardigrades (Phylum: Tardigrada) from mosses in the Monongahela National Forest, West Virginia. [Abstract]. Bull. Assoc. S. E. Biol. 43: 147. Utsugi, K. 1984. Tardigrades found in the mosses of cities of Japan. Zoological Society of Japan, 55th Annual Meeting, Morioka, Japan, 27-29 Sep 1984. (World Meeting Number 843 5010). Utsugi, K. and Ohyama, Y. 1991. Tardigrades in King George Island (Antarctica). Zool. Sci. 8: 1198. Watanabe, M. 2006. Anhydrobiosis in invertebrates. Appl. Entomol. Zool. 41: 15-31. Węglarska, B. 1957. On the encystation in Tardigrada. Zool. Poloniae 8: 315-325. Westh, P. and Kristensen, R. 1992. Ice formation in the freezetolerant eutardigrades Adorybiotus coronifer and Amphibolus

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nebulosus studied by differential scanning calorimetry. Polar Biol. 12: 693-699. Westh, P. and Ramløv, H. J. 1991. Trehalose accumulation in the tardigrade Adorybiotus coronifer during anhydrobiosis. J. Exper. Zool. 258: 303-311. Westh, P., Kristiansen, J., and Hvidt, A. 1991. Ice-nucleating activity in the freeze-tolerant tardigrade Adorybiotus coronifer. Compar. Biochem. Physiol. A: Physiol. 99: 401404. Wikipedia: Cryptobiosis. 2009. Accessed on 2 February 2010 at . Wright, J. C. 1988a. Structural correlates of permeability and tun formation in tardigrade cuticle: An image analysis study. J. Ultrastruc. Molec. Struc. Res. 101: 23-39.

Wright, J. C. 1988b. The tardigrade cuticle. I. Fine structure and the distribution of lipids. Tissue Cell 20: 745-758. Wright, J. C. 1989. Desiccation tolerance and water-retentive mechanisms in tardigrades. J. Exper.. Biol. 142: 267-292. Wright, J. C. 1991. The significance of four xeric parametres in the ecology of terrestrial Tardigrada. J. Zool. 224: 59–77. Wright, J. C. 2001. Cryptobiosis 300 years on from van Leeuwenhoek: What have we learned about tardigrades? Zool. Anz. [J. Compar. Zool.] 240: 563-582. Wright, J. C., Westh, P., and Ramløv, H. 1992. Cryptobiosis in Tardigrada. Biol. Rev. 67: 1-29. Zanten, B. O. van. 1974. The hygroscopic movement of the leaves of Dawsonia and some other Polytrichaceae. Bull. Soc. Bot. Fr., Colloque Bryol. 121: 63-66.

Glime, J. M. 2017. Tardigrade Reproduction and Food. Chapt. 5-2. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. Ebook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 18 July 2020 and available at .

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CHAPTER 5-2 TARDIGRADE REPRODUCTION AND FOOD TABLE OF CONTENTS Life Cycle and Reproductive Strategies .............................................................................................................. 5-2-2 Reproductive Strategies and Habitat ............................................................................................................ 5-2-3 Eggs ............................................................................................................................................................. 5-2-3 Molting......................................................................................................................................................... 5-2-7 Cyclomorphosis ........................................................................................................................................... 5-2-7 Bryophytes as Food Reservoirs ........................................................................................................................... 5-2-8 Role in Food Web ...................................................................................................................................... 5-2-12 Summary ........................................................................................................................................................... 5-2-14 Acknowledgments ............................................................................................................................................. 5-2-15 Literature Cited ................................................................................................................................................. 5-2-15

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CHAPTER 5-2 TARDIGRADE REPRODUCTION AND FOOD

Figure 1. Echiniscus, a parthenogenetic genus with at least 146 described species (Guidetti & Bertolani 2005). This genus is common on bryophytes and reproduces there. Photo by Martin Mach, with permission.

Life Cycle and Reproductive Strategies One means by which organisms survive in such changeable habitats as bryophytes is by progressing to a different life cycle stage to wait out the storm – or lack of one. Tardigrades are especially adept at this, as seen in Chapter 5-1 (diapause and cryptobiosis). In tardigrades, diapause and cryptobiosis can occur at any time and developmental stage. Here we will look at reproduction and its role in further providing an escape route, at least for the species, if not the individual, an even that often occurs on bryophytes (Figure 1). Hofmann (1987) considers that tardigrades must be able to reproduce quickly and in sufficient numbers when conditions are favorable because their life style is one of intermittent activity and inactivity, the latter in either a state of dormancy or cryptobiosis. This constraint of brief reproductive periods and the necessity for a few individuals to have sufficient offspring makes them r-strategists. They lack a defined carrying capacity and the population density is dependent upon the length of time since establishment in that location. Life history of tardigrades can differ among species, presumably providing somewhat different adaptive strategies. For example, Paramacrobiotus tonollii (Figure 2) requires 16 days for its embryonic development whereas

Macrobiotus sapiens requires only 12 days (Lemloh et al. 2011). Paramacrobiotus tonollii is larger than M. sapiens but the latter has a longer life span of 83 days.

Figure 2. Paramacrobiotus tonollii. Photo by Paul Bartels, with permission.

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Reproductive Strategies and Habitat

Eggs

Reproductive mechanisms do differ among tardigrades in different habitats (Bertolani 2001). Those living among bryophytes, as well as those of freshwater, leaf litter, and soil, commonly are parthenogenetic (Figure 1), or more rarely hermaphrodites that self-fertilize. Marine species, on the other hand, have separate sexes. Bertolani hypothesized that organisms living in isolated and unstable habitats (including bryophytes) have evolved cryptobiosis, parthenogenesis, self-fertilization, and passive dispersal, benefitting them in their challenging living conditions. Passive wind dispersal of tardigrades with mosses is already known and may be their primary dispersal strategy (Pilato 1979). As is common among many mosses, those tardigrades that have parthenogenesis (equivalent to vegetative reproduction in mosses, i.e. reproduction without males) do not also have self-fertilization. These reproductive strategies, as in bryophytes, permit tardigrades to reproduce when only one individual, or its egg (Figure 3), arrives to colonize a new location.

Depending on the species, there are two ways tardigrade eggs (including embryos) may be deposited. Some lay free eggs on their substrate (Figure 3), but others deposit them in the shed exuvia of a molt (Mach: The Water Bear; Figure 4).

Figure 4. A number of eggs can develop within one shed exuvia, as shown here for Hypsibius sp. Photo by Yuuji Tsukii, with permission.

Macrobiotus hufelandi (Figure 5) has two sexes – males do exist (Figure 5) (Mach 2010). It is one of the species having free eggs (Figure 6). Eggs deposited outside the exuviae generally have decorative processes (Figure 7) (Mach 2010). Kinchin (1994) suggests that the functions of the egg processes include anchorage of the egg to a substrate or a transporting medium, defensive structure against being eaten by other animals, water reservoir which slows down the desiccation process, and regulation of gas exchange between egg and environment.

Figure 3. Eggs of a species of Dactylobiotus. The clustering of eggs may be beneficial in protecting each other, but their inherent resistance to almost everything suggests that is probably not important. Photo by Yuuji Tsukii, with permission.

Dispersal in tardigrades seems independent of the tardigrade because it is passive dispersal. But Bertolani et al. (1990) considered that the differences in distribution and frequency of members of the genus Ramazzottius relate to their differences in reproductive modes. In their study of Ramazzottius species on bryophytes and lichens, they found that the sex ratio was strongly influenced by the type of reproduction, but also differed when samples were collected from tree trunks and limited rocky areas versus extensive rocky outcrops. Bertolani et al. found that bryophyte-dwelling Ramazzottius tardigrades from tree trunks or slightly rocky areas exhibited parthenogenesis and absence of male tardigrades. Among the rocky outcrops, there were always at least some males, although some parthenogenesis still occurred. Males are only useful if there is sufficient opportunity for contact with females. Perhaps the rocky outcrops provided less of a labyrinth and permitted the needed contact? Eggs provide light-weight, windborne propagules to disperse the species (Figure 3). [To clarify for botanists, some references tend to use the term egg for the zygote and sometimes even the developing organism (embryo) until it has hatched, like the hatching of a bird egg. Since I found the term egg used in my references, I shall use egg here as well.]

Figure 5. Macrobiotus hufelandi male. Photo by Martin Mach, with permission.

Figure 6. Egg of Macrobiotus hufelandi, demonstrating the decorative processes on this free-egg deposit. Photo by Martin Mach, with permission.

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Figure 10. Individual of Milnesium tardigradum with only three large eggs in the exuvia. Photo by Martin Mach, with permission.

Figure 7. Egg of Macrobiotus sp., demonstrating the highly decorative surface that is typical of eggs laid free of the organism. Photo by Martin Mach, with permission.

When healthy adult tardigrades discard their outer covering, many taxa deposit eggs in these shed exuviae (outer "skins") (Figure 8-Figure 11) (Bertolani et al. 2009). The eggs may be few or many (up to 30-40) and may differ even within the same species, as can be seen for Milnesium tardigradum in Figure 9-Figure 11 (Altiero et al. 2006). The number of eggs depends on the species, but also on the nutritional status of the individual female (Mach: The Water Bear). And it seems that some bears may even ingest their own eggs to improve their nutritional status. Egg development is poorly known. In Paramacrobiotus [=Macrobiotus] richtersi (Figure 12) it can be prolonged to 90 days or more if the eggs undergo desiccation and become resting eggs (Altiero et al. 2009). The non-resting (subitaneous) eggs may hatch in as little as 30-40 days.

Figure 8. These eggs reside in the shed exuvial "armor" of the parent and permit the tardigrade species to survive winter and desiccation. Photo by Martin Mach, with permission.

Figure 9. Individual of Milnesium tardigradum with eleven eggs in the shed exuvia. Photo by Martin Mach, with permission.

Figure 11. Developed eggs of Milnesium tardigradum with the buccal apparatus visible, indicating nearness to maturity. Photo by Martin Mach, with permission.

Figure 12. Paramacrobiotus richtersi. Photo from Science Photo Library, through Creative Commons.

The eggs generally develop within the exuvia until the fully-formed tardigrade is ready to leave the egg, as shown here for individuals in the genus Echiniscus (Figure 13Figure 19), requiring several weeks for completion (Mach: The Water Bear). The eggs are able to survive the same drying conditions as the adult; development stops during that dry state. The young tardigrades resemble the adults (Figure 20-Figure 23), but are smaller, requiring a series of molts as they grow. Growth occurs by cell enlargement rather than by addition of cells. Since the eggs often remain in the exuvia until they hatch (Figure 21), size would tend to reduce wind-dispersal of the pollen-grainsized eggs except when they are dispersed along with a substrate such as mosses.

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Figure 13. Echiniscus adult. Photo by Martin Mach, with permission.

Figure 17. Morula stage in the embryo development of Echiniscus. Photo by Martin Mach, with permission.

Figure 14. Echiniscus exuvia with eggs (embryos) after first division. Photo by Martin Mach, with permission.

Figure 18. This mature "egg" of Echiniscus can be found among bryophytes, and its smooth surface is typical of eggs that are kept within the exuvia. Note the buccal apparatus that signifies its late developmental stage. Photo by Martin Mach, with permission.

Figure 15. Echiniscus embryo after two divisions. Photo by Martin Mach, with permission.

In soil-dwelling Paramacrobiotus richtersi (Figure 12; also a known bryophyte dweller), temperature played a role in rate of development, survival rate, body growth, and generation time (Figure 24; Hohberg 2006). On the other hand, hatching time, first to fourth molts, and maturation time were dependent upon body size alone.

Figure 16. Multicellular Echiniscus embryo Martin Mach, with permission.

Figure 19. Echiniscus hatching from its eggs. Photo by Martin Mach, with permission.

Photo by

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Chapter 5-2: Tardigrade Reproduction and Food

Figure 20. Moss-dweller Macrobiotus derkai hatching. from a free "egg." Photo by Łukasz Kaczmarek and Łukasz Michalczyk, with permission.

Figure 23. Echiniscus young. Photo by Martin Mach, with permission.

Some tardigrades have found another safe site for their eggs. They can use the capsule of a moss as an egg depository (Mach: The Water Bear; Figure 25-Figure 26 ).

Figure 21. Despite the large number of eggs/embryos, it appears that most of them are successful in hatching into young tardigrades, as seen here for a species of Hypsibius. Photo by Yuuji Tsukii, with permission.

Figure 24. Effects of temperature on the development of Paramacrobiotus richtersi (Figure 12), starting with the day the tardigrades hatched. Body lengths are for hatching and first oviposition only. Redrawn from Hohberg (2006).

Figure 22. Echiniscus young and old. Note the long "hairs" extending from the body, giving the genus its name. Photo by Martin Mach, with permission.

Figure 25. Moss capsule with tardigrade (with green gut) and two white eggs. Photo by Martin Mach, with permission.

Chapter 5-2: Tardigrade Reproduction and Food

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Figure 26. This egg is protected by a moss capsule. Based on the decorations on the eggs, they appear to be close to Macrobiotus hufelandi. Photos by Martin Mach, with permission.

Molting Like its relatives in the Arthropoda, the tardigrade must molt (Figure 27-Figure 35). This process usually requires 5-10 days and occurs several times throughout its life (Walz 1982), including after sexual maturity while the body is still increasing in size (Nelson 1982). During molting, the old cuticle, claws, and lining of the fore- and hindgut are shed (Figure 28), causing a stage known as the simplex stage (Figure 35). Lacking its sclerified parts of the buccal-pharyngeal apparatus, the tardigrade cannot feed. It appears that tardigrades molt 4-12 times during their 3-30 months of active lives (Nelson 2002).

Figure 29. Eggs in the shed exuvia of Milnesium tardigradum. Photo by Martin Mach, with permission.

Figure 30. Milnesium tardigradum eggs in its shed exuvia. Photo by Martin Mach, with permission.

Figure 27. Milnesium tardigradum as it recedes from its cuticle in preparation for molting. Note the dark brown eggs that will soon be left behind. Photo by Martin Mach, with permission.

Figure 28. Milnesium tardigradum emerging from its exuvia during molting, leaving its claws, eggs, and various other parts behind. Photo by Martin Mach, with permission.

Cyclomorphosis It appears that cyclomorphosis (annual cycle of morphological change) occurs in tardigrades, although it has been demonstrated in only a few species. It has been documented in the marine species Halobiotus crispae (Kristensen 1982; Halberg et al. 2009). Likewise, Dastych (1993) demonstrated cyclomorphosis in a cryoconitedwelling species of Hypsibius (Figure 31), and in a bryophyte dweller. Furthermore, Rebecchi and Bertolani (1994) did demonstrate it for one species in the genus Bertolanius [=Amphibolus] (Figure 33), which does have moss-dwelling species. Kristensen (1982) studied the marine Halobiotus crispae cycle and found two morphs. In winter there is a pseudosimplex stage that hibernates and is sexually immature. These winter forms gather in large aggregations in protected areas where the aggregations increase chances for survival of the freezing temperatures. The population experiences synchronous development of gonads, hence all reaching sexual maturity and breeding simultaneously. But the cycle for other taxa and habitats, including bryophytes, remains to be explored (Nelson 2002).

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bryophytes, the stylet may be very long, permitting penetration of the thick cellulose walls of bryophytes. For example, Echiniscus testudo (Figure 37) feeds primarily on bryophytes (Morgan 1977). Diphascon (Figure 52), also a bryophyte dweller, has a flexible buccal tube with spiral rings resembling the extension on a vacuum cleaner. Small bryophyte dwellers may subsist on diatoms and bacteria that live epiphytically among the bryophytes [Bartels 2005; Tardigrada (Water Bears) 2005].

Figure 32. Hypsibius dujardini with 3 oocytes. Photo by Willow Gabriel, through EOL Creative Commons.

Figure 35. Pseudobiotus sp. shedding its cuticular exuvia and leaving its eggs/embryos behind. Photo by Paul Davison, with permission. Figure 33. Bertolanius volubilis cuticle with a type A cyst inside. Photo by Roberto Bertolani, with permission.

Figure 36. Echiniscus perviridis with green color, most likely due to its vegetarian diet. Echiniscus testudo is known to feed primarily on bryophytes. Photo by Łukasz Kaczmarek, with permission. Figure 34. Ramazzottius oberhaeuseri completing its molt out of its old cuticle. The emerging organism will remain in this simplex stage until it rebuilds its cuticular parts. Photo by Martin Mach, with permission.

Bryophytes as Food Reservoirs Bryophyte-dwelling tardigrades include both bryophyte-eating tardigrades and those with a variety of other feeding strategies, including carnivory. The tardigrade has a specially adapted pair of stylets (Figure 49) and a muscular pharynx (Figure 50-Figure 52) that produces a suction into the gut, permitting the tardigrade to suck fluids from the interior of a bryophyte or algal cell (Figure 53) or even small animals such as rotifers (Figure 54) and nematodes [Tardigrada (Water Bears) 2005]. In the family Echiniscidae (Figure 36), a common family on

Figure 37. Echiniscus testudo tun. Syred through Creative Commons.

Photo by Power &

Chapter 5-2: Tardigrade Reproduction and Food

Schill et al. (2011) consider the bryophytes to be a "rich food supply for both carnivorous and herbivorous species." These food sources include nematodes, rotifers, plant cells, algae (Figure 38), yeast, and bacteria, and for some, bryophytes. Schill and coworkers conducted a genetic tracer study (rbcL) on the guts of tardigrade species from various sites in Europe that demonstrated the presence of mosses from the Erpodiaceae [Aulacopilum hodgkinsoniae, Venturiella sinensis (Figure 39)] and Pottiaceae [Syntrichia (=Tortula) obtusissima (Figure 40)] in the guts of field-collected Macrobiotus sapiens, Grimmiaceae [Grimmia elongata (Figure 41), Coscinodon cribrosus (Figure 42), Schistidium strictum (Figure 43)] from Macrobiotus persimilis and Echiniscus granulatus, and the green alga Trebouxia (Figure 44) from Richtersius coronifer (Figure 38). For Macrobiotus sapiens they found no rbcL sequence demonstrating presence of the families Pottiaceae or Orthotrichaceae. It appears that Macrobiotus sapiens will only eat these latter two moss families when Grimmiaceae is not available, or that others had been digested completely before samples were extracted. The small tardigrade stylet makes it difficult for them to obtain cell contents from the moss genera Polytrichum (Figure 45), Dicranum (Figure 46), Leucobryum (Figure 47), and Racomitrium (Figure 48). Digestion in tardigrades is aided by the gut pH, with the foregut having an acidic environment and the midgut having a basic environment (Marcus 1928).

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Figure 40. Syntrichia obtusissima showing hair points. Photo by Claudio Delgadillo, with permission.

Figure 38. Richtersius coronifer, clinging to an algal cell. Photo by Martin Mach, with permission.

Figure 39. Venturiella sinensis, a moss eaten by Macrobiotus sapiens. Photo from Digital Museum, Hiroshima University, with permission, with permission.

Figure 41. Grimmia elongata, a moss eaten by Macrobiotus persimilis and Echiniscus granulatus. Photo by Michael Lüth, with permission.

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Figure 44. Trebouxia, a lichen symbiont that appeared in the guts of field collected Macrobiotus persimilis and Echiniscus granulatus. Photo by Yuuki Tsukii, with permission.

Figure 45. Polytrichum commune, a moss with thick leaves that make feeding by tardigrades difficult. Photo by Michael Lüth, with permission.

Figure 42. Coscinodon cribrosus, a moss that is suitable habitat and food for Macrobiotus persimilis and Echiniscus granulatus. Photo by Michael Lüth, with permission.

Figure 43. Schistidium strictum, a moss that is eaten by Macrobiotus persimilis and Echiniscus granulatus. Photo by Jan-Peter Frahm, with permission.

Figure 46. Dicranum scoparium, a moss with leaves that seem to make feeding by tardigrades difficult. Photo by Janice Glime.

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nebulosus are widespread in many habitats, including bryophytes [Tardigrada (Water Bears) 2005]. They eat nematodes, rotifers, and smaller tardigrades, but still use the stylet to suck out cell contents. Suzuki (2003) reared Milnesium tardigradum from the moss Bryum argenteum (Figure 55), using only rotifers [Lecane inermis, common in wet Sphagnum (Miller 1931)] as food.

Figure 47. Leucobryum glaucum, showing thick leaves that make tardigrade feeding difficult. Photo by James K. Lindsey, with permission.

Figure 50. This tardigrade has the stylets withdrawn into its head. The pharynx is in the center behind the stylets. Photo by Paul Davison, with permission.

Figure 48. Racomitrium macounii ssp macounii, a moss with leaves that seem to make feeding by tardigrades difficult, in Europe. Photo by Michael Lüth, with permission.

Figure 51. "Head" region of Milnesium tardigradum showing the pharynx. Photo by Martin Mach, with permission.

Figure 49. Echiniscus with the stylets protruding (out of focus). Photo by Martin Mach, with permission.

Tardigrades even consume smaller tardigrades. Larger species such as those of Macrobiotus (Figure 5) and Milnesium (Figure 9, Figure 51) consume smaller members such as Diphascon (Figure 52) and Hypsibius (Figure 64), as exhibited by remains of claws and buccal apparati (Figure 57) in the gut (Nelson 2002). Large carnivorous Eutardigrada such as Paramacrobiotus richtersi (Figure 12), Milnesium tardigradum (Figure 9, Figure 10, Figure 51, Figure 54), and Bertolanius

Figure 52. Pharynx (oval) of Diphascon, the organ that produces the suction for the stylets. Photo by Martin Mach, with permission.

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Figure 53. The green in this tardigrade is likely to be algae or moss. Photo by Paul Davison, with permission.

difference Suren interpreted to reflect a loss of bryophytes as a food source. It seems to be one of the few animals specifically adapted to obtaining the good stuff from the insides of the cells of bryophytes. Its stylets (Figure 49Figure 50) serve as a miniature needle and straw to puncture the cell and suck the nutrients from it. The pharynx (Figure 52) serves as a pump to draw fluids in through the stylets (Tardigrades, Bears of the Moss). It appears that eating bryophytes requires more than just the equipment to suck the good stuff out of the cell. The excretory system seems also to be altered. Węglarska (1990) found that in four genera of tardigrades, those that live among bryophytes have larger excretory organs relative to body size than do the freshwater species. The purpose of this added size remains a mystery. Ramazzotti and Maucci (1983) suggested that excretion probably occurs in four ways in tardigrades. At molting it occurs through the salivary glands. Likewise, when the cuticle is shed it removes accumulated excretory granules. It can occur through the wall of the midgut. And in the eutardigrades, it occurs through excretory glands. There is no study to determine how these various mechanisms might relate to a diet of bryophytes. Role in Food Web

Figure 54. Milnesium with the mastax of rotifers visible in the gut (black arrows). Photo by Martin Mach, with permission.

As seen above, tardigrades typically are either plant eaters or are carnivorous (Garey et al. 2008), including protozoa, nematodes, and rotifers (Figure 54), but also consume bacteria and fungi (Kinchin 1988). As noted in the earlier chapter on nematodes, they can be predators on nematodes that live in the same clump of moss (SánchezMoreno et al. 2008), making them important consumers and often the top carnivore. Both Paramacrobiotus [=Macrobiotus] richtersi (Figure 12) and Macrobiotus harmsworthi (Figure 56Figure 57) caused significant declines in the nematode populations, thus regulating the food web. In fact, a single P. richtersi dined on an average of 61 nematodes in a day! Unlike many of the slow-walking water bears, these carnivorous water bears are able to move swiftly to attack and devour their prey (Kristensen & Sørensen 2005). Davison (2005) reports that tardigrades lumber across the substrate, swinging their heads back and forth in search of food. When he offered them nematodes and rotifers, the tardigrades made no attempt to eat them. When he offered them a larger choice, the annelid Lumbriculus sp., a genus with known members that inhabit mosses, he found that they immediately approached it and began eating it.

Figure 55. Bryum argenteum, a moss known to house Milnesium tardigradum. Photo by Michael Lüth, with permission.

Suren (1988) attempted to determine the importance of bryophytes as food vs. simply substrate by using artificial mosses in high alpine New Zealand streams. When artificial structures were used, similar communities of invertebrates developed, but tardigrades appeared to be affected negatively by the absence of the bryophytes, a

Figure 56. Macrobiotus harmsworthi, a nematode predator. Photo by Paul J. Bartels, with permission.

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Figure 57. Macrobiotus harmsworthi buccal apparatus. Photo by Paul Bartels, with permission.

Tardigrade specialists have assumed that the buccal apparatus (Figure 58-Figure 60) indicates characteristics of the food, but no studies exist on the relationships of buccal apparati among the limnoterrestrial taxa (Nelson 2002).

Figure 60. Dactylobiotus dispar has a buccal apparatus similar to that of Paramacrobiotus areolatus. Photo by Martin Mach, with permission.

But tardigrades can have their predators too. Snails that live among the moss leaves could enjoy a meal of tardigrades (Fox 1966). The land snail Bulimulus guadalupensis (Figure 61) from Puerto Rico had evidence that all life cycle stages of the tardigrade Echiniscus molluscorum (see Figure 62) live in its feces (Fox & Garcia-Moll 1962). It is not clear if these passed unharmed through the gut or if they took advantage of the feces as a food source after defecation. It is even possible that eggs passed through the gut and hatched in the feces.

Figure 58. The three "filaments" and pharynx are the buccal apparatus of this Echiniscoides sigismundi. The pharynx resembles a pair of kidneys in contact with the three filaments. The outer two filaments are the stylets; the inner one is the buccal tube. The gut contains algal or plant material that has been ingested by this tidal zone species. Photo by Martin Mach, with permission.

Figure 61. The land snail Bulimulus guadalupensis is a known predator on moss-dwelling tardigrades. Photo by Gary Rosenberg at .

Figure 59. The three "filaments" and oval behind them are the buccal apparatus of Paramacrobiotus [=Macrobiotus] areolatus. The bulbous oval to the right of the three filaments (stylets and buccal tube) is the pharynx. Photo by Martin Mach, with permission.

Tardigrades have smaller predators as well. The fungus Ballocephala pedicellata (Figure 63) is known from the tardigrades Hypsibius dujardini (Figure 64) and Diphascon pingue complex (Figure 65) living in the moss Atrichum angustatum (Figure 66) (Pohlad & Bernard 1978). In this study, tardigrades with the fungus were only

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Chapter 5-2: Tardigrade Reproduction and Food

present in January and February in the collecting area in southeastern USA.

Figure 65. Diphascon pingue. Photo by Michael Collins, with permission.

Figure 62. Armor of Echiniscus sp. Material such as this is easy to identify in the feces of predators. Photo by Martin Mach, with permission.

Figure 66. Atrichum angustatum, a habitat for tardigrades, and in winter, their parasitic fungus Ballocephala pedicellata. Photo by Michael Lüth, with permission.

Summary Figure 63. Ballocephala sphaerospora zygospores in a tardigrade. Photo by George Barron, with permission.

Figure 64. Hypsibius dujardini, a host for the fungus Ballocephala pedicellata. Photo from Rpgch Wikimedia Commons.

Life cycle stages often provide a means of surviving changes in the environment. Bryophytedwelling tardigrades are usually parthenogenetic. They rarely are hermaphrodites, and parthenogenetic individuals do not self-fertilize. Tardigrade species may either lay free eggs or deposit them inside the exuvia as it is shed. External eggs are usually highly decorated, whereas those laid inside the exuvia tend toward smooth. The number of eggs varies up to 40, with the number depending on the species and nutritional status. Time required for development of the fertilized egg may be up to 90 days. Variability in development time permits bet-hedging. Temperature affects development rate, survival rate, and body growth, as well as affecting generation time. Young tardigrades resemble the adults and continue to grow by cell enlargement. Molting permits the tardigrade to expand its size and requires 5-10 days during which the tardigrade cannot eat and is less protected. Some species have more than one morph, where the winter morph may form aggregations that enhance survival of freezing conditions.

Chapter 5-2: Tardigrade Reproduction and Food

Tardigrades consume algae, bryophytes, fungi, protozoa, nematodes, rotifers, and smaller tardigrades. In many cases this is accomplished using a stylet that forms a straw for sucking cell contents. They suck in their prey with the pair of stylets, with the muscular pharynx producing suction. For whatever reason, bryophyte dwellers also have larger excretory organs than do tardigrades of other substrata. They play an important role in regulating the food web of bryophytes in some circumstances. Tardigrades are subject to predation by snails and even larger tardigrades. Fungi may extract nutrition from them.

Acknowledgments Like all of my chapters, this one is really the product of the efforts of many biologists. Roberto Bertolani provided an invaluable update to the tardigrade taxonomic names and offered several suggestions on the text to provide clarification or correct errors. Bryonetters have been wonderful in making their photographs available to me and seeking photographs from others. Paul Davison has been helpful in providing suggestions and offering images. Martin Mach and Yuuji Tsukii have given permission to use images that illustrate the species and life cycle stages. Łukasz Kaczmarek has provided me with references and contact information. Claudio Delgadillo was kind enough to take the picture of Syntrichia obtusissima just for this chapter. Martin Mach was kind enough to send me corrections for typos in the previous online version. My sister, Eileen Dumire, read and edited an earlier version of the chapter for me from the viewpoint of a non-biologist. And a big thank you goes to Michael Lüth for permission to use his many images and to all those who have contributed their images to Wikimedia Commons for all to use. I fear I have forgotten some who have helped – I have worked on this chapter for too many years.

Literature Cited Altiero, T., Rebecchi, L., and Bertolani, R. 2006. Phenotypic variations in the life history of two clones of Macrobiotus richtersi (Eutardigrada, Macrobiotidae). Hydrobiologia 558: 33-40. Altiero, T., Bertolani, R., and Rebecchi, L. 2009. Hatching phenology and resting eggs in Paramacrobiotus richtersi (Eutardigrada, Macrobiotidae). J. Zool. (in press). Bartels, P. 2005. "Little known" water bears? ATBI Quart. 6(2): 4. Bertolani, R. 2001. Evolution of the reproductive mechanisms in tardigrades – a review. Zool. Anz. 240(3-4): 247-252. Bertolani, R., Altiero, T., and Nelson, D. R. 2009. Tardigrada (water bears). Chapter 188. In: Likens, G. E. (Ed.). Vol. 2. Encyclopedia of Inland Waters. Elsevier, Oxford, pp. 443465. Bertolani, R., Rebecchi, L., and Beccaccioli, G. 1990. Dispersal of Ramazzottius and other tardigrades in relation to type of reproduction. Invert. Repro. Dev. 18: 153-157. Dastych, H. 1993. Redescription of the cryoconital tardigrade Hypsibius klebelsbergi Mihelčič, 1959, with notes on the

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microslide collection of the late Dr. F. Mihelčič. Veröff. Mus. Ferdinandeum 73: 5-12. Davison, P. G. 2005. Feeding water bears: A simple activity to connect the public with microorganisms. ATBI Quarterly 6(2): 5. Fox, I. 1966. On the significance of tardigrades in the digestive tract of snails. Proc. First Internat. Congr. Parasitol. 2: 1064. Fox, I. and Garcia-Moll, I. 1962. Echiniscus molluscorum, new tardigrade from the feces of the land snail, Bulimulus exilis (Gmelin) in Puerto Rico (Tardigrada: Scutechiniscidae). J. Parasitol. 48(2): 177-181. Garey, J. R., McInnes, S. J., and Nichols, P. B. 2008. Global diversity of tardigrades (Tardigrada) in freshwater. Develop. Hydrobiol. 198: 101-106. Guidetti, R. and Bertolani, R. 2005. Tardigrade taxonomy: an updated check list of the taxa and a list of characters for their identification. Zootaxa 845: 1-46. Halberg, K. A., Persson, D., Ramløv, H., Westh, P., Møbjerg Kristensen, R., and Møbjerg, N. 2009. Cyclomorphosis in Tardigrada: Adaptation to environmental constraints. J. Exper. Biol. 212: 2803-2811. Hofmann, I. 1987. Habitat preference of the most frequent mossliving Tardigrada in the area of Giessen (Hessen). In: Bertolani, R. (ed.). Selected Symposia and Monographs U.Z.I., 1, Mucchi, Modena, pp. 211-216. Hohberg, K. 2006. Tardigrade species composition in young soils and some aspects on life history of Macrobiotus richtersi J. Murray, 1911. Pedobiologia 50: 267-274. Kinchin, I. M. 1988. The tardigrade fauna of moss cushions. SSR Science Notes June, 1988, pp. 733-737. Kinchin, I. M. 1994. The Biology of Tardigrades. Blackwell Publishing Co., London, 186 pp. Kristensen, R. 1982. The first record of cyclomorphosis in Tardigrada based on a new genus and species from Arctic meiobenthos. Z. Zool. Syst. Evol.-forsch 20: 249-270. Kristensen, Reinhardt Møbjerg and Sørensen, Martin Vinther. 2005. Tardigrada (Water Bears) Accessed on 16 January 2010 at . Lemloh, M.-L., Brümmer, F., and Schill, R. O. 2011. Lifehistory traits of the bisexual tardigrades Paramacrobiotus tonollii and Macrobiotus sapiens. J. Zool. Syst. Evol. Res. 49(suppl 1): 58-61. Mach, Martin. 2010. The Water Bear web base Still Images I. Overview. Accessed on 27 January 2010 at . Marcus, E. 1928. Spinnentiere oder Arachnoidea IV: Bärtierchen (Tardigrada). Springer, Jena. Miller, H. M. 1931. Alternation of generations in the rotifer Lecane inermis Bryce: I. Life histories of the sexual and non-sexual generations. Biol. Bull. 60: 345-381. Morgan, C. I. 1977. Population dynamics of two species of Tardigrada, Macrobiotus hufelandii (Schultze) and Echiniscus (Echiniscus) testudo (Doyère), in roof moss from Swansea. J. Anim. Ecol. 46: 263-279. Nelson, D. R. 1982. Developmental biology of the Tardigrada. In: Harrison, F. and Cowden, R. (eds.). Developmental Biology of Freshwater Invertebrates. Alan R. Liss, New York, pp. 363-368. Nelson, D. R. 2002. Current status of the Tardigrada: Evolution and ecology. Integrat. Compar. Biol. 42: 652-659. Pilato, G. 1979. Correlations between cryptobiosis and other biological characteristics in some soil animals. Boll. Zool. 46: 319-332.

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Pohlad, B. R. and Bernard, E. C. 1978. A new species of Entomophthorales parasitizing tardigrades. Mycologia 70: 130-139. Ramazzotti, G. and Maucci, W. 1983. The phylum Tardigrada – 3rd edition: English translation by C. W. Beasley. Mem. Ist. Ital. Idrobiol. Dott. Marco de Marchi 41: 1B680. Rebecchi, L. and Bertolani, R. 1994. Maturative pattern of ovary and testis in eutardigrades of freshwater and terrestrial habitats. Invert. Reprod. Dev. 26: 107-118. Sánchez-Moreno, S., Ferris, H., and Guil, N.. 2008. Role of tardigrades in the suppressive service of a soil food web. Agric. Ecosyst. Environ. 124: 187-192. Schill, R. O., Jönsson, K. I., Fannkuchen, M., and Brümmer, F. 2011. Food of tardigrades: a case study to understand food choice, intake and digestion. J. Zool. Syst. Evol. Res. 49(suppl 1): 66-70. Suren, A. M. 1988. Ecological role of bryophytes in high alpine streams of New Zealand. Internat. Ver. Theor. Angew. Limnol. 23: 1412-1416.

Suzuki, A. C. 2003. Life History of Milnesium tardigradum Doyère (Tardigrada) under a Rearing Environment. Zool. Sci. 20: 49-57. Tardigrada (Water Bears). 2005. Based on Animal Classification. Grzimek's Animal Life Encyclopedia, Vol. 2. The Gale Group, Inc. Accessed on 27 February 2010 at Answers.com . Tardigrada (Water Bears). 2005. Accessed 14 June 2010 at < http://www.answers.com/topic/tardigrada-water-bearsbiological-family>. Walz, B. 1982. Molting in Tardigrada. A review including new results on cuticle formation in Macrobiotus hufelandi. In: Nelson, D. R. (ed.). Proceedings of the Third International Symposium on the Tardigrada, August 3–6, 1980, Johnson City, Tennessee. East Tennessee State University Press, Johnson City, pp. 129-147. Węglarska, B. 1990. Morphology of excretory organs in Eutardigrada. Acta Biol. Cracow Ser. Zool. 31: 63-70.

Glime, J. M. 2017. Tardigrade Habitats. Chapt. 5-3. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. Ebook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 18 July 2020 and available at .

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CHAPTER 5-3 TARDIGRADE HABITATS

TABLE OF CONTENTS Bryophyte Habitats.............................................................................................................................................. 5-3-2 Specificity ........................................................................................................................................................... 5-3-3 Habitat Differences ............................................................................................................................................. 5-3-3 Acid or Alkaline? ......................................................................................................................................... 5-3-3 Altitude ........................................................................................................................................................ 5-3-4 Polar Bryophytes .......................................................................................................................................... 5-3-6 Forest Bryophytes ........................................................................................................................................ 5-3-9 Epiphytes ..................................................................................................................................................... 5-3-9 Aquatic ....................................................................................................................................................... 5-3-12 Dry Habitats ............................................................................................................................................... 5-3-14 Vertical and Horizontal Distribution .......................................................................................................... 5-3-14 Summary ........................................................................................................................................................... 5-3-16 Acknowledgments ............................................................................................................................................. 5-3-16 Literature Cited ................................................................................................................................................. 5-3-16

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Chapter 5-3: Tardigrade Habitats

CHAPTER 5-3 TARDIGRADE HABITATS

Figure 1. Echiniscus sp., member of a genus that is common on bryophytes. Photo by Martin Mach, with permission.

Bryophyte Habitats Tardigrades exist in both aquatic and terrestrial habitats worldwide, and in both cases can be found with bryophytes (Figure 1) (and lichens and leaf litter) (Utsugi et al. 1997). The tropics seem unfavorable (Mathews 1938), perhaps for the same reasons that temperate stream bryophytes are uncommon in lowland tropical waters – they are warm and wet at the same time, encouraging bacterial and fungal growth. Most of the more than 900 known tardigrade species are limnoterrestrial (Garey et al 2008). That is, they live in a thin surface film of water, most commonly on bryophytes, lichens, algae, and other plants. They can only remain active while this film of water exists. Of the 910 species reviewed by Garey et al. (2008), only 62 species, in 13 genera, are truly aquatic and unknown from limnoterrestrial habitats. Nevertheless, many of the limnoterrestrial species can at least occasionally be found in freshwater. In these terrestrial habitats, pH of the substrate, oxygen tension, moisture content of the bryophyte, thickness of the bryophyte mat or cushion, and altitude (and its attendant conditions) all contribute to the habitat distribution. I have taken the liberty of changing the word "moss," used in many tardigrade studies, to "bryophyte." I have

learned from one of my kind tardigrade reviewers that people who study tardigrades often do not understand leafy liverworts and lump them into mosses. Hence, unless I could determine that the researcher definitely had in mind only mosses (and not also liverworts), I used the term bryophytes. I also learned that many ecologists include mosses in the category of "soil"! Others include them in "litter." While this lumping can be a useful concept for some aspects of functional ecology, one needs to be aware of it when searching for bryological literature or interpreting soil literature. Tardigrades are especially common on tree bark bryophytes (epiphytes), presumably due to having similar tolerances to drying (Crum 1976). They are known from all seven continents and up to 6600 m altitude in the Himalayas (Ehrenberg 1859 in Fontoura et al. 2009). Collins and Bateman (2001) examined factors affecting tardigrade distribution in Newfoundland and learned that in this case altitude and type of bedrock were important in determining tardigrade distribution. Moisture and rate of drying further defined their distribution. And in some cases, competitive exclusion or interspecific competition seemed to be determining factors for community composition.

Chapter 5-3: Tardigrade Habitats

Specificity Species assemblages seem to be similar throughout the world. Is this due to lack of taxonomic understanding or to widespread dispersal? In the following sections we will examine what we know about factors affecting tardigrade communities under different circumstances. It appears that many tardigrades have little preference for bryophytes versus lichens (Meyer & Hinton 2007). But even those cryptogams, supporting their wet-dry requirements, are usually not unique habitats for the tardigrades, with the same species of bryophytes and lichens also present in soil, leaf litter, and additional habitats. Several studies have attempted to show any species preferences for bryophytes, but typically with no success (Kathman & Cross 1991; Miller & Heatwole 1995; Meyer & Hinton 2007). Further evidence of nonspecificity is in their distribution. Meyer and Hinton (2007) report that the Nearctic realm shares 82 species of tardigrade with the Neotropical realm. Everything is everywhere! On the other hand, 30% of the Nearctic species are known from only one site. One of the problems in describing the tardigrade habitat is that substrate records are inconsistent or absent for many collections. But some studies have indicated that lichens and mosses may be preferred over other substrata. Working in China, Beasley et al. (2006) found 18 species of tardigrades from three provinces, primarily in lichens and bryophytes. One of the most common tardigrades among bryophytes is Milnesium tardigradum (Figure 21) and the less common Macrobiotus hibiscus. Hinton and Meyer (2008) reported these among liverworts (Jungermannia sp.; Figure 2) in a suburban lawn in central Georgia (USA).

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apparently cosmopolitan. Hinton and Meyer suggested that Biserovus bindae, Echiniscus cavagnaroi, Echiniscus kofordi, Hexapodibius christenberryae, Minibiotus fallax, and Parhexapodibius pilatoi, along with a new variant of Macrobiotus cf. hufelandi, may represent a distinctive southeastern USA regional fauna living among cryptogams. On the other hand, Echiniscus cavagnaroi and Echiniscus kofordi are known only in the Neotropical Region and in the Galapagos Islands, but Minibiotus fallax was described in Australia. These three species are frequently encountered in mosses and lichens.

Habitat Differences Several tardigrade researchers have considered five types of bryophyte habitats (Mihelčič 1954/55, 1963; Ramazzotti 1962; Hofmann 1987): • • • •

bryophytes that are submerged bryophytes that are permanently moist bryophytes growing in shady places and rarely dry bryophytes that dry out frequently and receive direct sunlight regularly • bryophytes that are extremely exposed and often dry for a longer period. Ito (1999) was able to identify six groupings of tardigrades, based on habitat preference, in his altitudinal study on Mt. Fuji, Japan. Kaczmarek et al. (2011) likewise found altitudinal relationships in Costa Rica. They furthermore found a higher diversity among mosses than among lichens or liverworts, although they admitted to a possible bias due to unequal sampling. Tardigrades from these substrates were most common from 2000 to 2400 m asl and above 3200 m asl. Tardigrades do not have much control over their dispersal, typically depending on dispersal of the substrate. This may help to explain the observations on two morphotypes of the moss Grimmia. In this case, the tardigrade distribution was very patchy. There were no differences in distribution patterns on the two Grimmia morphotypes, despite their representation of different moisture conditions (Bettis 2008). Bettis suggested that the greater rainfall during the winter of observation might account for the lack of difference. But tardigrades are well known for their great tolerance of extremes, so their greatest limitation may be dispersal. Acid or Alkaline?

Figure 2. Jungermannia atrovirens, member of a genus where tardigrades have been found in a lawn in Georgia, USA. Photo by Michael Lüth, with permission.

Liverworts are rarely mentioned in tardigrade studies. However, Hinton and Meyer (2007) reported Echiniscus virginicus and Milnesium tardigradum from liverworts. In their study, they collected handfuls of mosses, liverworts, and lichens from 54 parishes in Louisiana, USA. They found 51 species in the region: 19 in Texas, 16 in Louisiana, 10 in Mississippi, 33 in Alabama, 3 in Georgia, and 15 in Florida. Of these 51 cryptogam dwellers, 20 are widely distributed in the region and 18 are

Bartels (2005) reported greater diversity in limestone habitats than elsewhere. It appears that acid can be an uncomfortable or lethal milieu. Hypsibius dujardini (Figure 13) had reduced activity after only five minutes at pH 3 and died at pH 2.8. Even at pH 4.0, it had reduced activity after 30 hours. But in Giessen, Germany, Hofmann (1987) found a somewhat different relationship. The four most abundant species [Macrobiotus hufelandi (Figure 7), Ramazzottius (formerly in Hypsibius) oberhaeuseri (Figure 3), Milnesium tardigradum (Figure 21), and Echiniscus testudo (Figure 4)] had similar preferences for alkaline substrata, but the remaining species, as a group, had a preference for the acidic habitats, thus presenting greater tardigrade species diversity among the acid substrata and the mosses that inhabited them.

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Chapter 5-3: Tardigrade Habitats

Figure 3. Ramazzottius oberhaeuseri, a bryophyte dweller that cannot tolerate constant moisture. Photo by Martin Mach, with permission.

Hingley (1993) found only two genera in her acid peatland studies: Diphascon (D. scoticum) (Figure 29) and Macrobiotus (Figure 6). To that Pilato (2009) added Bindius triquetrus from Sphagnum (Figure 5) in Sicily. Distribution is patchy (Romano et al. 2001), requiring greater sampling effort. One must wonder, is the paucity of reports of aquatic tardigrades on bryophytes a realistic representation of a meager aquatic fauna, or are there simply too few studies that have looked for them?

Figure 4. Echiniscus testudo tun on a bryophyte leaf. Photo by Power & Syred, through Creative Commons.

altitudinal groupings (lowland, upland, montane, etc.) (Ramazzotti & Maucci 1983; Dastych 1987, 1988). Collins and Bateman (2001) found that in Newfoundland, Canada, the lowland class could be further divided into locations up to100 m and those above (101-200 m). Table 1 compares the altitudinal abundance of 45 species of tardigrades associated with bryophytes on mountains in British Columbia, Canada (~48-60ºN), with those of riparian epiphytes (inhabiting trees on banks of natural water courses) in Alabama, USA (~33ºN). Although the latitudes are quite different, six species are common to both), but six species differ. The three most abundant Alabama species were common to both, but the very common Macrobiotus hufelandi (Figure 7) was absent in the Alabama collections. These data suggest that there may be more than just chance determining the species and abundance differences. But not all altitudinal studies have supported these conclusions (e.g. Kathman & Cross 1991). It is interesting that Meininger and Spatt (1988) found that altitude was not influential in determining distribution and abundance of moss-dwelling tardigrades in Alaska, USA. Likewise, Guil et al. (2009) found that the altitudinal differences could be explained by differences in soil, climate, vegetation structure, and litter type. Nevertheless, Ramazzotti and Maucci (1983) considered certain species to occur only above 500 m. This may simply be a lack of sufficient collecting – they claimed that Macrobiotus harmsworthi (Figure 6) was one of these "montane" species, but Dastych (1985) later reported it from locations between 0 and 1100 m altitude on Spitsbergen Island, Norway. Furthermore, Dastych (1980, 1988), showed a large correlation between Tardigrada species and altitude in the Tatra Mountains in Poland. Certainly latitude must be considered in making comparisons of altitude. And local moisture regimes are likely to play a major role in altitudinal relationships.

Figure 6. Macrobiotus harmsworthi, a common tardigrade on bryophytes and elsewhere. Photo by Paul J. Bartel, with permissions. Figure 5. Sphagnum fuscum, a species that forms hummocks where a tardigrade could find moisture but usually avoid being flooded. Photo by Michael Lüth, with permission.

Altitude Many researchers have shown a relationship between altitude and the distribution of tardigrades (RodriguezRoda 1951; Nelson 1973, 1975; Ramazzotti & Maucci 1983; Dastych 1985, 1987, 1988; Beasley 1988), suggesting that species richness increases with altitude. Bertolani and Rebecchi (1996) found that some species were typical of high altitudes or latitudes. Some researchers have even classified the tardigrades based on

Figure 7. Macrobiotus hufelandi, one of the most abundant tardigrades on bryophytes. Photo by Paul J. Bartels, with permission.

Chapter 5-3: Tardigrade Habitats

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Table 1. Altitudinal distribution of numbers of tardigrades in eleven bryophyte samples each, from six altitudes on five mountains on Vancouver Island, British Columbia, Canada, from Kathman & Cross 1991, and from 108 riparian epiphytic bryophyte samples representing 6 sites at Choccolocco Creek, Alabama, USA, from Romano et al. 2001. Those highlighted in grey occur in both sites.

Astatumen trinacriae Bertolanius [=Amphibolus] weglarskae Calohypsibius ornatus Diphascon [=Hypsibius] scoticum Diphascon belgicae Diphascon iltisi Diphascon modestum Diphascon nodulosum Diphascon pingue sl Diphascon prorsirostre Diphascon recamieri Echiniscus cf. arctomys Echiniscus horningi Echiniscus mauccii Echiniscus quadrispinosus Echiniscus sp. n. Echiniscus wendti Hypechiniscus gladiator Hypsibius convergens Hypsibius dujardini Insuetifurca arrowsmithi Isohypsibius lunulatus Isohypsibius sattleri Isohypsibius woodsae Itaquascon pawlowskii Macrobiotus crenulatus Macrobiotus echinogenitus (Figure 8) Macrobiotus harmsworthi Macrobiotus hufelandi Macrobiotus islandicus Macrobiotus lazzaroi Macrobiotus sp. A Mesocrista spitzbergensis Milnesium tardigradum Minibiotus cf. intermedius Minibiotus intermedius Murrayon hibernicus Paramacrobiotus [=Macrobiotus] areolatus Paramacrobiotus[=Macrobiotus] richtersi Platicrista cheleusis Pseudechiniscus goedeni Pseudechiniscus juanitae Ramazzottius baumanni Ramazzottius oberhaeuseri Testechiniscus laterculus SUM OF INDIVIDUALS NUMBER OF SPECIES

Altitude (m) 750 1050

150

450

1350

>1525

0 0 22 1 0 12 14 70 4 49

0 3 18 0 1 0 4 318 38 47

0 10 11 0 0 1 1 45 16 2

0 4 13 6 0 4 16 7 3 3

0 2 30 2 0 0 17 40 8 13

2 3 16 1 0 0 26 27 5 1

2 6 0

1 3 0

1 0 2

3 3 14

3 1 4

5 7 1

2 0 199

3 0 203

3 0 188

0 0 78

38 0 54

3 0 26

0 6 96 0 1 1 0 177 3039

0 9 49 0 0 0 0 459 1710

40 0 28 0 0 0 10 284 2061

0 0 4 2 2 0 79 44 1116

1 0 8 0 0 0 48 8 1586

0 0 0 0 1 0 0 10 662

10 1 5 21

0 0 1 24

0 0 0 2

0 0 2 0

0 0 2 2

0 0 0 4

2 0 31 0 8 0 0 18 11 0

1 0 16 0 1 5 0 44 2 0

12 0 0 0 2 0 3 8 0 0

3 14 0 0 13 0 0 7 1 0

0 1 0 0 10 0 0 5 1 39

0 0 0 1 13 2 0 3 0 0

3808 27

2960 23

2730 21

1421 24

1923 24

819 22

riparian 3

28 16 24

1

737 1

87 27 476 4 44

1448 12

Figure 8. Macrobiotus echinogenitus, a tardigrade living on riparian bryophytes at Choccolocco Creek Alabama, USA. Photo by Paul J. Bartels, with permission.

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Chapter 5-3: Tardigrade Habitats

Using PCA analysis, Kathman and Cross (1991) also reported no relationship between altitude and abundance on Vancouver Island, British Columbia, on the western coast of Canada (Table 1 suggests a decreasing trend in number of individuals might be present). Collins and Bateman (2001) later reported that altitude was one of the major determining factors in tardigrade distribution in Newfoundland, eastern coast of Canada. Rodriguez-Roda (1951 in Kathman & Cross 1991) found that altitude had a distinct effect on the abundance of tardigrades in Spain, with numbers increasing with altitude and reaching a maximum between 1000 and 2000 m. Dastych (1980) likewise found that tardigrades in the Tatra Mountains of Poland increased with altitude, again with the maximum numbers between 1000 and 2000 m. In one of his later studies, Dastych (1985) reported a seemingly opposite effect, demonstrating that the number of species and individuals decreased with increasing altitude in West Spitsbergen, Norway. The differences between studies may be a matter of scale and the fact that only some species are affected by altitude within the study range, but moisture regimes are likely to differ as well. Nelson (1973, 1975) found that only seven of the 21 bryophyte-dwelling species on Roan Mountain, Tennessee, USA, were affected by altitude. Bertrand (1975) found three altitudinal groups in the Aigoual Mountains of France. Beasley (1988) divided the tardigrades from 10523567 m in New Mexico, USA, into altitudinal ranges. Ito (1999) examined tardigrades at 17 stations on Mt. Fuji, Japan, finding little distributional pattern related to altitude (950-2380 m). Rather, the distribution related to habitat. Collins and Bateman (2001) found that tardigrades of Newfoundland, Canada, were affected by both altitude (Table 2) and type of bedrock, but that moisture content and rate of desiccation of the mosses and lichens where they live also contributed to their distributional pattern. Table 2. Decline in number of species with increasing altitude in Newfoundland. From Collins and Bateman 2001.

Altitude

Total number of species found at each altitude

0->100 m 101->200 m 200+ m

28 15 8

Mean number of tardigrade species at each site 2.75 1.75 1.88

Differences in techniques, lack of or differences in statistical analyses to support purported differences, and misidentification could contribute to the apparent differences in relationships among these studies, but moisture regimes most likely play a major role. In some cases, competitive exclusion appears to play a role (Collins & Bateman 2001). Nevertheless, it is likely that the effect of altitude, if it exists, depends in part on both latitude and scale. Polar Bryophytes Because of their relative abundance, and the predominance of mosses and lichens in the Antarctic flora,

we have learned some interesting aspects of their faunal ecology and physiology there. As early as 1976, 23 species of tardigrades were known from Antarctica (Jennings 1976). That's not bad for that early date in a place with limited vegetation, harsh climate, and limited opportunity for collecting, not to mention the distance to be travelled for colonization. Most of these tardigrade species have worldwide distribution (Venkataraman 1998). In the short Antarctic summers, the tardigrades multiply quickly, using parthenogenesis. Unlike most habitats elsewhere, the tardigrades in the Antarctic moss turf do exhibit a vertical zonation pattern. Schwarz et al. (1993) found that protozoa, rotifers, nematodes, and tardigrades dominated the moss-dominated flushes at Canada Glacier, southern Victoria Land, Antarctica. Mites were of less importance. These invertebrates occurred in the range of 5 to 10.83 mm depth in the moss clumps. Post melt samples had a greater percentage of all groups of organisms in the upper 5 mm of mosses compared to those at that depth range in the premelt samples, suggesting either migration or rapid reproduction once melting occurred. Venkataraman (1998), in clumps of Bryum argenteum (Figure 9) from continental Antarctica, found that the tardigrades only live down to 15 cm depth in the 30 cm deep turfs. If they prefer to eat rotifers, they can only find those down to 10 cm. Their slow movement could limit the distance they can reasonably move downward and still return for food in a timely manner. Ramazzotti (1972) estimated that tardigrades could travel an average of 17.7 cm h-1, seemingly enough speed to travel another 10 cm, but perhaps not in the cold and not during the daytime feeding period when downward movement would be most beneficial to avoid drying. Temperature may play a role in the zonation of these Antarctic bryophytes. Bryophyte temperatures in the Antarctic can differ considerably from those of the air and may provide a warm refuge for activity even on cold days. Bryophytes exhibit a sharp temperature zonation on sunny days when there is no snow cover (Jennings 1979). The surface is subject to evaporative cooling in the polar winds while the moss layer immediately below that interface is quiet and often dark in color, absorbing the heat like a black body, as seen by the temperatures shown in Figure 10. Hence, in summer the moss turf has temperatures much higher than that of the air and at the beginning and end of the summer season the temperatures fluctuate around freezing for a considerable time, even if the mosses are snow-covered. Sohlenius and Boström (2006) described tardigrade communities from moss cushions on four ice-free mountains (nunataks) in Antarctica. Tardigrades occurred in 32% of the 91 samples of mosses. No invertebrates at all occurred in 8% of the samples. They considered stochastic processes (random events) accounted for the uneven colonization of the moss cushions, most likely as a result of random dispersal. They suggested that nematodes seem to offer competition, whereas the rotifers seem to provide a food choice.

Chapter 5-3: Tardigrade Habitats

Figure 9. Bryum sp. cushions in Antarctica, demonstrating the deep turfs that can house tardigrades. Photo courtesy of Catherine Beard.

Tardigrades manage to survive the extreme cold of the long Antarctic winter as a tun (see Chapter 4-6). But other physiological adaptations are needed to combat the extremes of temperature that can be experienced in a single Antarctic day (see for example Figure 10). Both Bertolanius nebulosus and Richtersius coronifer (Figure 11) endure ice formation as they proceed through the onset of freezing temperatures (Westh & Kristensen 1992). Both are able to supercool to -6 or -7ºC before they succumb to freezing. These two tardigrades are common in Polar areas, as well as elsewhere. Richtersius coronifer (Figure 11) spends its Arctic winters in drought-resistant mosses as a frozen or dry individual. Bertolanius nebulosus has adopted a somewhat different strategy, spending its cold period in moist mosses and algae as a frozen cyst, or occasionally as an egg or adult.

Figure 10. Comparison of moss surface temperature with that of 1.5 cm depth in moss cover on Signy Island in the Antarctic on three days in February, 1973. Redrawn from Jennings 1979.

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Figure 11. Richtersius coronifer, a tardigrade that survives Arctic winters on mosses in a frozen state. Photo by Martin Mach, with permission.

It appears that adapted tardigrades can survive extreme conditions for a long time. Newsham et al. (2006) experimented in a way that might be considered cruelty to animals (but not unlike Mother Nature herself). They partially dried a bit of the leafy liverwort Cephaloziella varians from Rothera Point on the Wright Peninsula, Adelaide Island, western Antarctic Peninsula, then stored it frozen at -80ºC for six years and two months. They then rapidly thawed the liverwort at 10ºC. You guessed it! Tardigrades survived, along with nematodes and a bdelloid rotifer. Only two individuals [Diphascon sp. (see Figure 12), Hypsibius cf. dujardini (Figure 13)] out of fifteen tardigrades (13%) made it, but that is still remarkable! The eleven individuals of Macrobiotus furciger (Figure 14) and one of Echiniscus sp. did not. Nematodes fared a bit better, with 31% survival out of 159 individuals.

Figure 12. Diphascon scoticum, a moss-dwelling representative of a genus in which one member survived storage at -80ºC for six years! Photo by Łukasz Kaczmarek, with permission.

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Chapter 5-3: Tardigrade Habitats

Figure 13. Hypsibius dujardini, a species that survived -80ºC for six years. Photo by Rpgch, through Wikimedia Commons.

Figure 14. Macrobiotus furciger. Photo by Smithsonian Institution, through EOL Creative Commons.

Sømme and Terje Meier (1995) examined Echiniscus jenningsi (Figure 15), Macrobiotus furciger (Figure 14), and Diphascon chilenense from Müihlig-Hofmannfjella, Dronning Maud Land, Antarctica to ask similar winter survival questions. They compared both hydrated and dehydrated individuals. After 600 days at -22ºC, both hydrated and dehydrated tardigrades had high survival rates. After 3040 days, the dry individuals still had a high rate of survival. However, at -80ºC hydrated Echiniscus jenningsi (Figure 15) did not fare as well as the others, decreasing in survivorship as time increased from 7 to 150 days. At -180ºC, all hydrated individuals of these three species rapidly died, but all dehydrated species had good survivorship after 14 days at -180ºC.

Figure 15. Echiniscus jenningsi. Photo by Smithsonian Institution, through EOL Creative Commons.

It is incredible that some Antarctic tardigrades can survive temperatures as low as -80ºC in a hydrated state (Sømme & Meier 1995; Sømme, 1996)! The ability to survive short periods in a hydrated condition just below a freezing temperature is important to survival in bryophyte clumps that regularly warm in the day and freeze at night. Ice crystals on the bryophytes would most likely help to dehydrate the tardigrades as temperatures plummeted to well below freezing, facilitating their survival during the long and very cold winters. Furthermore, the ability to survive low temperatures for years would permit tardigrades to survive dispersal across the ice or remain viable within it until a suitable habitat or conditions are reached. But how do the rapidly changing temperatures of the environment affect the ability of the tardigrade to move about on the bryophyte to seek food? The beneficial acclimation hypothesis (BAH) predicts that animals will have their best performance at the temperature to which they are acclimated. Li and Wang (2005) tested this hypothesis with the moss-dwelling species Macrobiotus hufelandi (Figure 7, Figure 19), collected from the Qinling Mountains in central China. They acclimated the water bears to 2 and 22ºC for two weeks. Using walking speed and percentage of time moving, they compared performance at the acclimation temperature with that at the alternate temperature. They found that both walking speed and percentage of time moving was significantly faster when the tardigrades were kept at their acclimation temperature than when they were placed at the higher or lower experimental temperature. But in the Antarctic, we have seen that such extreme temperature fluctuations within a single day are not unusual. Could this be a threat to the water bears, who must find food, often adhering bacteria and algae, on the moss? And others eat nematodes and other moving targets. One factor to consider is that in the experiments of Li and Wang, only 1.5 minutes acclimation were provided at the new temperature before measurements began, lasting another 3-5 minutes. This seems unrealistic as a representation of nature. The next question to ask is how fast can the tardigrades acclimate to a new temperature? Danger may lurk among the Antarctic bryophytes. Gray et al. (1982) isolated eighteen taxa of predaceous fungi from among Antarctic mosses and soil samples. Among these eight different trapping mechanisms were present. The fungus Monacrosporium ellipsosporum seemed to be associated primarily with calcicolous mosses. Although these are nematode-trapping fungi, they may also catch the occasional tardigrade. More importantly, it indicates that the moss habitat is suitable for parasitic fungi that might attack other invertebrate groups such as tardigrades. Miller et al. (1996) actually looked at the role of bryophytes vs other cryptogamic substrata in harboring tardigrades on the Windmill Islands in East Antarctica. Pseudechiniscus suillus (Figure 16), Macrobiotus sp. (see Figure 7, Figure 19), Hypsibius antarcticus, Ramajendas frigidus, Diphascon chilenense, and Diphascon pingue (Figure 17) occurred among mosses and lichens. Three of these had positive associations with each other and with bryophytes: Pseudechiniscus suillus, Hypsibius

Chapter 5-3: Tardigrade Habitats

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antarcticus, and Diphascon chilenense. These three species had a strong negative association with algae and lichens.

Figure 18. Platicrista angustata. Photo by Michael Collins, with permission.

Figure 16. Pseudechiniscus suillus group, an Antarctic bryophyte dweller that avoids lichens. Photo by P. J. Bartels, with permission.

Figure 19. Macrobiotus hufelandi. Photo by Martin Mach, with permission.

Figure 17. Diphascon pingue, a moss and lichen dweller Swedish forests and in the Antarctic. Photo by Michael Collins, with permission.

Forest Bryophytes Forests have a much more tempered climate than the Antarctic. Trees reduce the rate of water loss and shade the bryophytes and their fauna from the heat of the bright sun. Jönsson (2003) examined bryophytes in Swedish forests and found sixteen species of tardigrades, five of which were previously undescribed for that region [Murrayon dianeae (Figure 36), Isohypsibius sattleri, Platicrista angustata (Figure 18), Diphascon belgicae, D. pingue (Figure 17)]. Jönsson found that the pine forest had the most species compared to clearcut areas, but that abundance differed little from that of clearcut areas. Of the sixteen species of tardigrades recorded, the cosmopolitan Macrobiotus hufelandi (Figure 19) was by far the most abundant. The weft growth form seemed to harbor more tardigrades than did other bryophyte growth forms.

Schuster and Greven (2007) conducted a 54-month study of the tardigrade fauna of the moss Rhytidiadelphus squarrosus in the Black Forest of Germany. They found 24 species, dominated by Macrobiotus hufelandi (56%), M. richtersi (18%), and Diphascon pingue (12%). Diversity tended to be higher in winter, but the three dominant species generally declined in winter and increased from spring until fall. Rainfall, humidity. and temperature seemed to play a major role in changes in seasonal abundance. Epiphytes Whereas forest floor bryophyte dwellers are protected by snow in winter, bryophytes on trees (epiphytes) are often above the winter snow level. In summer they have intermittent wet and dry periods and in winter they often have exposures to extreme temperatures, lacking the protective cover of snow. In the Cincinnati, Ohio, USA area, bark-inhabiting bryophytes provide homes to numerous tardigrades, with the greatest species richness in environs of high humidity and clean air (Meininger et al. 1985). Hence, cities afford a less hospitable environment due to the lower humidity and decreased air quality.

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Chapter 5-3: Tardigrade Habitats

Despite their seemingly unfriendly habitat, epiphytic bryophytes are particularly suitable as a tardigrade habitat. Indeed, this habitat seems to house the most species. The frequency of wetting and drying of these bryophytes seems to be most suitable to the tardigrade life cycle. Richness seems to run about 4-16 species. Hooie and Davison (2001; Hooie Tardigrade diversity) found the following tardigrades associated with the epiphytic moss Ulota crispa (Figure 20) on four tree species (Acer saccharum, Acer rubrum, Betula lenta, Tilia americana) in the Great Smoky Mountains National Park, USA: Echiniscus cf. oihonnae Echiniscus virginicus Hypechiniscus gladiator Macrobiotus hufelandi (Figure 19) Milnesium tardigradum (Figure 21) Minibiotus cf. pustulatus Paramacrobiotus tonollii (Figure 23) Pseudechiniscus (Figure 22) Figure 21. Milnesium tardigradum. A cosmopolitan bryophyte inhabitant. Photo by Martin Mach, with permission.

Figure 22. Pseudechiniscus juanitae. Bartels, with permission.

Photo by Paul J.

Figure 20. Ulota crispa, an epiphytic moss that houses a number of tardigrade species. Photo by Michael Lüth, with permission.

In a study of riparian tardigrades, Romano et al. (2001) reported on the tardigrades on epiphytic bryophytes in Alabama, USA. Using 108 samples, they extracted 1588 tardigrades from three tree species on six sampling dates. Like Riggin (1962) for forest bryophytes, Romano et al. (2001) found that Macrobiotus species (Figure 6-Figure 8) were the most abundant (1358 of the 1588 tardigrades, 86%). They found no differences among tree species, bryophyte species, or seasons, but there were site differences, possibly suggesting dispersal limitations. Although relative humidity and temperature did not seem to influence abundance, precipitation did. Interestingly, as precipitation increased, the number of tardigrades decreased. Beasley (1981) found that higher humidity resulted in lower tardigrade abundance in the Caribbean National Rain Forest at Luquillo, Puerto Rico. This further supports the hypothesis that periods of anhydrobiosis are required in the life of a tardigrade and that lack of them shorten the length of life. However, if only active periods are considered, there may be little difference.

Figure 23. Paramacrobiotus tonollii, a tardigrade known from Ulota crispa in the Smoky Mountains. Photo by Martin Mach, with permission.

Briones et al. (1997) suggested that during periods of high precipitation the film of water surrounding the bryophytes may become anoxic, killing the tardigrades. This could especially be a problem in the riparian zone, where the bryophytes, and hence the tardigrades, were under water during several collection periods. Diversity of tardigrades was somewhat low in the Alabama, USA,

Chapter 5-3: Tardigrade Habitats

riparian sites (Table 1), with only twelve species overall (Romano et al. 2001). Mosses included Anomodon (Figure 24), Leucodon (Figure 25), and Schwetschkeopsis (Figure 26), all epiphytes.

Figure 24. Anomodon rugelii, an epiphytic moss. Photo by Michael Lüth, with permission.

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As in the Alabama study, Jerez Jaimes (2002) found a low diversity of only seven species on the moss Calymperes palisotii (Figure 27) on six species of trees on the campus of the University of Puerto Rico at Mayagüez. The highest species richness was on Mangifera indica (mango) and Swietenia macrophylla (mahogany). When collections came from trees and shrubs from all 67 counties in Florida, only 20 species of tardigrades were found (Meyer 2006, 2008). Meyer also found no specificity for a particular bryophyte (47 species included) or ecological region, but there was specificity for bryophytes as opposed to foliose lichens. There seemed to be an avoidance of one moss species, Aulacomnium heterostichum (Figure 28), with no tardigrades occurring on it (Meyer 2008). Bartels and Nelson (2006), working in the Great Smoky Mountains National Park, USA, increased the number of known species in the park from three to 42 from multiple substrates, a further testimony to how little known these organisms are. Bartels and Nelson found that more tardigrades occurred in bryophytes at breast height on a tree than at the tree bases, perhaps again relating to longer or more frequent dry periods. Diphascon [=Hypsibius] scoticum (Figure 29), a very common tardigrade, inhabits mosses on logs (Cushman, pers. comm. 1970). It would be interesting to compare the log-dwelling tardigrade taxa with those living on epiphytic bryophytes of the same species. Presumably, the log habitat would have longer moist periods. Based on the findings discussed above of Bartels and Nelson (2006) and Romano et al. (2001), one might expect more on the epiphytes, where alternating wet and dry periods might fit better with the apparent dormancy requirements of the tardigrades.

Figure 25. Leucodon sciuroides, an epiphytic moss that compresses and curls its branches upward when it dries. Photo by Michael Lüth, with permission.

Figure 26. Schwetskeopsis fabronia , an epiphyte from Asia and North America, and home for tardigrades. Photo by Misha Ignatov, with permission.

Figure 27. Calymperes palisotii, a moss that had the lowest tardigrade diversity on the University of Puerto Rico campus, Mayagüez. Photo by Claudio Delgadillo Moya, with permission.

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Chapter 5-3: Tardigrade Habitats

Figure 28. Aulacomnium heterostichum, a moss that seems to be avoided by tardigrades. Photo by Janice Glime.

Figure 30. Dactylobiotus [=Macrobiotus] dispar. Photo by Martin Mach, with permission.

Figure 29. Diphascon scoticum, a very common tardigrade, one that has been found among mosses on logs. Photo by Paul J. Bartels, with permission.

Figure 31. Dactylobiotus [=Macrobiotus] dispar. Photo by Martin Mach, with permission.

Aquatic Few truly aquatic tardigrades seem to exist. Garey et al. (2008) reported that of the more than 900 species of tardigrades they reviewed, most live in the water film on the surface of bryophytes, lichens, algae, and other photosynthetic organisms. Of their 910 species of tardigrades, only 62 (in 13 genera) were truly aquatic. In New Zealand, tardigrades represented only 2.4% of the fauna among the bryophytes of an unshaded stream in the alpine zone on South Island (Suren 1991a). In a more extensive study there, they represented only 1.2% of the invertebrates collected among mosses in 103 streams in one study (Suren 1993). Similar small numbers were present among the moss Fontinalis antipyretica (Figure 34) in the Czech Republic, where one stream had 1541 per mL of moss (0.6% of the fauna in the moss) and another stream had only 545 per mL (0.1%) (Vlčková et al. 2001/2002). In the Colorado Rocky Mountains, Elgmork and Sæther (1970) found most of the tardigrades Dactylobiotus cf macronyx (formerly Macrobiotus macronyx) associated with algae, but they also reported them from Fontinalis beds (Figure 34) and other submerged mosses. In streams, flow velocity may cause tardigrades to seek refuge among mosses. Suren (1992) reported high densities of tardigrades (Dactylobiotus [=Macrobiotus] dispar; Figure 30-Figure 31) associated with mosses in alpine streams of New Zealand. In an earlier study Suren (1991b) found that the colonization of this species on artificial mosses was at a reduced density compared to that on mosses [Fissidens rigidulus (Figure 32), Cratoneuropsis relaxa, Bryum blandum (Figure 33)].

Figure 32. Fissidens rigidulus from New Zealand, a good tardigrade habitat. Photo by Bill and Nancy Malcolm, with permission.

Figure 33. Bryum blandum from New Zealand, where tardigrade density is greater than that on artificial mosses. Photo by Jan-Peter Frahm, with permission.

Chapter 5-3: Tardigrade Habitats

Suren (1992) reported densities of 3120 and 8160 per m2 on the mosses in two trials, whereas they reached only 1760 and 1600 on the artificial substrata. He suggested that the high periphyton biomass among mosses provided a good food source that made this a good habitat for the tardigrades. This suggestion is supported by the largest percentage of variation (24.2%) being explained by the ultra-fine particulate matter (UFPOM). The abundance of tardigrades on bryophytes was 10 times that found on stream gravel. Linhart et al. (2002) examined scattered clumps of the aquatic moss Fontinalis antipyretica (Figure 34) and found that whereas several groups of invertebrates were distributed among the clumps in relation to stream flow, this was not the case for tardigrades. Even though the researchers showed that fine organic matter trapped within the moss mat was determined by flow velocity, this did not seem to be a determining factor in tardigrade distribution.

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Suren (1992) also considered the possibility that the mosses offered shelter from the rapid flow of water elsewhere. These tiny organisms are often in the leaf axils of the mosses, where they have almost no effect from the strong flow, yet the oxygen diffusion could be fairly rapid. But as yet, no data seem to support cause and effect of flow velocity and tardigrade distribution. Living in a stream is challenging for a tardigrade. Using 22 animals, Shcherbako et al. (2010) found that Milnesium tardigradum (Figure 21) could manage in water with a mean velocity of 23.3 mm/h, moving at a mean speed of 19.8 mm/h in the light and 29 mm/h in the dark, making snails look like track stars! Bryophytes provide a safe refuge from fast-moving waters. Eles and Repas (2009) described the stream tardigrades as having faster motion and longer claws than their terrestrial counterparts. In New Zealand, Suren (1992) found that the tardigrade Dactylobiotus dispar (Figure 31) represented about 6.6% of the fauna on mosses in unshaded streams compared to 0.6% on gravel. In shaded streams they occupied only 5.3% of the bryophyte fauna compared to 0.4% on gravel. But not all wet habitats seem to be very suitable for tardigrades. Kaczmarek (pers. comm. 29 January 2010) has reminded me that most of the water-dwelling tardigrades are in fact marine. Those that are truly freshwater aquatic species live on algae or plants (including bryophytes), in the sand, or in sediments. The genus Murrayon (Figure 36) is unusual among the water-dwelling tardigrades in that some aquatic individuals lay their eggs in the shed shells of cladocerans (Bertolani et al. 2009).

Figure 34. Fontinalis antipyretica in flowing water. Photo by Michael Lüth, with permission.

In the Italian Alps Borealibius zetlandicus occurred on Warnstorfia exannulata (Figure 35; Rebecchi et al. 2009). This tardigrade species is known only from boreo-alpine areas, where it typically occurs in sediment, submerged aquatic mosses, or Sphagnum (Figure 5). But like many other tardigrades, this one has a wide habitat range, including the Barents Sea and terrestrial mosses and soil that rarely dry out. This boreal habitat distribution for this species is possible because this species is able to survive freezing. But the populations of the species studied are unable to survive desiccation.

Figure 35. Warnstorfia exannulatus, home for the tardigrade Borealibius zetlandicus in the Italian Alps. Photo by Michael Lüth, with permission.

Figure 36. Murrayon dianeae, an aquatic tardigrade. Photo by Michael Collins, with permission.

Emergent bryophytes may be especially comfortable for some species of tardigrades. One of the more "friendly" environments is in association with Barbula [=Didymodon] tophacea (Figure 37-Figure 38), a well-known rockforming moss, above the wet zone.

Figure 37. Barbula [=Didymodon] tophacea, an emergent moss known to house 84 tardigrades per gram. Photo by Barry Stewart, with permission.

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Chapter 5-3: Tardigrade Habitats

The aquatic habitat is also sometimes hospitable to hygrophilous species that are more typical among moist mosses or species with a wide tolerance range for moisture conditions (Nelson & Marley 2000). On the other hand, Hypsibius dujardini (Figure 13) is known from moist mosses, but it is primarily aquatic (McFatter et al. 2007). Parhexapodibius pilatoi was found among mosses on a stream bank in central Georgia (McFatter et al. 2007), but otherwise is known only from Michigan (Bernard 1977).

Figure 38. Barbula [=Didymodon] tophacea, an emergent moss showing the numerous possibilities for resting in leaf axils. Photos by Michael Lüth, with permission.

Dry Habitats Although collectors looking for a rich bryophyte flora would most likely ignore the desert, some bryophytes depend on its predominately dry nature. The cryptogamic crust of prairies and deserts has its own tardigrade fauna (Brantley & Shepherd 2002). In this habitat of bryophytes, lichens, Cyanobacteria, and algae, long dry periods are guaranteed. The occasional wet periods make it a suitable tardigrade habitat. As already discussed, Meininger and Spatt (1988), working along Dalton Highway in the tundra adjacent to the trans-Alaska Pipeline, found that road dust had a major impact on both the mosses and the tardigrades. The mosses near the road represented xerophytic species tolerant of high calcium. Consequently, the tardigrades likewise were taxa tolerant of drier conditions. Because of the moisture limitations on other kinds of taxa, the tardigrades near the road were mostly fungivorous and algal feeders; those farther from the road, where Sphagnum (Figure 5) was able to grow, tended to be more omnivorous and carnivorous. These habitat differences caused differences in tardigrade fauna between roadside bryophytes and more distant Sphagnum species. Vertical and Horizontal Distribution It seems likely that some vertical distribution within the bryophyte mat should occur. These could be defined by light levels. The presence of eyespots (Figure 41) in at least some members of Tardigrada was reported by Greven (2007), with responses to light varying from none to both positive and negative. Beasley (2001) reported negative photokinesis in the common tardigrade Macrobiotus hufelandi (Figure 19), a common moss dweller. Rather than being attracted to or from the light, they increased

their rate of movement. Since light indicates sun intensity, it also is an indicator of the likelihood of drying, making the response to move quickly away from light an adaptive one. Vertical differences in tardigrade distribution are known from soil (Leetham et al. 1982). Nevertheless, as noted elsewhere, there seems to be little evidence for vertical position differences or migration of tardigrades in mosses; only one tardigrade (Echiniscus viridissimus) seems more common near the upper portion of the moss (Nelson & Adkins 2001). Wright (1991) found that in xeric habitats this species does not migrate vertically to the C zone as the moss dehydrates, even though other species do at the same time. Data from the Antarctic suggest that temperature may play a role in the vertical positioning of tardigrades there. On Signy Island, 80% of the tardigrades occurred in the upper 6 cm of moss, and usually 70% were in the top 3 cm (Jennings 1979). One factor that contributes to this limited distribution is that the turf below 7-8 cm is anaerobic (lacking oxygen), making it inhospitable for the tardigrades. In moss-dominated flushes near the Canada Glacier in southern Victoria Land, Antarctica, the invertebrates, including tardigrades, occurred at a mean depth ranging 5-10.83 mm (Schwarz et al. 1993). As discussed above, the relative number of organisms increased near the surface in post-melt mosses. This is not necessarily a direct temperature response; it could result from changes in light or humidity associated with the melt. Schuster et al. (2009) examined the microclimate within a cushion of the moss Rhytidiadelphus loreus (Figure 39). They found that the deep layers had lower daytime and higher nighttime temperatures than ambient (in this case, air temperature). Oxygen was similar throughout the cushion, but CO2 increased greatly with depth. The six species of tardigrades were concentrated in the green-brown layer of the moss. The authors suggested that light and oxygen had little impact on the distribution but that CO2 kept the tardigrades from occupying lower positions and that temperature might cause migrations within the upper portion.

Figure 39. Rhytidiadelphus loreus. Photo by Michael Lüth, with permission.

Differences in horizontal distribution may be the result of microhabitat differences such as shade vs sun or distance from water. But they can also be a simple result of passive, random dispersal and the slow-moving nature of the animal. Degma et al. (2011) sampled Hypnum cupressiforme (25 samples; Figure 40) to try to determine

Chapter 5-3: Tardigrade Habitats

the causes of horizontal positioning of tardigrades on that species. They found 224 tardigrades in the species Milnesium tardigradum (Figure 41), Hypsibius convergens (Figure 42), H. microps, Diphascon pingue (Figure 17), Astatumen trinacriae (Figure 43), Macrobiotus hufelandi (Figure 7), and Minibiotus sp. (Figure 44) They found no significant moisture gradient among the moss plants. The distribution of the tardigrade species was aggregated, but the number of species (richness) was random. There was no relationship of tardigrade species distribution to moisture.

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But even these species may not be able to tolerate desiccation. Macrobiotus hufelandi (Figure 7) and Hypsibius dujardini (Figure 13) are unable to live in habitats that desiccate quickly, perhaps explaining their association with the slow-drying bryophytes (Wright 1991). Other bryophyte dwellers [Milnesium tardigradum (Figure 41) and Ramazzottius (formerly Hypsibius) oberhaeuseri (Figure 3)] may not tolerate constant moisture, but these two species avoid high insolation and rapid desiccation, again making bryophytes a suitable habitat.

Figure 43. Astatumen trinacriae. Photo by Paul J. Bartels, with permission.

Figure 40. Hypnum cupressiforme, home of seven species of tardigrades. Photo by Michael Lüth, with permission.

Figure 44. Minibiotus intermedius. Miller, through Flickr.

Figure 41. Eyespots of Milnesium tardigradum. Photo by Martin Mach, with permission.

Figure 42. Hypsibius convergens. Kaczmarek, with permission.

Photo by Łukasz

Photo by William

Competition and food relations are often determinants of the species assemblages. For example, Milnesium tardigradum (Figure 41) may be found with two Hypsibius species that it can use for food (Wright 1991). Competition may account for the negative associations among Macrobiotus hufelandi (Figure 7), Paramacrobiotus richtersi (Figure 45), and Isohypsibius prosostomus (see Figure 46) in xeric habitats.

Figure 45. Paramacrobiotus richtersi. Creative Commons.

Photo through

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Chapter 5-3: Tardigrade Habitats

images, and my appreciation goes to all those who have contributed their images to Wikimedia Commons for all to use. Martin Mach was kind enough to send me corrections for typos in the previous online version. Thank you to my sister, Eileen Dumire, for providing the view of a novice on the readability of the text. Tardigrade nomenclature is based on Degma et al. 2010.

Literature Cited

Figure 46. Isohypsibius asper. Photo by Smithsonian Institution through EOL Creative Commons.

Summary Tardigrades live on both terrestrial and aquatic bryophytes, with the lowest numbers in the tropics. Epiphytic mosses seem to be especially suitable for them. Altitude is influential on species and numbers in some cases, but in others researchers were unable to find any differences. There are indications that the greatest numbers on higher mountains occur at mid elevations. Latitude, scale, and moisture availability most likely play a role. Numbers of species in most studies are modest. In forests, studies reported herein range 7-42 species. In one study, the number of species was greater in the pine forest, but the number of individuals differed little from that of clearcut areas. Macrobiotus hufelandi seems to be the most common species just about everywhere. The most common genera are Echiniscus, Macrobiotus, Diphascon, and Hypsibius, and the genera recently segregated from them. Tardigrades comprised only 1.2% of the invertebrates in an expansive New Zealand study and Hingley found only two taxa in peatlands. Dry habitats may pose food limitations; constantly wet ones may be unfavorable to their longevity. Although mosses get wet and dry on top first, it appears that tardigrades have little ability to migrate and do not even seem to be arranged in vertical assemblages. But, they have eyespots, indicating that light may play some role in their locations.

Acknowledgments Roberto Bertolani provided an invaluable update to the tardigrade taxonomic names and offered several suggestions on the text to provide clarification or correct errors. Łukasz Kaczmarek has provided me with references, images, contact information, and a critical read of an earlier version of the text. Martin Mach and Yuuji Tsukii have given permission to use images that illustrate the species and life cycle stages. And a big thank you goes to Michael Lüth for permission to use his many bryophyte

Bartels, P. 2005. "Little known" water bears? ATBI Quart. 6(2): 4. Bartels, P. J. and Nelson, D. R. 2006. A large-scale, multihabitat inventory of the phylum Tardigrada in the Great Smoky Mountains National Park, USA: A preliminary report. Hydrobiologia 558: 111-118. Beasley, C. W. 1981. Some Tardigrada from Puerto Rico. Tex. J. Sci. 33: 9-12. Beasley, C. W. 1988. Altitudinal distribution of Tardigrada of New Mexico with the description of a new species. Amer. Midl. Nat. 120: 436-440. Beasley, C. W. 2001. Photokinesis of Macrobiotus hufelandi (Tardigrada, Eutardigrada). Zool. Anz. 240: 233-236. Beasley, C. W., Kaczmarek, £., and Michalczyk, £. 2006. New records of tardigrades from China, with zoogeographical remarks. Biol. Lett. 43(1): 13-20. Bernard, E. C. 1977. A new species of Hexapodibius from North America, with a redescription of Diphascon belgicae (Tardigrada). Trans. Amer. Microsc. Soc. 96: 476-482. Bertolani, R. and Rebecchi, L. 1996. The tardigrades of Emilia (Italy). II. Monte Rondinaio. A multihabitat study on a high altitude valley of the northern Apennines. Zool. J. Linn. Soc. 116: 3-12. Bertolani, R., Altiero, T., and Nelson, D. R. 2009. Tardigrada (water bears). Chapter 188. In: Likens, G. E. (ed.). Volume 2. Encyclopedia of Inland Waters. Elsevier, Oxford, pp. 443-465. Bertrand, M. 1975. Répartition des tardigrades "terrestres" dans le massif de l'Aigoual. Vie Milieu 25: 283-298. Bettis, C. J. 2008. Distribution and abundance of the fauna living in two Grimmia moss morphotypes at the McKenzie Table Mountain Preserve, Fresno County, California. M.S. Thesis, California State University, Fresno, CA, 65 pp. Brantley, S. and Shepherd, U. 2002. Microarthropods on different types of cryptobiotic crusts in pinyon-juniper habitat. Abstracts of the 87th Annual Meeting of the Ecological Society of America and the 14th Annual International Conference of the Society for Ecological Restoration, 4-9 August, 2002, Tucson, AZ. Briones, M. J. I., Ineson, P., and Piearce, T. G. 1997. Effects of climate change on soil fauna; responses of enchytraeids, Diptera larvae and tardigrades in a transplant experiment. Appl. Soil Ecol. 6: 117-134. Collins, M. and Bateman, L. 2001. The ecological distribution of tardigrades in Newfoundland. Zool. Anz. 240: 291-297. Crum, H. 1976. Mosses of the Great Lakes Forest, revised edition. Contributions from the University Herbarium, Univ. Mich., Ann Arbor. Vol. 10, 404 pp. Dastych, H. 1980. Tardigrades from the Tatra National Park. Polska Akademia Nauk Zaklad Zoologii Systematycznej i Doswiadczalnej. Monografie Fauny Polski Vol. 9. 232. pp. Dastych, H. 1985. West Spitsbergen Tardigrada. Acta Zool. Cracov. 28(3): 169-214.

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Dastych, H. 1987. Altitudinal distribution of Tardigrada in Poland. In: Bertolani, R. (ed.). Selected Symposia and Monographs U.Z.I., 1, Mucchi, Modena, pp. 45-56. Dastych, H. 1988. The Tardigrada of Poland. Monografie Fauny Polski. Vol. 16. Institute of Systematics and Evolution of Animals, Polish Academy of Sciences, Kraków, Poland, pp. 1-255. Degma, P., Bertolani, R., and Guidetti, R. 2010. Actual checklist of Tardigrada species (Ver. 11:26-01-2010). Accessed 18 June 2010 at . Degma, P., Stanislav, K., and Sabatovicova, L. 2011. Horizontal distribution of moisture and Tardigrada in a single moss cushion. J. Zool. Syst. Evol. Res. 49 (Suppl 1): 71-77. Eles, A. and Repas, L. 2009. New dates and new methods for research of freshwater Tardigrada fauna of city Szekesfehervar. In: Greven, H., Hohberg, K., and Schill, R. O. 11th International Symposium on Tardigrada. Conference Guide. Tübingen, Germany, 3-6 August 2009, pp. 56-57. Elgmork, K. and Sæther, O. R. 1970. Distribution of invertebrates in a high mountain brook in the Colorado Rocky Mountains. Univ. Colorado Studies Ser. Biol. 31: 355. Fontoura, P., Pilato, G., Morais, P., and Lisi, O. 2009. Minibiotus xavieri, a new species of tardigrade from the Parque Biológico de Gaia, Portugal (Eutardigrada: Macrobiotidae). Zootaxa 2267: 55-64. Garey, J. R., McInnes, S. J., and Nichols, P. B. 2008. Global diversity of tardigrades (Tardigrada) in freshwater. Hydrobiologia 595: 101-106. Gray, N. F., Wyborn, C. H. E., and Smith, R. I. L. 1982. Nematophagous fungi from the maritime Antarctic. Oikos 38: 194-201. Greven, H. 2007. Comments on the eyes of tardigrades. Arthropod Struc. Dev. 36: 401-407. Guil, N., Hortal, J., Sánchez-Moreno, S., and Machordom, A. 2009. Effects of macro and micro-environmental factors on the species richness of terrestrial tardigrade assemblages in an Iberian mountain environment. Landscape Ecol. 24: 375390. Hingley, M. 1993. Microscopic Life in Sphagnum. Naturalists' Handbook 20. [i-iv]. Richmond Publishing Co. Ltd., Slough, England, pp. 1-64. 58 fig. 8 pl. Hinton, J. G., and Meyer, H. A. 2007. Distribution of limnoterrestrial Tardigrada in Georgia and the Gulf Coast states of the United States of America with ecological remarks. J. Limnol. 66(Suppl. 1): 72-76. Hinton, J. G. and Meyer, H. A. 2008. Tardigrades from Fayette County, Georgia. Georgia J. Sci. 66: 30-32. Hofmann, I. 1987. Habitat preference of the most frequent mossliving Tardigrada in the area of Giessen (Hessen). In: Bertolani, R. (ed.). Selected Symposia and Monographs U.Z.I., 1, Mucchi, Modena, pp. 211-216. Hooie, A. K. and Davison, P. G. 2001. Tardigrade diversity in the moss Ulota crispa from tree canopies in the Great Smoky Mountains National Park – A preliminary report. J. Alabama Acad. Sci. 72: 91. Abstract. Ito, M. 1999. Ecological distribution, abundance and habitat preference of terrestrial tardigrades in various forests on the northern slope of Mt. Fuji, central Japan. Zool. Anz. 238: 225-234.

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Jennings, P. G. 1976. The Tardigrada of Signy Island, South Orkney Islands, with a note on the Rotifera. Brit. Antarct. Surv. Bull 44: 1-25. Jennings, P. G. 1979. The Signy Island terrestrial reference sites: X. Population dynamics of Tardigrada and Rotifera. Brit. Antarct Surv. Bull. 47: 89-105. Jerez Jaimes, J. H., Narváez, E. X., and Restrepo, R. 2002. Tardígrados en musgos de la Reserva el Diviso (Santander, Colombia). Revista Colombiana de Entomología 28(2): 199206. Jönsson, K. I. 2003. Population density and species composition of moss-living tardigrades in a boreo-nemoral forest. Ecography 26: 356-364. Kaczmarek, L., Gołdyn, B., Wełnicz, W., and Michalczyk, L. 2011. Ecological factors determining Tardigrada distribution in Costa Rica. J. Zool. Syst. Evol. Res. 49: 78-83. Kathman, R. D. and Cross, S. F. 1991. Ecological distribution of moss-dwelling tardigrades on Vancouver Island, British Columbia, Canada. Can. J. Zool. 69: 122-129. Leetham, J. W., McNary, T. J., Dodd, J. L., and Lauenroth, W. K. 1982. Response of soil nematodes, rotifers and tardigrades to three levels of season-long sulfur dioxide exposures. Water Air Soil Pollut. 17: 343-356. Li, X.-C. and Wang, L.-Z. 2005. Effects of temperature and thermal acclimation on locomotor performance of Macrobiotus hufelandi Schultze (Tardigrada: Macrobiotidae). Current Zool. (formerly Acta Zoologica Sinica) 51: 516-520. Linhart, J., Vlcková, S., and Uvíra, V. 2002. Moss-dwelling meiobenthos and flow velocity in low-order streams. Acta Universitatis Palackianae Olomucensis Facultas Rerum Naturalium (2001-2002) Biologica 39/40: 111-122. Mathews, G. B. 1938. Tardigrada of North America. Amer. Midl. Nat. 19: 619-627. McFatter, M. M., Meyer, H. A., and Hinton, J. G. 2007. Nearctic freshwater tardigrades: A review. In: Pilato, G. and Rebecchi, L. (guest eds.). Proceedings of the Tenth International Symposium on Tardigrada. J. Limnol. 66(Suppl. 1): 84-89. Meininger, C. A. and Spatt, P. D. 1988. Variations of tardigrade assemblages in dust-impacted Arctic mosses. Arct. Alp. Res. 20: 24-30. Meininger, C. A., Uetz, G. W., and Snider, J. A. 1985. Variation in epiphytic microcommunities (tardigrade-lichen-bryophyte assemblages) of the Cincinnati, Ohio area. Urban Ecol. 9(1): 45-62. Meyer, H. A. 2006. Small-scale spatial distribution variability in terrestrial tardigrade populations. Hydrobiologia 558: 133139. Meyer, H. A. 2008. Distribution of tardigrades in Florida. Southeast. Nat. 7: 91-100. Meyer, H. A. and Hinton, J. 2007. Limno-terrestrial Tardigrada of the Nearctic Realm. In: Pilato, G. and Rebecchi, L. (eds.). Proceedings of the Tenth International Symposium on Tardigrada. J. Limnol. 66(Suppl. 1): 97-103. Mihelcic, F. 1954/55. Zur Ökologie der Tardigraden. Zool. Anz. 153: 250-257. Mihelcic, F. 1963. Moose als Lebensstätten für Tardigraden. Der Schlern 37: 179-181. Miller, W. R. and Heatwole, H. 1995. Tardigrades of the Australian Antarctic Territories: The Mawson Coast, East Antarctica. Invert. Biol. 114(1): 27-38. Miller, W. R., Miller, J. D., and Heatwole, H. 1996. Tardigrades of the Australian Antarctic Territories: The Windmill Islands, East Antarctica. Zool. J. Linn. Soc. 116: 175-184.

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Nelson, D. R. 1973. Ecological distribution of tardigrades on Roan Mountain, Tennessee – North Carolina. Ph. D. dissertation, University of Tennessee, Knoxville. Nelson, D. R. 1975. Ecological distribution of Tardigrada on Roan Mountain, Tennessee – North Carolina. In: Higgins, R. P. (ed.). Proceedings of the First International Symposium on Tardigrades. Mem. Ist. Ital. Idrobiol., Suppl. 32: 225-276. Nelson, D. R. and Adkins, R. G. 2001. Distribution of tardigrades within a moss cushion: Do tardigrades migrate in response to changing moisture conditions? Zool. Anz. 240: 493-500. Nelson, D. R. and Marley, N. J. 2000. The biology and ecology of lotic Tardigrada. Freshwat. Biol. 44: 93-108. Newsham, K. K., Maslen, N. R., and McInnes, S. J. 2006. Survival of Antarctic soil metazoans at -80ºC for six years. CryoLetters 27(5): 269-280. Pilato, G. 2009. Bindius triquetrus gen. nov. sp. nov. (Eutardigrada, Hypsibiidae) from Sicily (Italy). Zootaxa 2058: 62-68. Ramazzotti, G. 1962. Il Phylum Tardigrada. Mem. Ist. Ital. Idrobiol. 14: 1-595. Ramazzotti, G. 1972. Phylum Tardigrada (Seconda edizione aggionata). Vol. II. Mem. Ist. ital. Idrobiol. 28: 1-732. Ramazzotti, G. and Maucci, W. 1983. Il Phylum Tardigrada. [The phylum Tardigrada - 3rd edition: English translation by C. W. Beasley.]. Mem. Ist. Ital. Idrobiol. 41: 1-1011. Rebecchi, L., Boschini, D., Cesari, M., Lencioni, V., Bertolani, R., and Guidetti, R. 2009. Stress response of a boreo-alpine species of tardigrade, Borealibius zetlandicus (Eutardigrada, Hypsibiidae). J. Limnol., 68(1): 64-70. Riggin, G. T. Jr. 1962. Tardigrada of Southwest Virginia: With the Addition of a Description of a New Marine Species from Florida. Technical Bull. 152, Virginia Agricultural Experiment Station, Blacksburg, VA. 145 pp. Rodrigues-Roda, J. 1951. Alguños datos sobre la distribucion de los Tardigrados españoles. Bol. Real Soc. Esp. Hist. Nat. Secc. Biol. 49: 75-83. Romano, F. A. III., Barreras-Borrero, B., and Nelson, D. R. 2001. Ecological distribution and community analysis of Tardigrada from Choccolocco Creek, Alabama. Zool. Anz. 240: 535-541. Schuster, R. and Greven, H. 2007. A long-term study of population dynamics of tardigrades in the moss Rhytidiadelphus squarrosus (Hedw.) Warnst. J. Limnol. 66: 141-151. Schuster, R., Spiertz, I., Lösch, R., and Greven, H. 2009. Microclima and vertical distribution of tardigrades in a moss cushion. In: Greven, H., Hohberg, K., and Schill, R. O.

11th International Symposium on Tardigrada. Conference Guide. Tübingen, Germany, 3-6 August 2009, p. 59. Schwarz, A.-M. J., Green, J. D., Green, T. G. A., and Seppelt, R. D. 1993. Invertebrates associated with moss communities at Canada Glacier, southern Victoria Land, Antarctica. Polar Biol. 13: 157-162. Shcherbako, D., Schill, R. O., Brümmer, F., Nissen, M., and Blum, M. 2010. Movement behaviour and video tracking of Milnesium tardigradum Doyère, 1840 (Eutardigrada Apochela). Contrib. Zool. 79: 33-38. Sohlenius, B. and Boström, S. 2006. Patch-dynamics and population structure of nematodes and tardigrades on Antarctic nunataks. Eur. J. Soil Biol. 42(Suppl. 1): S321S325. Somme, L. 1996. Anhydrobiosis and cold tolerance in tardigrades. Eur. J. Entomol. 93: 349-357. Somme, L. and Terje Meier, T. 1995. Cold tolerance in Tardigrada from Dronning Maud Land, Antarctica. Polar Biol. 15: 1432-2056. Suren, A. M. 1991a. Bryophytes as invertebrate habitat in two New Zealand alpine streams. Freshwat. Biol. 26: 399-418. Suren, A. M. 1991b. Assessment of artificial bryophytes for invertebrate sampling in two New Zealand alpine streams. N. Z. J. Marine Freshwat. Res. 25: 101-112. Suren, A. M. 1992. Enhancement of invertebrate food resources by bryophytes in New Zealand alpine headwater streams. N. Z. J. Marine Freshwat. Res. 26: 229-239. Suren, A. 1993. Bryophytes and associated invertebrates in firstorder alpine streams of Arthur's Pass, New Zealand. New Zeal. J. Marine Freshwat. Res. 27: 479-494. Utsugi, K., Hiraoka, T., and Nunomura, N. 1997. On the relations between tardigrade fauna and bryophyte flora in Toyama Prefecture. Bull. Toyama Sci. Mus. 20: 57-71. Venkataraman, K. 1998. Studies on Phylum Tardigrada and Other Associated Fauna, South Polar Skua and Bird and Mammal Logging During 1994 - 1995 Expedition. Fourteenth Indian Expedition to Antarctica, Scientific Report, 1998. Department of Ocean Development, Technical Publication No. 12: 221-243. Vlcková, S., Linhart, J., and Uvíra, V. 2001/2002. Permanent and temporary meiofauna of an aquatic moss Fontinalis antipyretica Hedw. Acta Univers. Palack. Olom. Biol. 3940: 131-140. Westh, P. and Kristensen, R.-M. 1992. Ice formation in the freeze-tolerant eutardigrades Adorybiotus coronifer and Amphibolus nebulosus studied by differential scanning calorimetry. Polar Biol. 12: 693-699. Wright, J. C. 1991. The significance of four xeric parameters in the ecology of terrestrial Tardigrada. J. Zool. 224: 59-77.

Glime, J. M. 2017. Tardigrades: Species Relationships. Chapt. 5-4. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. Ebook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 18 July 2020 and available at .

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CHAPTER 5-4 TARDIGRADES: SPECIES RELATIONSHIPS TABLE OF CONTENTS Species Relationships .......................................................................................................................................... 5-4-2 Life Forms.................................................................................................................................................... 5-4-7 Liverworts .................................................................................................................................................... 5-4-9 Substrate Comparisons ...................................................................................................................................... 5-4-11 Finding New Species......................................................................................................................................... 5-4-12 Summary ........................................................................................................................................................... 5-4-14 Acknowledgments ............................................................................................................................................. 5-4-14 Literature Cited ................................................................................................................................................. 5-4-15

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CHAPTER 5-4 TARDIGRADES: SPECIES RELATIONSHIPS

Figure 1. SEM of tardigrades on a leafy liverwort.

Photo by Łukasz Kaczmarek and Łukasz Michalczyk, with permission.

Species Relationships Tardigrades occur on both mosses and liverworts (Figure 1). Since bryophytes vary widely in structure, compactness, and moisture-holding nature, one would expect that some bryophytes would be more suitable for tardigrades than others, causing specificity. But is that really the case? Although Hofmann and Eichelberg (1987), in Lahnau near Giessen, Germany, found a correlation between species of tardigrade and degree of moisture in their preferred mosses, there seemed to be no example of a single species of tardigrade preferring a single species of moss. It appeared that species of bryophyte was not an important factor for most tardigrades. A number of studies name the bryophytes where the tardigrades have been found, but quantitative approaches

are limited. For example, Degma (2006) found Echiniscus reticulatus on the moss Ctenidium molluscum (Figure 2) and Testechiniscus spitsbergensis from the mosses Tortella tortuosa (Figure 3), Ctenidium molluscum (Figure 2), Distichium capillaceum (Figure 4), and Ditrichum flexicaule (Figure 5-Figure 6) in Slovakia. Baxter (1979) did find differences in the tardigrades on several moss species in Ireland. These represented different life forms as well as habitats. Some of their more specific finds include stream bank mosses that had Diphascon oculatum (Figure 7). Polytrichum (Figure 8), with its more open structure, had Diphascon scoticum (Figure 9). Hypsibius dujardini (Figure 1) was abundant, accompanied by Isohypsibius tuberculatus, on the turfs of Rhytidiadelphus squarrosus (Figure 10).

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Figure 6. View inside cushion of Ditrichum flexicaule, a habitat for Testechiniscus spitsbergensis. Photos by Michael Lüth, with permission. Figure 2. Ctenidium molluscum, a moss that is home to Echiniscus reticulatus, among others. Photo by Michael Lüth, with permission.

Figure 7. Diphascon oculatum, an inhabitant of streambank mosses. Photo by Björn Sohlenius, Swedish Museum of Natural History, with permission.

Figure 3. Tortella tortuosa, a Slovakian habitat for Testechiniscus spitsbergensis. Photo by Michael Lüth, with permission.

Figure 8. Polytrichum, a moss with spreading leaves that provide limited tardigrade habitat. Photo by Michael Lüth, with permission.

Figure 4. Distichium capillaceum, a known tardigrade habitat. Photo by Michael Lüth, with permission.

Figure 5. Ditrichum flexicaule, a habitat for Testechiniscus spitsbergensis. Photos by Michael Lüth, with permission.

Figure 9. Diphascon scoticum, a tardigrade that is able to inhabit Polytrichum. Photo by Paul J. Bartels, with permission.

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Figure 10. Rhytidiadelphus squarrosus, where Baxter (1979) found Isohypsibius tuberculatus and Diphascon scoticum. Photo by Michael Lüth, with permission.

Figure 11. Lembophyllum divulsum, a home for 16 tardigrade species in New Zealand. Photo by Li Zhang, with permission.

Figure 12. Hypnum cupressiforme, the moss with the most tardigrade species in the New Zealand study by Horning et al. (1978), shown here on rock and as a pendant epiphyte. Photos by Michael Lüth, with permission.

Horning et al. (1978) examined the tardigrades on 21 species of mosses in New Zealand and listed the tardigrade species on each (Table 1). Some moss species clearly had more tardigrade species than others, ranging from 1 on Syntrichia rubra to 17 on Hypnum cupressiforme (Figure 12). Lembophyllum divulsum (Figure 11) had 16 species. Hopefully lists like the one provided by Horning et al. (1978) will eventually permit us to determine the characteristics that foster tardigrade diversity and abundance. Perhaps the moss Hypnum cupressiforme (Figure 12) had the most tardigrade species among the mosses in New Zealand because of its own wide habitat range there. However, Degma et al. (2005) found that distribution of the number of tardigrade species on this moss in their Slovakia sites was random, as supported by a Chi-square goodness of fit test. But this still does not preclude the assertion that its ubiquitous nature on a wide range of habitats in New Zealand may account for the greater number of species of tardigrades on Hypnum cupressiforme in the New Zealand study. A kind of vertical zonation occurs among tardigrades on trees that is the reverse of that sometimes found within a moss cushion. In the Great Smoky Mountains National Park, the number of tardigrade species among epiphytes at breast height was greater than the number of species found at the base (Bartels & Nelson 2006). This may relate to the

need for dry periods, but it could also relate to differences in predators and possibly even to dispersal patterns. In their study of Chinese mosses Beasley and Miller (2007) found that Heterotardigrada (armored tardigrades) were better represented than were Eutardigrada (unarmored tardigrades), a factor the authors attribute to the xerophilic moss samples and the locality, which has hot, dry summers, very cold, dry winters, low summer rainfall, and high winds (Fullard 1968). The Heterotardigrada have armor, which may account for their ability to withstand the dry habitat. These tardigrades also have cephalic (head) appendages with a sensorial function, a character lacking in the Eutardigrada, but so far their function has not been related to a bryophyte habitat. Beasley and Miller found little specificity, but most of the mosses were xerophytic and exhibited similar moisture requirements. They did find that Echiniscus testudo (Figure 13) occurred on a wider variety of mosses than did other tardigrade species. On Roan Mountain in Tennessee and North Carolina, Nelson (1973, 1975) found no specificity among 21 tardigrade species on 25 bryophyte species. Hunter (1977) in Montgomery County, Tennessee, and Romano et al. (2001) in Choccolocco Creek in Alabama, USA, again were unable to find any dependence of tardigrades upon a particular species of bryophyte in their collections.

Chapter 5-4: Tardigrades: Species Relationships

5-4-5

Table 1. Tardigrade species found on the most common moss taxa in New Zealand. From Horning et al. 1978. Breutelia elongata

Breutelia pendula

Bryum campylothecium Bryum dichotomum Bryum truncorum

Dicranoloma billardieri Dicranoloma grossialare

Dicranoloma menziesii Dicranoloma robustum

Dicranoloma trichopodum

Hypnum cupressiforme

Lembophyllum divulsum

Macrobiotus hibiscus Macrobiotus liviae Milnesium tardigradum Minibiotus intermedius Diphascon prorsirostre Diphascon scoticum Doryphoribius zyxiglobus Hypechiniscus exarmatus Macrobiotus hibiscus Macrobiotus liviae Milnesium tardigradum Hypsibius convergens Isohypsibius sattleri Minibiotus intermedius Hypsibius wilsoni Macrobiotus coronatus Macrobiotus liviae Diphascon chilenense Diphascon scoticum Isohypsibius sattleri Isohypsibius wilsoni Macrobiotus coronatus Macrobiotus furciger Macrobiotus liviae Macrobiotus recens Paramacrobiotus areolatus Paramacrobiotus richtersi Ramazzottius oberhaeuseri Hypechiniscus exarmatus Macrobiotus hibiscus Diphascon prorsirostre Hypechiniscus exarmatus Hypsibius dujardini Isohypsibius cameruni Isohypsibius sattleri Limmenius porcellus Macrobiotus anderssoni Macrobiotus hibiscus Macrobiotus liviae Milnesium tardigradum Pseudechiniscus novaezeelandiae Macrobiotus hibiscus Macrobiotus liviae Paramacrobiotus areolatus Echiniscus bigranulatus Macrobiotus anderssoni Macrobiotus furciger Macrobiotus liviae Milnesium tardigradum Pseudechiniscus juanitae Echiniscus quadrispinosus Echiniscus q. brachyspinosus Macrobiotus furciger Pseudechiniscus lateromamillatus Diphascon alpinum Diphascon bullatum Echiniscus quadrispinosus Echiniscus spiniger Hypsibius dujardini Macrobiotus anderssoni Macrobiotus coronatus Macrobiotus furciger Macrobiotus hibiscus Macrobiotus liviae Macrobiotus recens Milnesium tardigradum Oreella mollis Paramacrobiotus areolatus Pseudechiniscus novaezeelandiae Pseudechiniscus juanitae Ramazzottius oberhaeuseri Diphascon alpinum Doryphoribius zyxiglobus Hypsibius convergens Isohypsibius sattleri Macrobiotus anderssoni Macrobiotus coronatus

Macrobiotus furciger Macrobiotus hibiscus Macrobiotus liviae Macrobiotus recens Macrobiotus subjulietae Milnesium tardigradum Minibiotus intermedius Paramacrobiotus areolatus Pseudechiniscus novaezeelandiae Pseudechiniscus juanitae Macromitrium erosulum Macrobiotus furciger Macrobiotus hibiscus Macrobiotus liviae Pseudechiniscus juanitae Macromitrium longipes Doryphoribius zyxiglobus Hypsibius convergens Macrobiotus recens Minibiotus intermedius Porotrichum ramulosum Diphascon alpinum Diphascon scoticum Doryphoribius zyxiglobus Echiniscus bigranulatus Hypsibius convergens Macrobiotus anderssoni Macrobiotus coronatus Macrobiotus furciger Macrobiotus hibiscus Macrobiotus liviae Macrobiotus rawsoni Minibiotus aculeatus Pseudechiniscus lateromamillatus Pseudechiniscus novaezeelandiae Pseudechiniscus juanitae Racomitrium crispulum Calohypsibius ornatus Diphascon alpinum Echiniscus quadrispinosus Echiniscus zetotrymus Hebesuncus conjungens Hypsibius convergens Isohypsibius wilsoni Macrobiotus anderssoni Macrobiotus coronatus Macrobiotus furciger Macrobiotus hibiscus Macrobiotus orcadensis Milnesium tardigradum Oreella minor Paramacrobiotus areolatus Pseudechiniscus juanitae Racomitrium lanuginosum Diphascon scoticum Echiniscus quadrispinosus brachyspinosus Echiniscus vinculus Hebesuncus conjungens Macrobiotus furciger Milnesium tardigradum Minibiotus intermedius Oreella mollis Pseudechiniscus juanitae Racomitrium ptychophyllum Echiniscus quadrispinosus Echiniscus velaminis Hebesuncus conjungens Hypechiniscus exarmatus Hypsibius dujardini Macrobiotus furciger Milnesium tardigradum Minibiotus intermedius Oreella mollis Syntrichia princeps Hypsibius convergens Ramazzottius oberhaeuseri Isohypsibius wilsoni Macrobiotus coronatus Macrobiotus recens Milnesium tardigradum Pseudechiniscus novaezeelandiae Syntrichia rubra Diphascon scoticum Tortula subulata var. serrulata Diphascon scoticum Paramacrobiotus areolatus

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Chapter 5-4: Tardigrades: Species Relationships

Table 2. Distribution of tardigrades on specific mosses in Xinjiang Uygur Region, China, based on herbarium specimens. From Beasley & Miller 2007.

tardigrade

Figure 13. Echiniscus testudo tun. Syred through Creative Commons.

numb/samples

Bryodelphax asiaticus Cornechiniscus holmeni

1/1 18/5

Echiniscus blumi

4/4

Echiniscus canadensis

82/7

Echiniscus granulatus

8/3

Echiniscus testudo

11/4

Echiniscus trisetosus

33/5

Macrobiotus mauccii Milnesium asiaticum

2/2 10/4

Milnesium longiungue

4/2

Milnesium tardigradum

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Photo by Power &

Hofmann developed a preference coefficient: n Pn = (Tn/Sn) Σ 100(Ti/Si) i=1 where P = preference index for category n of observed factor n = index of observed category T = number of tardigrade populations of a single species S = number of samples in category The preference indices will add up to 100%. The categories can be the five bryophyte habitat groups listed by Mihelčič 1954/55, 1963; Ramazzotti 1962, and Hofmann 1987 or other groupings defined for the purpose.

Paramacrobiotus alekseevi 5/4

moss

Pseudoleskeella catenulata Grimmia tergestina Mnium laevinerve Schistidium sp. Abietinella abietina Schistidium sp. Grimmia laevigata Grimmia ovalis Grimmia tergestina Grimmia longirostris Schistidium trichodon Schistidium sp. Grimmia anodon Grimmia longirostris Grimmia tergestina Lescuraea incurvata Pseudoleskeella catenulata Schistidium sp. Abietinella abietina Grimmia ovalis Pseudoleskeella catenulata Schistidium sp. Grimmia anodon Grimmia tergestina Grimmia ovalis Schistidium sp. Grimmia laevigata Grimmia ovalis Grimmia tergestina Grimmia ovalis Orthotrichum sp. Brachythecium albicans Schistidium sp.

Table 3. Preference of moss species by tardigrades, using five moss species plus the remaining species combined (total = 43 species) as the habitat categories, based on 106 samples from Giessen, Germany (Hofmann 1987).

Ceratodon purpureus samples (%) Macrobiotus hufelandi Ramazzottius oberhaeuseri Milnesium tardigradum Echiniscus testudo mean empty samples

Grimmia pulvinata

Bryum argenteum

Syntrichia ruralis

Syntrichia montana

Other

19 16 18 13 11

9 18 27 23 20

7 18 29 15 20

7 18 17 20 9

6 21 0 23 34

52 8 8 6 6

14.5 25

22.0 7

20.0 9

16.0 9

19.5 11

7.0 38

Figure 14. Macrobiotus hufelandi. Photo by Martin Mach, with permission.

Chapter 5-4: Tardigrades: Species Relationships

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associations and one positive association between species of tardigrades. Likewise, in Georgia and the Gulf Coast, USA, Hinton and Meyer (2007) found Milnesium tardigradum (Figure 16), Minibiotus intermedius (Figure 39), and Minibiotus furcatus among mosses, whereas Echiniscus cavagnaroi, E. kofordi (see Figure 15), and Minibiotus fallax were in both mosses and lichens.

Figure 15. Adult Echiniscus sp.. Photo by Martin Mach, with permission.

Kathman and Cross (1991) found that species of bryophyte had no influence on the distribution or abundance of tardigrades from five mountains on Vancouver Island, British Columbia, Canada. In fact, Kathman and Cross (1991) were unable to find any correlation with altitude or aspect throughout a span from 150 to 1525 m. They concluded that it was the presence of bryophyte that determined tardigrade presence, not the species of bryophyte, altitude, or locality. Despite a lack of specificity among the tardigrades, 39 species inhabited these 37 species of mountain bryophytes, comprising 14,000 individuals. Several researchers contend that any terrestrial species of tardigrade can be found on any species of moss, given the "appropriate microhabitat conditions" (Bertrand 1975; Ramazzotti & Maucci 1983). If these tardigrade bryophyte specialists find no differences among the bryophytes, can we blame the ecologists for lumping all the bryophytes in their studies as well? In collections from Giessen, Germany, the most common tardigrade species, the cosmopolitan Macrobiotus hufelandi (Figure 14), had no preference for any moss species (Hofmann 1987). But lack of influence of bryophyte species may not always be the case. Hofmann (1987) used a preference index to show that four out of sixteen tardigrades from Giessen had distinct preferences among five moss species and that they seemed to prefer cushion mosses over sheet mosses. Also contrasting with the above researchers, Bertolani (1983) found that there seemed to be a species relationship between tardigrades and coastal dune mosses. It is possible that this is again related to moisture. The moisture relationship might also explain why mosses on rotten logs seem to have few tardigrades. Could it be that they are too wet for too long? Meyer (2006a, b, 2008) found 20 species of tardigrades among 47 species of mosses, liverworts, lichens, and ferns in Florida. There were some tardigrade species that were significantly associated with either mosses or lichens, but, as in most other studies, there was no convincing evidence for associations with any plant species substratum. Despite the lack of substrate specificity, there were three significant negative

Figure 16. Milnesium tardigradum, an inhabitant of both mosses and liverworts. Photo by Björn Sohlenius, Swedish Museum of Natural History.

Life Forms There is some indication that species differences may exist, based on life form. The bryophyte form can affect the moisture-holding capacity and rate of loss of moisture. That foregoing evidence suggests that the moisture-holding capacity of cushion mosses was probably a desirable trait in that habitat. On the other hand, Beasley (1990) found that more samples of clubmosses (Lycopodiaceae – tracheophytes) (75%) had tardigrades than did mosses (46%) or liverworts (0%) in Gunnison County, Colorado. There seems to be a preference for cushions among the most common species [Macrobiotus hufelandi (Figure 14), Ramazzottius oberhaeuseri (Figure 17), Milnesium tardigradum (Figure 16), and Echiniscus testudo (Figure 13)] (Hofmann 1987). But the less frequent species are commonly found among sheet mosses. The ubiquitous Macrobiotus hufelandi seems to have no preference for moss shape.

Figure 17. Ramazzottius oberhaeuseri. Photo by Martin Mach, with permission.

Jönsson (2003), working in the forests of Sweden, found that wefts had more tardigrades than other moss forms. Kathman and Cross (1991) likewise found that tardigrades from Vancouver Island were more common on weft-forming mosses than on turfs, suggesting that the thick carpets of the wefts were more favorable habitat than the thinly clustered turfs with their thick rhizoidal mats and

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Chapter 5-4: Tardigrades: Species Relationships

attached soil. Contrasting with some of these findings, and the preference for cushion mosses in the study by Hofmann (1987), Diane Nelson (East Tennessee State University, Johnson City, pers. comm. in Kathman & Cross 1991) found no preference for sheet or cushion mosses in her Roan Mountain, Virginia, USA study. Rather, those tardigrades were more common in thin, scraggly mosses or in small tufts than in thick cushion mosses. Sayre and Brunson (1971) compared tardigrade fauna on mosses in 26 North American collections from a variety of habitats and substrata (Figure 18). They found that mosses of short stature in the Thuidiaceae (Figure 19) and Hypnaceae (Figure 20) had the highest frequencies of tardigrades. Other moss-dwellers were found in fewer numbers on members of the moss families Orthotrichaceae (epiphytic and rock-dwelling tufts; Figure 21), Leucobryaceae (cushions on soil and tree bases; Figure 22), Polytrichaceae (tall turfs on soil; Figure 23), Plagiotheciaceae (low mats on soil and tree bases; Figure 24), and Mniaceae (mats & wefts on soil; Figure 25).

Figure 20. Hypnum revolutum (Hypnaceae), representing a family that includes low-stature mosses that had among the highest frequencies of tardigrades in 26 North American collections (Sayre & Brunson 1971). Photo by Michael Lüth, with permission.

Figure 21. Orthotrichum pulchellum, an epiphytic moss in the Orthotrichaceae. This family is among those with lower numbers of tardigrades in the North American study of Sayre & Brunson (1971) compared to families of mat-forming species. Photo by Michael Lüth, with permission. Figure 18. Relative frequency of tardigrades on bryophytes of various North American substrata. Redrawn from Sayre & Brunson 1971.

Figure 19. Thuidium delicatulum (Thuidiaceae), a lowstature moss that is a good tardigrade habitat. Photo by Michael Lüth, with permission.

Figure 22. Leucobryum glaucum, a cushion moss in the Leucobryaceae. This family of mosses had lower numbers of tardigrades than those found in the mat-forming mosses in 26 North American collections (Sayre & Brunson 1971). Photo by Michael Lüth, with permission.

Chapter 5-4: Tardigrades: Species Relationships

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Collins and Bateman (2001), studying tardigrade fauna of bryophytes in Newfoundland, Canada, found that rate of desiccation of the mosses affected distribution of tardigrades, and this suggests that bryophyte species and life forms that dehydrate quickly should have fewer individuals and probably different or fewer species than those that retain water longer. In different climate regimes, that rate will differ. This may explain a preference for cushions in some locations and not in others. Data are needed on humidity within the various life forms of bryophytes, correlated with tardigrade densities, to try to explain why different life forms seem to be preferred in different locations. Liverworts Figure 23. Polytrichum juniperinum, a moss in the Polytrichaceae. This family of mosses tends to have low numbers of tardigrades (Sayre & Brunson 1971). The tardigrades do live among them often nestle in the leaf bases where water evaporates more slowly. Photo by Michael Lüth, with permission.

I would expect liverworts, with their flat structure, to have at least some differences in tardigrade communities (Figure 1). But reports on liverwort inhabitants are limited, at least in part due to lack of knowledge about bryophytes on the part of the tardigrade specialists and an equal lack of knowledge of tardigrades by bryologists. Hinton and Meyer (2009) found two species of tardigrades [Milnesium tardigradum (Figure 16) and Macrobiotus hibiscus], both also common among mosses, in samples of the liverwort Jungermannia sp. (Figure 26). In the Gulf Coast states, USA, Hinton and Meyer (2007) found Echiniscus virginicus among liverworts.

Figure 24. Plagiothecium denticulatum, a low-growing soil moss in Plagiotheciaceae, a family with limited numbers of tardigrade dwellers (Sayre & Brunson 1971). The flattened growth habit provides few protective chambers, perhaps accounting for the lower numbers. Photo by Michael Lüth, with permission. Figure 26. The leafy liverwort Jungermannia sphaerocarpa, representing a genus from which tardigrades are known. Photo by Michael Lüth, with permission.

Figure 25. Plagiomnium cuspidatum, a soil moss in the Mniaceae, a family with limited numbers of tardigrade dwellers (Sayre & Brunson 1971). The spreading nature of the vertical shoots and the flattened nature of the horizontal shoots would most likely not provide many protective chambers for the tardigrades. Photo by Michael Lüth, with permission.

Liverworts may actually house some interesting differences as a result of their underleaves (Figure 27) and flattened life form (Figure 28). In their New Zealand study, Horning et al. (1978) found that among the liverworts (Table 4), Porella elegantula (Figure 27) had the most species (16). The folds and underleaves of this genus form tiny capillary areas where water is held, perhaps accounting for the large number of species. Interestingly, the tardigrade Macrobiotus snaresensis occurred on several liverwort species [4 Lophocolea species, Plagiochila deltoidea (Figure 29)], but did not appear in any moss collections. Of 150 liverwort samples (26 species), 27% had tardigrades, with a total of 16 species, mean of 2.8 species, range 1-9. In 107 samples of foliose lichens, 60.7% had tardigrades, mean 2.2 species, range 1-11.

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Chapter 5-4: Tardigrades: Species Relationships

It appears that at least some other researchers have paid attention to liverworts. Christenberry (1979) found Echiniscus kofordi and E. cavagnaroi on liverworts in Alabama, USA. Hinton and Meyer (2009) found Milnesium tardigradum (Figure 16) and Macrobiotus hibiscus in a liverwort sample from Georgia, USA. Michalczyk and Kaczmarek (2006) found a new species, Paramacrobiotus magdalenae (Figure 30, Figure 31), on liverworts in Costa Rica. Newsham et al. (2006) identified the tiny leafy liverwort Cephaloziella varians and used it to experiment on the effects of low temperature storage on tardigrades and other Antarctic invertebrates.

Figure 27. Porella elegantula, showing the underleaves and folds that create numerous capillary spaces. Photo by Jan-Peter Frahm, with permission.

Figure 30. Paramacrobiotus magdalenae egg. Photo by Łukasz Kaczmarek and Łukasz Michalczyk, with permission.

Figure 28. Underside of leafy liverwort with two tardigrades. Photo by Łukasz Kaczmarek and Łukasz Michalczyk, with permission.

Figure 31. Paramacrobiotus areolatus. Photo by Martin Mach, with permission.

Figure 29. Plagiochila deltoidea, a leafy liverwort that forms large patches in wet ground in New Zealand. This is a known habitat for tardigrades. Photo by Clive Shirley, Hidden Forest , with permission.

Just what do we mean by "appropriate habitat conditions"? The bryophytes only occur in conditions that are appropriate for them, hence defining the conditions for the tardigrades. And the bryophytes create habitat conditions of moisture due to their morphology and substrate preference. Lack of species preference in many studies may result from methods that were insensitive to subtle differences or that failed to control for microhabitat differences. Usually no statistical tests were employed, sample sizes were small, and enumeration was often simple presence/absence data.

Chapter 5-4: Tardigrades: Species Relationships

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Table 4. Species of tardigrades found on 13 liverwort species in New Zealand and surrounding islands. From Horning et al. 1978. Liverwort Species Lophocolea innovata Lophocolea. minor Lophocolea. subporosa Lophocolea semiteres Lophocolea subporosa: Lophocolea sp. Metzgeria decipiens

Metzgeria decrescens

Plagiochila deltoidea

Tardigrade Species Macrobiotus snaresensis Macrobiotus snaresensis Macrobiotus snaresensis Diphascon chilenense Macrobiotus coronatus Diphascon scoticum Hypsibius dujardini Macrobiotus snaresensis Macrobiotus liviae Echiniscus spiniger Isohypsibius sattleri Paramacrobiotus areolatus) Macrobiotus furciger Macrobiotus coronatus Minibiotus intermedius Macrobiotus liviae Macrobiotus snaresensis Milnesium tardigradum Pseudechiniscus novaezeelandiae Diphascon scoticum Macrobiotus recens Macrobiotus snaresensis Milnesium tardigradum Echiniscus bigranulatus Hypechiniscus exarmatus Hypsibius convergens Isohypsibius cameruni

Substrate Comparisons Meyer (2006b) extended the comparison of substrata in Florida, USA, to include not only liverworts, mosses, and foliose lichens, but also ferns. He found 20 species of tardigrades on 47 species of plants and lichens. They found that some species were positively associated with mosses or with foliose lichens, but as in most other studies, there was no association with any particular plant or lichen species. Guil et al. (2009a) reviewed tardigrades and their habitats (altitude, habitat characteristics, local habitat structure or dominant leaf litter type, and two bioclimatic classifications), including bryophytes and leaf litter at various elevations. They were able to show some habitat preference. Species richness was most sensitive to bioclimatic classifications of macroenvironmental gradients (soil and climate), vegetation structure, and leaf litter type. A slight altitude effect was discernible. These relationships suggest that differences among bryophyte species should exist where bryophyte species occupy different environmental types or maintain different microenvironments within a habitat. But it also suggests that within the same habitat, bryophytes of various life forms should provide different moisture regimes, hence creating species relationship differences. In a different study in the Iberian Peninsula (extreme southwestern Europe), Guil et al. (2009b) found that leaf litter habitats showed high species richness and low abundances compared to rock habitats (mosses and lichens), which had intermediate species richness and high abundances. Tree trunk habitats (mosses and lichens) showed low numbers of both richness and abundances. One might conclude that the moisture of these habitats is

Liverwort Species

Plagiochila fasciculata Plagiochila obscura Plagiochila strombifolia Porella elegantula

Tardigrade Species Macrobiotus anderssoni Macrobiotus liviae Macrobiotus recens Macrobiotus snaresensis Diphascon scoticum Macrobiotus furciger Macrobiotus coronatus Macrobiotus liviae Pseudechiniscus juanitae Macrobiotus anderssoni Macrobiotus furciger Doryphoribius zyxiglobus Echiniscus vinculus Diphascon alpinum Diphascon bullatum Diphascon prorsirostre Hypsibius convergens Isohypsibius sattleri Macrobiotus anderssoni Macrobiotus furciger Macrobiotus coronatus Macrobiotus hibiscus Minibiotus intermedius Minibiotus aculeatus Macrobiotus liviae Milnesium tardigradum Pseudechiniscus novaezeelandiae

the overall determining factor, and this should coincide with bryophyte species groups on the large scale. Miller et al. (1996) found six species of tardigrades in lichen and bryophyte samples on ice-free areas at Windmill Islands, East Antarctica. The tardigrade species Diphascon chilenense (see Figure 32), Acutuncus antarcticus (formerly Hypsibius antarcticus; see Figure 33), and Pseudechiniscus juanitae (=Pseudechiniscus suillus; Figure 34) showed a positive association with bryophytes and a negative association with algae and lichens.

Figure 32. Diphascon sp., member of one of the most common bryophyte-dwelling genera. Photo by Martin Mach, with permission.

Meyer and Hinton (2007) reviewed the Nearctic tardigrades (Greenland, Canada, Alaska, continental USA, northern Mexico). They found that one-third of the species occur in both cryptogams (lichens and bryophytes) and soil/leaf litter (Table 5). Few tardigrades occurred exclusively in soil/leaf litter habitats. Although many

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Chapter 5-4: Tardigrades: Species Relationships

occurred among both bryophytes and lichens, 18 species occurred only in bryophytes. It is likely that bryophytes offer a better moisture environment, but this has not been tested.

Figure 33. permission.

Hypsibius.

Photo by Yuuji Tsukii, with

lichen, moss, & stream habitats). Whereas it is not unusual for the soil, lichens, and mosses to have similar fauna and richness, it seems a bit unusual for the stream habitat to be as rich. Amphibolus cf. weglarskae and Diphascon cf. ramazzottii were the only species found only on bryophytes among those four substrates. Horning et al. (1978) collected from soil, fungi, algae, bryophytes, lichens, marine substrata, freshwater substrata, and litter in New Zealand and surrounding islands. From bryophyte and lichen habitats, they found that all 14 of the most abundant species occurred in at least three of the five "plant" categories (three lichen forms, liverworts, and mosses). Among these, the highest occurrence was among mosses. Although Milnesium tardigradum (Figure 16) was slightly more abundant on lichens than on mosses, the combined numbers on mosses and liverworts was still higher. Horning et al. identified the bryophytes and lichens and presented the species of tardigrades on each (Table 1, Table 4, Table 6). In 559 moss samples, 45.8% had tardigrades, mean of 1.8 species, range 1-8 (Table 1). Of 55 species of tardigrades known for New Zealand, 45 occurred on mosses.

Finding New Species

Figure 34. Pseudechiniscus juanitae. Bartels, with permission.

Photo by Paul J.

Table 5. Comparison of tardigrades inhabiting their primary substrates in the Nearctic realm. Only species present on that substrate in at least three sites are included. From Meyer & Hinton 2007.

Substrate category Cryptogams only Both cryptogams and soil/leaf litter Soil/leaf litter only Both bryophyte and lichen Bryophyte only Lichen only

The common appearance of tardigrades among bryophytes causes those who seek to describe new taxa to go first to the mossy habitats. In this spirit, Kaczmarek and Michalczyk (2004a) found the new species of mossdwelling Doryphoribius quadrituberculatus in Costa Rica. From mosses in China they described the new species Bryodelphax brevidentatus (Kaczmarek et al. 2005) and B. asiaticus (Figure 35; Kaczmarek & Michalczyk 2004b), as did Li and coworkers for Echiniscus taibaiensis (Wang & Li 2005), Isohypsibius taibaiensis (Li & Wang 2005), Isohypsibius qinlingensis (Li et al. 2005a), Pseudechiniscus papillosus (Li et al. 2005b), Pseudechiniscus beasleyi, Echiniscus nelsonae, and E. shaanxiensis (Li et al. 2007), and Tumanov (2005) for Macrobiotus barabanovi and M. kirghizicus. Pilato and Bertolani (2005) described Diphascon dolomiticum from Italy.

number of species 64 27 3 50 18 5

Beasley (1990) conducted a similar study in Colorado, USA. Out of 135 samples of liverworts, mosses, lichens, and club mosses (Lycopodiaceae), they found 20 species in 55 samples. There were no tardigrades on liverworts (!), but they were on 46% of mosses and 43% of lichens. The big surprise is that 75% of the clubmosses had tardigrades. In the Great Smoky Mountains National Park, Bartels and Nelson (2006) found that the number of species differed little among the substrates they sampled (soil,

Figure 35. Bryodelphax asiaticus. Photo through Creative Commons.

Chapter 5-4: Tardigrades: Species Relationships

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Table 6. Comparison of numbers of individuals and percentage of individuals of each of 14 tardigrade species on liverworts, mosses, and lichens in collections from New Zealand and surrounding islands. The remaining ones were on other non-plant substrata. Number of samples is in parentheses. From Horning et al. 1978.

Pseudechiniscus novaezeelandiae Pseudechiniscus juanitae Macrobiotus harmsworthi Macrobiotus hibiscus Minibiotus intermedius Milnesium tardigradum Hypsibius dujardini Paramacrobiotus areolatus Echiniscus bigranulatus Hypechiniscus gladiator Diphascon scoticum Macrobiotus liviae Macrobiotus anderssoni Macrobiotus furciger

n

liverworts % (150)

mosses % (559)

lichens % (239)

46 43 89 90 65 143 32 58 18 21 35 72 63 89

8.70 6.98 5.62 7.78 7.69 7.69 10.53 3.45 5.56 19.05 11.43 8.33 11.11 12.36

56.50 44.19 55.06 60.00 41.54 35.66 50.00 60.34 38.89 61.90 65.71 56.94 42.86 50.56

23.90 27.91 34.83 17.78 32.30 37.06 2.63 18.97 38.89 9.50 11.43 18.06 22.22 22.47

New species from South Africa are no surprise, as enumeration of small organisms in that country is barely out of its infancy. Kaczmarek and Michalczyk (2004c) described the new species Diphascon zaniewi in the Dragon Mountains there. Other species found there were more cosmopolitan: Hypsibius maculatus (previously known only from Cameroon and England), H. convergens (Figure 36), Paramacrobiotus cf. richtersi (Figure 37), and Minibiotus intermedius (Figure 38-Figure 39).

Figure 38. Minibiotus intermedius. Miller through Flickr.

Photo by William

Figure 36. Hypsibius convergens, a common moss-dweller. Photo by Björn Sohlenius, Swedish Museum of Natural History, with permission.

Figure 39. Minibiotus intermedius mouth. Photo by Łukasz Kaczmarek and Łukasz Michalczyk, with permission.

Figure 37. Paramacrobiotus richtersi, a common bryophyte dweller. Photo by Science Photo Library through Creative Commons.

Likewise, in South America, Michalczyk and Kaczmarek (2005) described Calohypsibius maliki as a new species from Chile; Michalczyk and Kaczmarek (2006) described Echiniscus madonnae (Figure 40) from

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Chapter 5-4: Tardigrades: Species Relationships

Peru, all from bryophytes. In Argentina they described Macrobiotus szeptyckii and Macrobiotus kazmierskii (Kaczmarek & Michalczyk 2009). In 2008 Degma et al. described another new species [Paramacrobiotus derkai (Figure 41)] from Chile, a country where only 29 species had previously been described.

Figure 42. Macrobiotus marlenae. Photo by Kaczmarek and Łukasz Michalczyk, with permission.

Łukasz

Figure 40. Echiniscus madonnae, a moss dweller from Peru. Photo by Łukasz Kaczmarek & Łukasz Michalczyk, with permission.

Figure 43. Echiniscus viridianus. Photo by Paul J. Bartels, with permission.

Summary

Figure 41. Paramacrobiotus derkai emerging from egg. Photo by Łukasz Kaczmarek, with permission.

In Portugal, lichens and mosses provided the new species Minibiotus xavieri to Fontoura and coworkers (2009). In Cyprus, Kaczmarek and Michalczyk (2004d) described Macrobiotus marlenae (Figure 42). Macrobiotus kovalevi proved to be a new species from mosses in New Zealand (Tumanov 2004). Clearly, mosses have been a favorite sampling substrate for tardigrade seekers (Kaczmarek & Michalczyk 2009) and most likely hold many more undescribed species around the world. Even when new species are collected, they are not always identified or diagnosed in a timely manner. This can result in their ultimate description from multiple locations. Such is the case for Echiniscus viridianus (Figure 43), a new species described by Pilato et al. (2007) from Alabama and New Mexico, USA, and from the Azores Islands, all from mosses.

Most studies indicate no correlation between bryophyte species and tardigrade species. There is limited indication that cushions may have more species, but in other studies thin mats have more than cushions. Other studies indicate they are more common on weftforming mosses than on turfs. Open mosses like Polytrichum seem to be less suitable as homes. There may be some specificity for liverworts rather than mosses, as for example Macrobiotus snaresensis in New Zealand. Unfortunately, many researchers have not identified the bryophyte taxa in tardigrade faunistic studies. A common garden study including several bryophyte species and tardigrades of the same or different species could be revealing.

Acknowledgments Roberto Bertolani provided an invaluable update to the tardigrade taxonomic names and offered several suggestions on the text to provide clarification or correct errors. Łukasz Kaczmarek has provided me with references, images, contact information, and a critical read of an earlier version of the text. Martin Mach and Yuuji Tsukii have given permission to use images that illustrate the species. Michael Lüth has given permission to use his many bryophyte images, and my appreciation goes to all

Chapter 5-4: Tardigrades: Species Relationships

those who have contributed their images to Wikimedia Commons for all to use. Martin Mach was kind enough to send me corrections for typos in the previous online version. Thank you to my sister, Eileen Dumire, for providing the view of a novice on the readability of the original text (it has been much expanded since then). Tardigrade nomenclature is based on Degma et al. 2010.

Literature Cited Bartels, P. J. and Nelson, D. R. 2006. A large-scale, multihabitat inventory of the phylum Tardigrada in the Great Smoky Mountains National Park, USA: A preliminary report. Hydrobiologia 558: 111-118. Baxter, W. H. 1979. Some notes on the Tardigrada of North Down, including one addition to the Irish fauna. Irish Nat. J. 19: 389-391. Beasley, C. W. 1990. Tardigrada from Gunnison Co., Colorado, with the description of a new species of Diphascon. Southw. Nat. 35: 302-304. Beasley, C. W. and Miller, W. R. 2007. Tardigrada of Xinjiang Uygur Autonomous Region, China. Proceedings of the Tenth International Symposium on Tardigrada. J. Limnol. 66(Suppl. 1): 49-55. Bertolani, R. 1983. Tardigardi muscicoli delle dune costiere Italiane, con descrizione di una nuova specie. Atti Soc. Tosc. Sci. Nat. Mem. Ser. B 90: 139-148. Bertrand, M. 1975. Répartition des tardigrades "terrestres" dans le massif de l'Aigoual. Vie Milieu 25: 283-298. Christenberry, D. 1979. On the distribution of Echiniscus kofordi and E. cavagnaroi (Tardigrada). Trans. Amer. Microsc. Soc. 98: 469-471. Collins, M. and Bateman, L. 2001. The ecological distribution of tardigrades in Newfoundland. Zool. Anz. 240: 291-297. Degma, P. 2006. First records of two Heterotardigrada (Tardigrada) species in Slovakia. Biologia Bratislava 61: 501-502. Degma, P., Bertolani, R., and Guidetti, R. 2010. Actual checklist of Tardigrada species (Ver. 11:26-01-2010). Accessed 18 June 2010 at < http://www.tardigrada.modena.unimo.it/miscellanea/Actual %20checklist%20of%20Tardigrada.pdf>. Degma, P., Simurka, M., and Gulanova, S. 2005. Community structure and ecological macrodistribution of moss-dwelling water bears (Tardigrada) in Central European oak-hornbeam forests (SW Slovakia). Ekologia (Bratislava) 24(suppl. 2): 59-75. Degma, P., Michalczyk, Ł., and Kaczmarek, Ł. 2008. Macrobiotus derkai, a new species of Tardigrada (Eutardigrada, Macrobiotidae, huziori group) from the Colombian Andes (South America). Zootaxa 1731: 1-23. Fontoura, P., Pilato, G., Morais, P., and Lisi, O. 2009. Minibiotus xavieri, a new species of tardigrade from the Parque Biológico de Gaia, Portugal (Eutardigrada: Macrobiotidae). Zootaxa 2267: 55-64. Fullard, H. 1968. China in Maps. George Philip & Sons, Ltd., London, 25 pp. Guil, N., Hortal, J., Sanchez-Moreno, S. and Machordom, A. 2009a. Effects of macro and micro-environmental factors on the species richness of terrestrial tardigrade assemblages in an Iberian mountain environment. Landscape Ecol. 24: 375390.

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Guil, N., Sanchez-Moreno, S., and Machordom, A. 2009b. Local biodiversity patterns in micrometazoans: Are tardigrades everywhere? System. Biodiv. 7: 259-268. Hinton, J. G. and Meyer, H. A. 2007. Distribution of limnoterrestrial Tardigrada in Georgia and the Gulf Coast states of the United States of America with ecological remarks. In: Pilato, G. and Rebecchi, L. (guest eds.). Proceedings of the Tenth International Symposium on Tardigrada. J. Limnol. 66(Suppl. 1): 72-76. Hinton, J. G. and Meyer, H. A. 2009. Tardigrades from Fayette County, Georgia. Georgia J. Sci. 67(2): 30-32. Hofmann, I. 1987. Habitat preference of the most frequent mossliving Tardigrada in the area of Giessen (Hessen). In: Bertolani, R. (ed.). Biology of Tardigrades. Selected Symposia and Monographs U.Z.I., Mucchi, Modena Italia, pp 211-216. Hofmann, I. and Eichelberg, D. 1987. Faunistisch-oekologische Untersuchungen zur Habitat-praeferenz moosbewohnender Tardigraden. [Ecological investigations of the habitat preference of moss-inhabiting tardigrades.]. Zool. Beitr. 31(1): 61-76. Horning, D. S., Schuster, R. O., and Grigarick, A. A. 1978. Tardigrada of New Zealand. N. Zeal. J. Zool. 5: 185-280. Hunter, M. A. 1977. A study of Tardigrada from a farm in Montgomery County, Tennessee. MS Thesis, Austin Peay State University, Clarksville, TN, 61 pp. Jönsson, K. I. 2003. Population density and species composition of moss-living tardigrades in a boreo-nemoral forest. Ecography 26: 356-364. Kaczmarek, Ł. and Michalczyk, Ł. 2004a. First record of the genus Doryphoribius Pilato, 1969 from Costa Rica (Central America) and description of a new species Doryphoribius quadrituberculatus (Tardigrada: Hypsibiidae). Genus 15: 447-453. Kaczmarek, Ł. and Michalczyk, Ł. 2004b. A new species Bryodelphax asiaticus (Tardigrada: Heterotardigrada: Echiniscidae) from Mongolia (Central Asia). Raffles Bull. Zool. 52: 599-602. Kaczmarek, Ł. and Michalczyk, Ł. 2004c. Notes on some tardigrades from South Africa, with the description of Diphascon (Diphascon) zaniewi sp. nov. (Eutardigrada: Hypsibiidae). Zootaxa 576: 1-6. Kaczmarek, Ł. and Michalczyk, Ł. 2004d. New records of Tardigrada from Cyprus with a description of the new species Macrobiotus marlenae (hufelandi group) (Eutardigrada: Macrobiotidae). Genus 15: 141-152. Kaczmarek, Ł. and Michalczyk, Ł. 2009. Two new species of Macrobiotidae, Macrobiotus szeptyckii (harmsworthi group) and Macrobiotus kazmierskii (hufelandi group) from Argentina. Acta Zool. Cracoviensia, Seria B - Invertebrata 52: 87-99. Kaczmarek, Ł., Michalczyk, Ł., and Degma, P. 2005. A new species of Tardigrada Bryodelphax brevidentatus sp. nov. (Heterotardigrada: Echiniscidae) from China (Asia). Zootaxa 1080: 33-38. Kathman, R. D. and Cross, S. F. 1991. Ecological distribution of moss-dwelling tardigrades on Vancouver Island, British Columbia, Canada. Can. J. Zool. 69: 122-129. Li, X.-C. and Wang, L.-Z. 2005. Isohypsibius taibaiensis sp. nov. (Tardigrada, Hypsibiidae) from China. Zootaxa 1036: 55-60. Li, X.-C., Wang, L.-Z. and Yu, D. 2005a. Isohypsibius qinlingensis sp. nov. (Tardigrada, Hypsibiidae) from China. Zootaxa 980: 1-4.

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Chapter 5-4: Tardigrades: Species Relationships

Li, X.-C., Wang, L.-Z., and Yu, D. 2007. The Tardigrada fauna of China with descriptions of three new species of Echiniscidae. Zool. Stud. 46(2): 135-147. Li, X.-C., Wang, L.-Z., Liu, Y., and Su, L.-N. 2005b. A new species and five new records of the family Echiniscidae (Tardigrada) from China. Zootaxa 1093: 25-33. Meyer, H. A. 2006a. Interspecific association and substrate specificity in tardigrades from Florida, southeastern United States. Hydrobiologia 558: 129-132. Meyer, H. A. 2006b. Small-scale spatial distribution variability in terrestrial tardigrade populations. Hydrobiologia 558: 133-139. Meyer, H. A. 2008. Distribution of tardigrades in Florida. Southeast. Nat. 7: 91-100. Meyer, H. A. and Hinton, J. G. 2007. Limno-terrestrial Tardigrada of the Nearctic realm. J. Limnol. 66(Suppl. 1): 97-103. Michalczyk, Ł. and Kaczmarek, Ł. 2005. The first record of the genus Calohypsibius Thulin, 1928 (Eutardigrada: Calohypsibiidae) from Chile (South America) with a description of the new species Calohypsibius maliki. New Zeal. J. Zool. 32: 287-292. Michalczyk, Ł. and Kaczmarek, Ł. 2006. Revision of the Echiniscus bigranulatus group with a description of a new species Echiniscus madonnae (Tardigrada: Heterotardigrada: Echiniscidae) from South America. Zootaxa 1154: 1-26. Mihelcic, F. 1954/55. Zur Ökologie der Tardigraden. Zool. Anz. 153: 250-257. Mihelcic, F. 1963. Moose als Lebensstätten für Tardigraden. Der Schlern 37: 179-181. Miller, W. R., Miller, J. D., and Heatwole, H. 1996. Tardigrades of the Australian Antarctic Territories: The Windmill Islands, East Antarctica. Zool. J. Linn. Soc. 116: 175-184. Nelson, D. R. 1973. Ecological distribution of tardigrades on Roan Mountain, Tennessee - North Carolina. Ph. D. dissertation, University of Tennessee, Knoxville.

Nelson, D. R. 1975. Ecological distribution of Tardigrada on Roan Mountain, Tennessee - North Carolina. In: Higgins, R. P. (ed.). Proceedings of the First International Symposium on Tardigrades. Mem. Ist. Ital. Idrobiol. Suppl. 32: 225-276. Newsham, K. K., Maslen, N. R., and McInnes, S. J. 2006. Survival of Antarctic soil metazoans at -80ºC for six years. CryoLetters 27(5): 269-280. Pilato, G. and Bertolani, R. 2005. Diphascon (Diphascon) dolomiticum, a new species of Hypsibiidae (Eutardigrada) from Italy. Zootaxa 914: 1-5. Pilato, G., Fontoura, P., and Lisi, O. 2007. Remarks on the Echiniscus viridis group, with the description of a new species (Tardigrada, Echiniscidae). In: Pilato, G. and Rebecchi, L. (guest eds.). Proceedings of the Tenth International Symposium on Tardigrada. J. Limnol. 66(Suppl. 1): 33-39. Ramazzotti, G. and Maucci, W. 1983. The phylum Tardigrada 3rd edition: English translation by C. W. Beasley. Mem. Ist. Ital. Idrobiol. Dott. Marco de Marchi 41: 1B680. Romano, F. A. III., Barreras-Borrero, B., and Nelson, D. R. 2001. Ecological distribution and community analysis of Tardigrada from Choccolocco Creek, Alabama. Zool. Anz. 240: 535-541. Sayre, R. M. and Brunson, L. K. 1971. Microfauna of moss habitats. Amer. Biol. Teacher Feb. 1971: 100-102, 105. Tumanov, D. V. 2004. Macrobiotus kovalevi, a new species of Tardigrada from New Zealand (Eutardigrada, Macrobiotidae). Zootaxa 406: 1-8. Tumanov, D. V. 2005. Two new species of Macrobiotus (Eutardigrada, Macrobiotidae) from Tien Shan (Kirghizia), with notes on Macrobiotus tenuis group. Zootaxa 1043: 3346. Wang, L.-Z. and Li, X.-C. 2005. Echiniscus taibaiensis sp. nov. and a new record of Echiniscus bisetosus Heinis (Tardigrada, Echiniscidae) from China. Zootaxa 1093: 39-45.

Glime, J. M. 2017. Tardigrade Densities and Richness. Chapt. 5-5. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. Ebook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 18 July 2020 and available at .

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CHAPTER 5-5 TARDIGRADE DENSITIES AND RICHNESS TABLE OF CONTENTS Densities and Richness ........................................................................................................................................ 5-5-2 Europe .......................................................................................................................................................... 5-5-4 North America ........................................................................................................................................... 5-5-10 South America and Neotropics .................................................................................................................. 5-5-12 Asia ............................................................................................................................................................ 5-5-12 Africa ......................................................................................................................................................... 5-5-13 Antarctic and Arctic ................................................................................................................................... 5-5-13 Seasonal Variation ............................................................................................................................................ 5-5-15 Patchiness .......................................................................................................................................................... 5-5-17 Summary ........................................................................................................................................................... 5-5-18 Acknowledgments ............................................................................................................................................. 5-5-18 Literature Cited ................................................................................................................................................. 5-5-19

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Chapter 5-5: Tardigrade Densities and Richness

CHAPTER 5-5 TARDIGRADE DENSITIES AND RICHNESS

Figure 1. Echiniscus, the genus with the most species among mosses. Photo by Martin Mach, with permission.

Densities and Richness But just how common are these bryophyte-dwelling tardigrades (Figure 1)? I think the largest reported density I have found in the literature is 22,000 individuals per gram of dry moss (Mathews 1938), but that is an old number and may well have been replaced. These animals seem to be especially adapted for the bryophyte habitat (Jerez et al. 2002), achieving densities as great as 2,000,000 individuals per square meter of Bryum argenteum (Figure 2) (Brusca & Brusca 1990). (Is that greater than 22,000 per gram?) Nelson (2002) reminds us that densities of these animals are highly variable and conditions for optimum development of the population are unknown (see also Kinchin 1994). Factors such as temperature and moisture (Franceschi et al. 1962-1963; Morgan 1977; Briones et al. 1997), air pollution (Steiner 1994a, b, c, 1995), and food availability (Hallas & Yeates 1972) all influence population density. And it appears that random dispersal may be a major factor, since both population density and species diversity vary considerably between adjacent microhabitats that appear to be identical (Nelson 2002).

Figure 2. Bryum argenteum exhibiting the tight leaves that provide capillary spaces where tardigrades can enjoy prolonged water retention. Photo by Michael Lüth, with permission.

Among the more extensive studies is that of Kathman and Cross (1991) on Vancouver Island, British Columbia, Canada. They collected from mosses at six altitudes on five mountains and found 39 species among 37 moss species, with 13,696 individuals in all. However, as noted in Bertolani's (1983) study, the species of moss did not seem to be important.

Chapter 5-5: Tardigrade Densities and Richness

Horning et al. (1978) collected from soil, fungi, algae, bryophytes, lichens, marine substrata, freshwater substrata, and litter in New Zealand and surrounding islands. They provide summaries of the tardigrade species from each bryophyte species. From their 1354 collections, they represented 577 terrestrial habitats. All 14 of the more abundant tardigrade species occurred in at least three of the five "plant" categories (three lichen forms, liverworts, and mosses). Among these, the highest occurrence was among mosses, except for Milnesium tardigradum (Figure 3), which occurred more often among lichens. They reported the number of species on each bryophyte, but not the density of individuals. As in other studies, moisture seemed to play a major role. They considered the "plant" categories, arranged from dry to moist, to be crustose lichen > fruticose lichen > foliose lichen > liverworts & mosses. The foliose lichens and mosses served as habitat for more tardigrade species than did the liverworts, crustose lichens, or fruticose lichens. Liverworts housed 30 tardigrade species on 26 liverwort species.

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Figure 4. Moss-covered roof that has been sampled along the edge. Photo by Susan Moyle-Studlar, with permission.

Figure 5. Ramazzottius oberhaeuseri, a common mossdweller, including those on roofs. Photo by Martin Mach, with permission.

Figure 3. Milnesium tardigradum, a moss dweller that seems to have a slight preference for lichens. Photo by Martin Mach, with permission.

Roof mosses (Figure 4) have their share of tardigrade fauna; Morgan (1977) recorded densities of four tardigrade species [Macrobiotus hufelandi (Figure 15), Milnesium tardigradum (Figure 3), Ramazzottius oberhaeuseri (Figure 5), Echiniscus testudo (Figure 6)] of up to 823 individuals per gram of the mosses Ceratodon purpureus (Figure 7) and Bryum argenteum (Figure 8) on roofs in Swansea, Wales. In total, Morgan collected 32,552 tardigrades from these two mosses on just three roof locations at the University College of Swansea. Even new species might be abundant in many parts of the world. This is an under-collected group, as suggested by finding very common species for the first time in some countries. Kristensen et al. (2009) found more than 200 individuals of a new species of Bryodelphax (see Figure 9) in a "very small moss sample." And these were cohabiting with Macrobiotus hufelandi (Figure 15) and Milnesium tardigradum (Figure 3).

Figure 6. Echiniscus testudo tun on a moss leaf. Photo by Power & Syred through Creative Commons.

Figure 7. Ceratodon purpureus, another common roof moss that can house innumerable tardigrades. Photo by Michael Lüth, with permission.

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Chapter 5-5: Tardigrade Densities and Richness

Figure 10. Hypnum cupressiforme, home of abundant tardigrades. Photo by Michael Lüth, with permission.

Figure 8. Bryum argenteum, a common roof moss that can house innumerable tardigrades. Photo by Michael Lüth, with permission.

Figure 11. Hylocomium splendens, a good habitat for tardigrades. Photo by Janice Glime.

Figure 9. Bryodelphax asiaticus. Photo through Creative Commons.

Europe One might expect the knowledge of European tardigrades to be the most complete, partly because the taxonomy of the bryophytes has been known longer than in many other countries, including North America, and partly because of the interest of Europeans in natural history. Some European mosses have abundant tardigrades: Hypnum cupressiforme (Figure 10), Hylocomium splendens (=Hypnum parietinum) (Figure 11), and Sanionia uncinata (Figure 12), as well as Grimmia (might include Schistidium; Figure 13) and Tortula (Marcus 1928a; probably includes Syntrichia; Figure 14) and may contain up to 20,000 individuals per 1 g of air-dried moss (Marcus 1928b).

Figure 12. Sanionia uncinata, a suitable tardigrade habitat. Photo by Janice Glime.

Chapter 5-5: Tardigrade Densities and Richness

5-5-5

Figure 13. Grimmia elongata cushions. Photo by Michael Lüth, with permission.

Figure 16. Eurhynchium swartzii, a pleurocarpous moss that is known to house tardigrades. Photo by Kristian Peters through Wikimedia Commons.

Figure 14. Tortula intermedia cushion. Photo by Michael Lüth, with permission.

Figure 17. Barbula tophacea, an acrocarpous moss that houses tardigrades. Photo by Michael Lüth, with permission.

In a boreal forest in Sweden, Jönsson (2003) found sixteen species of tardigrades on mosses, including the widespread Macrobiotus hufelandi (Figure 15) as the most common. Among these, five were new to Sweden. They also found that the forest tended to have more tardigrade species than did a clear-cut area, but overall abundance within a species differed little between these two habitats.

Hofmann and Eichelberg (1987) found sixteen species, including two undescribed, among mosses at Lahnau, near Giessen, Germany. Maucci (1980) collected 2686 samples of bryophytes and found 23 species of tardigrades. In Sardinia, Pilato and Sperlinga (1975) likewise found sixteen species of tardigrades among the bryophytes. These included Macrobiotus nuragicus and M. arguei as new species. Isohypsibius pappi, I. sattleri (formerly I. bakonyiensis), and Hypsibius convergens (Figure 18) were new for Sardinia. It seems that finding new species within tardigrade communities is a fairly common occurrence.

Figure 15. Macrobiotus hufelandi, a dominant species on Rhytidiadelphus squarrosus (Figure 19) in the Black Forest, Germany. Photo by Martin Mach, with permission.

In the Tihany Peninsula, Hungary, Felföldy and Iharos (1947) found modest numbers, with 38 individuals per gram of the moss Eurhynchium swartzii (Figure 16) and 84 per gram among clones of Barbula [formerly in Didymodon] tophacea (Figure 17).

Figure 18. Hypsibius convergens, one of the most common of bryophyte dwellers. Photo by Łukasz Kaczmarek and Łukasz Michalczyk, with permission.

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Chapter 5-5: Tardigrade Densities and Richness

Schuster and Greven (2007) followed the inhabitants of the moss Rhytidiadelphus squarrosus (Figure 19) in the Black Forest in Germany for 54 months (Table 1). They uncovered 19,909 individuals comprising 24 species. The dominant species were Macrobiotus hufelandi (Figure 15; 56%), Paramacrobiotus richtersi (Figure 20; 18%), and Diphascon pingue (Figure 21; 12%). In contrast to the Oregon study, the highest diversity occurred in winter, whereas the number of individuals declined in winter, then increased from spring until autumn, as in Oregon. On the other hand, D. rugosum (Figure 22), Hypsibius dujardini (Figure 23), and H. cf. convergens (Figure 18) exhibited peaks in winter. Water-loving species were most numerous in the moist season, whereas euryhydric species increased when it was relatively dry and sunny. During the course of the 54 months, 14 of the 24 species remained, whereas species succession/change occurred among the others.

Figure 19. Rhytidiadelphus squarrosus, the home for 24 rotifer species in The Black Forest of Germany. Photo by Michael Lüth, with permission.

Figure 20. Paramacrobiotus richtersi, one of the most common and abundant of the bryophyte tardigrades. Photo by Science Photo Library through Creative Commons.

Figure 21. Diphascon pingue. Photo by Michael Collins, with permission.

Figure 22. Diphascon rugosum, a tardigrade that peaks in winter in Oregon, USA. Photo by Björn Sohlenius, Swedish Museum of Natural History, with permission.

Figure 23. Hypsibius dujardini, a moss dweller that has its peak population in winter in the Black Forest of Germany. Photo by Bob Goldstein, with permission.

Species such as Diphascon oculatum (Figure 24) that had reasonable numbers on Rhytidiadelphus squarrosus (Figure 19), but for which no eggs were found (Schuster & Greven 2007), might deposit eggs at a different season than those sampled. It is unlikely that they would deposit eggs in a different habitat/location from that of the adults because of their limited mobility. On the other hand, rare species occurring only once, e.g. Mesocrista spitzbergensis (Figure 25) [note – this is a name change from M. spitzbergense, required to make the gender agree with that of the genus (Degma et al. 2010)], may have been an accidental arrival on Rhytidiadelphus squarrosus, or generally rare. It would be interesting to know the longevity and life cycle of rare species.

Figure 24. Diphascon oculatum, an inhabitant of Rhytidiadelphus squarrosus (Figure 19). Photo by Björn Sohlenius, Swedish Museum of Natural History, with permission.

Chapter 5-5: Tardigrade Densities and Richness

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Table 1. Comparison of total number of individuals (in order of dominance), eggs in exuviae, dominances, and frequencies for each tardigrade species collected on Rhytidiadelphus squarrosus (Figure 19) in the Black Forest of Germany within the investigation period of 54 months. Asterisks indicate species found at least once in each year of study. From Schuster & Greven 2007.

Species

N. individuals Eggs / Exuviae

Dominance (%) Frequency (%)

*Macrobiotus hufelandi (Schultze 1833) *Paramacrobiotus richtersi (Murray 1911) *Diphascon pingue sl (Marcus 1936) *Hypsibius scabropygus (Cuénot 1929) *Macrobiotus patiens (Pilato et al. 2000) *Hypsibius dujardini (Doyère 1840) *Diphascon rugosum (Bartos 1935) *Isohypsibius prosostomus (Thulin 1928) *Hypsibius convergens (Urbanowicz 1925) *Hypsibius pallidus (Thulin 1911) *Hypsibius cfr. convergens *Milnesium tardigradum (Doyère 1840) *Diphascon oculatum (Murray 1906) *Diphascon prorsirostre (Thulin 1928) Isohypsibius pappi (Iharos 1966) Hypsibius sp. Diphascon nobilei (Binda 1969) Minibiotus cfr. poricinctus Minibiotus cfr. scopulus Diphascon scoticum (Murray 1905) Minibiotus intermedius (Plate 1888) Diphascon bullatum (Murray 1905) Diphascon higginsi (Binda 1971) Mesocrista spitzbergensis (Richters 1903)

11118 3600 2359 429 403 390 348 294 246 246 164 101 77 63 24 12 8 8 6 5 5 1 1 1

448 179 170 15 7 58 22 29 18 13 8 4 0 1 7 0 0 0 0 0 0 0 0 0

55.84 18.08 11.85 2.15 2.02 1.96 1.75 1.48 1.24 1.24 0.82 0.51 0.39 0.32 0.12 0.06 0.04 0.04 0.03 0.03 0.03 0.01 0.01 0.01

Sum

19909

979

100.00

Figure 25. Mesocrista spitzbergensis, an inhabitant of Rhytidiadelphus squarrosus. Photo by Björn Sohlenius, Swedish Museum of Natural History, with permission.

In Scotland, Morgan (1976) found that bryophyte and lichen dwellers represented the highest number of tardigrades as well as having the greatest species diversity. In Wales, Morgan (1974) found tardigrades numbering 2287x103 m-2 among mosses. By contrast, Hallas and Yeates (1972) found only 12x103 m-2 in soil and litter in Danish forests. Studies on abundance reveal a wide range of densities. Degma and coworkers (2003, 2006; & Pecalková 2003; et al. 2004, 2005) have provided us with records of tardigrades on named species of mosses in Slovakia. On Hypnum cupressiforme (Figure 10) in Central European

100 100 100 78.5 87.9 72.9 48.6 67.3 46.7 65.4 31.8 48.6 41.1 39.3 16.8 2.8 2.8 3.7 5.6 2.8 3.7 0.9 0.9 0.9

oak-hornbeam forests of Slovakia, Degma et al. (2005) found 3050 tardigrades [21 species in two families (Hypsibiidae & Macrobiotidae)] from 79 moss samples. As in many other studies they were unable to demonstrate any of 12 environmental variables that accounted for the distribution of the tardigrades. Rather, they found that the distribution of species was random. Nevertheless, in his 2003 study, Degma found particular tardigrades on particular bryophytes (Figure 26Figure 43): Eremobiotus alicatai on mosses Brachythecium rutabulum (Figure 26) and Eurhynchium hians (Figure 27); Isohypsibius pappi on these two as well as on B. reflexum (Figure 28), Homalothecium sericeum (Figure 29), Hypnum cupressiforme (Figure 10), Mnium stellare (Figure 31), and Rhynchostegium megapolitanum (Figure 32); Isohypsibius josephi on Amblystegium serpens (Figure 33) and Brachythecium starkei (Figure 34); Diphascon iltisi on Campylium halleri (Figure 35); Astatumen trinacriae (Figure 36) on Brachythecium rutabulum (Figure 26), Homalothecium sericeum (Figure 29), Hypnum cupressiforme (Figure 10), Isothecium alopecuroides (Figure 30), Leskeella nervosa (Figure 37), Paraleucobryum longifolium (Figure 38), and Pterigynandrum filiforme (Figure 39); Isohypsibius dastychi in unidentified moss. Degma and Pecalková (2003) reported Diphascon belgicae in Brachythecium reflexum (Figure 28); Calohypsibius schusteri and Itaquascon pawlowskii in Hypnum cupressiforme (Figure 10). In 2006 Degma reported Echiniscus cf. reticulatus on Ctenidium molluscum (Figure 40); Testechiniscus

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spitsbergensis on Ctenidium molluscum (Figure 40), Distichium capillaceum (Figure 41), Ditrichum flexicaule (Figure 42), and Tortella tortuosa (Figure 43). But are these just chance findings, or is there a preference? It is interesting that all but the last three and Paraleucobryum longifolium (Figure 38) are mat-forming mosses. Was this a preference of the tardigrade or the collector? Or simply a consequence of the habitat?

Figure 29. Homalothecium sericeum, a mat-forming moss that is home to Astatumen trinacriae and Isohypsibius pappi. Note the branches turned to one side. Photo by Michael Lüth, with permission.

Figure 26. Brachythecium rutabulum, a mat-forming moss that is home to Astatumen trinacriae, Eremobiotus alicatai, and Isohypsibius pappi. Photo by Michael Lüth, with permission.

Figure 30. Isothecium alopecuroides, home to Astatumen trinacriae and Isohypsibius pappi. Photo by Biopix through EOL Creative Commons.

Figure 27. Eurhynchium hians, a mat-forming moss that is home to Eremobiotus alicatai and Isohypsibius pappi. Photo by Michael Lüth, with permission. Figure 31. Mnium stellares, home to Isohypsibius pappi. Photo by Michael Lüth, with permission.

Figure 28. Brachythecium reflexum, a mat-forming moss that is home to Diphascon belgicae and Isohypsibius pappi. Photo by Michael Lüth, with permission.

Figure 32. Rhynchostegium megapolitanums, home to Isohypsibius pappi. Note the droplets of water adhering to the leaves, making this a good limnoterrestrial habitat. Photo by Michael Lüth, with permission.

Chapter 5-5: Tardigrade Densities and Richness

Figure 33. Amblystegium serpens, home to Isohypsibius josephi. Photo by Michael Lüth, with permission.

Figure 34. Brachythecium starkei, home to Isohypsibius josephi. Photo by Michael Lüth, with permission.

Figure 35. Campylium halleri, home to Diphascon iltisi. Photo by Michael Lüth, with permission.

Figure 36. Astatumen trinacriae. Photo by Paul J. Bartels, with permission.

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Figure 37. Leskeella nervosa, home to Astatumen trinacriae. Note the bulbils at the tips of branches. Photo by Michael Lüth, with permission.

Figure 38. Paraleucobryum longifolium, a cushion former on rocks, home to Astatumen trinacriae. Photo by Michael Lüth, with permission.

Figure 39. Pterigynandrum filiforme, home to Astatumen trinacriae. Photo by Michael Lüth, with permission.

Figure 40. Ctenidium molluscum, home to Echiniscus cf. reticulatus and Testechiniscus spitsbergensis. Photo by Michael Lüth, with permission.

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It is clear that neglect of the bryophyte habitat is neglect of tardigrades in general. Based on species-area curves, Bartels and Nelson (2007) estimated the greatest species richness among bryophytes in their comparison of habitats in the Great Smoky Mountains, USA, although their actual numbers showed about equal numbers of species among the terrestrial habitats: Aquatic Soil Lichen Moss Total Figure 41. Distichium capillaceum, a cushion former, home to Testechiniscus spitsbergensis. Photo by Michael Lüth, with permission.

Figure 42. Ditrichum flexicaule, exhibiting tight vertical leaves, home to Testechiniscus spitsbergensis. Photo by Michael Lüth, with permission.

Figure 43. Tortella tortuosa, a cushion former, home to Testechiniscus spitsbergensis. Photo by Michael Lüth, with permission.

North America The neglect of tardigrades has not escaped North America. Meyer (2006a) lamented that only one species of tardigrade had been reported from Florida. By sampling 47 species of mosses, liverworts, lichens, and ferns from trees and shrubs in all 67 counties of Florida, he found 20 species of tardigrades. Like other studies discussed here, he could find no association between tardigrade species and any particular bryophyte or lichen species. He did, however, find differences between species occurring on lichens and mosses in general.

29 39 35 37 140

Among the additional species most likely to contribute to the predicted number of bryophyte dwellers are a number of species found there on other substrata, that are known from bryophytes elsewhere but not found in the necessarily limited sampling in this study. Meyer et al. (2003) examined populations among a variety of habitats in central Florida and Ouichita Mountains, Arkansas, USA. They found the tardigrades to be both diverse and abundant, varying greatly within the same species among mosses on different rocks and trees. For example, in an extreme case a tree exhibited three species with numerous individuals while the adjacent tree had none. Four adjacent cores yielded from 0 to 86 individuals, totalling 5 species. This type of distribution is consistent with the patchiness discussed below and supports the hypothesis of random dispersal followed by aggregation resulting from reproduction without migration. Paul Davison (pers. comm. 21 June 2006), working in Alabama, USA, contends that tardigrades are best found on "scrappy mosses" that occur in harsh environments. These include those on the face of concrete steps or rock and concrete walls, rooftops, or bark of city trees. In fact, some researchers have suggested that the tardigrades might require a dry period during their lives to survive. Using such mosses, drying, and crumbling them through a 0.5 cm screen over a dish pan can yield as many as 70 tardigrades in just 5 mL of processed extract. A more modest flora was in evidence in the collections from Southwestern Virginia, USA (Riggin 1962). In 434 collections of mosses and lichens, Riggin found only 694 individual tardigrades – hardly a story of high densities on a broad scale. These were represented by 26 species. Macrobiotus seems to be among the most common genera on bryophytes, including North American collections where Riggin found 63% of the Virginia bryophyte (moss?) and lichen collections housing members of this genus. In a study of both the Upper and Lower Peninsulas of Michigan, USA, Meyer et al. (2011) revealed 28 species of tardigrades from mosses, liverworts, lichens, and leaf litter, of which 19 were from bryophytes [Echiniscus blumi, E. merokensis, E. virginicus, E. wendti, Pseudechiniscus facettalis, P. suillus (Figure 44), Milnesium tardigradum (Figure 3), Hypsibius arcticus (Figure 45), Ramazzottius baumanni, R. oberhaeuseri (Figure 5), Diphascon alpinum, D. nodulosum (Figure 46), Astatumen trinacriae (Figure 36), Macrobiotus echinogenitus, M. hufelandi (Figure 15), Minibiotus intermedius (Figure 47), Fractonotus caelatus, Paramacrobiotus areolatus (Figure 48), P. tonollii (Figure 49)]. Of the 28, 18 species were considered to be cosmopolitan. They found only one new

Chapter 5-5: Tardigrade Densities and Richness

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species, and it was not a bryophyte dweller. Although Ramazzotti and Maucci (1983) reported that more than ten taxa of tardigrades can often occur in a single bryophyte sample, and the range is generally 2-6, Meyer et al. found diversity on Michigan bryophytes to usually be at the lower end of this range.

Figure 47. Minibiotus intermedius. Miller through Flickr. Figure 44. Pseudechiniscus juanitae. Bartels, with permission.

Photo by William

Photo by Paul J.

Figure 48. Paramacrobiotus [=Macrobiotus] areolatus. Photo by Martin Mach, with permission.

Figure 45. Hypsibius arcticus. Photo from Smithsonian Institution through EOL Creative Commons.

Figure 49. Paramacrobiotus tonollii. Bartels, with permission.

Figure 46. Diphascon nodulosum. Collins, with permission.

Photo by Michael

Photo by Paul J.

Nelson and Hauser (2012) collected epiphytic mosses and liverworts in a natural area in Oregon, USA. Out of 1102 invertebrates collected, the tardigrades ranked second, exceeded only by the mites (Acari). They pointed out the need for water sampling (washing samples) to find tardigrades. These animals did not show up in the Berlese

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Chapter 5-5: Tardigrade Densities and Richness

extraction used by many collectors. Their collections reveal at least six or seven different taxa of tardigrades from each epiphytic moss water sample, a number that brings the patchy distribution of tardigrades into question. They considered the tardigrades to be well represented for a group with approximately 1000 species, compared to mites with approximately 50,000 species.

species could not be identified. The best known bryophyte dweller among these was Milnesium tardigradum (Figure 54). Echiniscus testudo (Figure 6) was found among the greatest number of bryophyte species. The majority of species were in the Heterotardigrada, possibly due to the higher elevation of the samples and the arid nature of the habitats.

South America and Neotropics Numbers of species and density varies widely among tardigrade collections. Claps et al. (2009) found 28 species in 10 genera in a sub-Antarctic Nothofagus forest (18) and plateau (13) in the Rio Negro province of Argentina. In Costa Rica, Kaczmarek et al. (2009, 2011) found more than 7000 tardigrade individuals in 700 samples of lichens, mosses, and liverworts. These comprised 64 species in 18 genera, but the average number of species per sample was not more than three. They found altitude to be an important factor in distribution, with the highest diversity in the range of 1400-2000 m asl (35 species, 55% frequency). Only 18 species (28% frequency) occurred in the range of 2400-2800 m asl. In the range of 2000-2400 m asl the number of individuals was high. Then at 3200 m asl the frequency (70%) and abundance increased again. Surprisingly, they found a significantly higher presence in the urban and agricultural habitats than they did in natural habitats. Although 24 species had very defined habitat preferences, with the highest frequency in humid habitats, substrate and plant type were not important in their habitat choice. Asia Unfortunately, much of the Asian literature is lost to the western world because of our lack of skill in reading the languages. But according to Beasley et al. (2006), the knowledge of tardigrades in China is meager. And ecological studies seem to be totally wanting. Many of the studies are simply reports of collections made by outsiders (e.g. Mathews 1937a, b; Bartos 1963; Pilato 1974; Beasley et al. 2006). Pilato (1974) found six species of tardigrades in Chinese bryophyte communities and identified three new species: Bryodelphax [=Echiniscus] sinensis, Macrobiotus mandalaae, and Macrobiotus mauccii. Yang (2002) reported on tardigrades from bryophytes in Yunnan Province. Beasley et al. (2006) reported only 18 species from a wide geographic range (3 provinces) in China, with 12 of these species occurring on mosses [Echiniscus nepalensis, Pseudechiniscus jiroveci, Murrayon hibernicus, Hypsibius pallidus, Isohypsibius sattleri, Doryphoribius flavus, Diphascon pingue (Figure 21), Diphascon scoticum (Figure 50), Diphascon prorsirostre, Mesocrista spitsbergensis (Figure 51), Platicrista angustata (Figure 52), Milnesium tardigradum (Figure 3)] and 1 on a liverwort [Cornechiniscus lobatus (see Figure 53)]. Of the 18 species reported, 8 were new to China! It is likely that a much larger fauna exists but has not been explored – or translated. In 2007, Beasley and Miller published a list of tardigrades from Xinjiang Uygur Autonomous Region, China, based on bryophyte specimens from the Missouri Botanical Garden. They found only 78 tardigrades among the 270 specimens of bryophytes, comprising 12 species. Of these 12, 7 were new to China. Several additional

Figure 50. Diphascon scoticum. Kaczmarek, with permission.

Photo by Łukasz

Figure 51. Mesocrista spitsbergensis. Photo by Björn Sohlenius Swedish Museum of Natural History, with permission.

Figure 52. Platicrista angustata, a species that occurs on mosses in China. Photo by Michael Sullivan, with permission.

International knowledge of the Japanese tardigrade fauna suffers from the same language barrier. Mathews, who also named a number of Chinese taxa, reported on the Japanese tardigrades in 1936/37. More recently, Ito (1999) made an ecological study on the north slope of Mt. Fuji,

Chapter 5-5: Tardigrade Densities and Richness

sampling soil, mosses, and lichens. The number of soil tardigrades ranged 8,050 m-2 to 75,500 m-2. Their density was as high as the density of soil arthropods such as mites (Acari) and springtails (Collembola). A few of these showed a relationship with altitude (950-2380 m asl), but typically the dominant species for a habitat did not change much among locations. On the other hand, they changed considerably between habitats at a single location.

Figure 53. Cornechiniscus cornutus. Mach, with permission.

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Schirmacher Oasis, Mitra (1999) examined 36 sites and found only two tardigrade species. Here they are also patchily distributed, nevertheless usually having the highest densities among these three groups of organisms. The ubiquitous and very common moss inhabitant, Macrobiotus sp., is present there, on the sub-Antarctic Marion Island (McInnes et al. 2001). Other tardigrades present include Milnesium cf. tardigradum (Figure 54) and Echiniscus sp. (Figure 55). Gut analysis of M. tardigradum revealed the presence of bdelloid rotifers and even other tardigrades (Diphascon sp.). Sohlenius and Boström (2006) also noted predation by tardigrades on rotifers in East Antarctica. On the nunataks (mountain peaks that penetrate the ice sheet) in continental Antarctica, distribution of tardigrades is patchy, with the greatest abundance occurring within moss cushions and guano (accumulated excrement of seabirds and bats) from bird colonies (Swedish Museum of Natural History 2009). Nine tardigrade taxa have been identified in the Swedish studies.

Photo by Martin

The Japanese certainly have not ignored the tardigrades. They have made important contributions to the physiology (Horikawa & Higashi 2004; Horikawa et al. 2006) and space biology (Horikawa 2008; Ono et al. 2008) of these organisms. There are also good studies on the ecology of soil species. But ecological studies on bryophyte-dwelling taxa are hard to find. Africa

Figure 54. Milnesium tardigradum, a cosmopolitan moss inhabitant. Photo by Yuuji Tsukii, with permission.

Although little is known about them, Africa sports its share of moss-dwelling tardigrades. Pilato and Pennisi (1976) reported 21 species of tardigrades among the mosses in their collections from Cyrenaica (eastern coast of Libya), two of which represented the first members of their genera in Africa. A third, Isohypsibius brulloi, was a new species. Binda (1984) found thirteen species of mossdwelling tardigrades in South Africa and Mozambique. Meyer and Hinton (2009) found only nine species of tardigrades among mosses and lichens in KwaZulu-Natal, South Africa, bringing the total number of species from soil, mosses, and lichens to 61 in southern Africa. But aside from species records, tardigrade-bryophyte ecological studies seem to be rare or non-existent for Africa.

Figure 55. Echiniscus, a ubiquitous genus that occurs on mosses in the Antarctic. Photo by Martin Mach, with permission.

Antarctic and Arctic Unlike Asia, Africa, and South America (McInnes 1994), tardigrades are fairly well studied in polar climates, especially in the Antarctic. In the Antarctic, bryophytes, as well as lichens and algae, provide important habitats for tardigrades, rotifers, and nematodes (Utsugi & Ohyama 1991; Sohlenius et al. 2004). Most invertebrates decrease in abundance as one approaches the poles, but Jennings (1979) found that tardigrades actually increase in abundance in the Antarctic tundra. Peters and Dumjahnn (1999) found 15 species in ten genera in their 249 cushion moss samples from Disko Island, West Greenland. On the other hand, in his moss studies on the Antarctic

On Signy Island off the coast of Antarctica, Jennings (1979) found five species of tardigrades that occurred at both of the sampling sites: Echiniscus capillatus, E. meridionalis, Hypsibius dujardini (Figure 23), Diphascon alpinum, Diphascon pingue sensu lato (Figure 21; or may be Diphascon polare, D. dastychi, or D. victoriae), and Macrobiotus furciger (Figure 56). Other less common taxa were Diphascon scoticum (Figure 50), Isohypsibius renaudi (Figure 57), and Isohypsibius asper (Figure 58). Jennings conducted sampling for two years and found maximum populations of 309x103 m-2 in moss communities of Polytrichum strictum - Chorisodontium aciphyllum (Figure 59-Figure 61). In the Calliergidium austro-

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Chapter 5-5: Tardigrade Densities and Richness

stramineum – Calliergon sarmentosum – Sanionia uncinata communities (Figure 12; Figure 62; Figure 63) they found a maximum of 71x103 m-2. Reproductive potential is high, with increases of 3- to 4-fold in a single year. Hallas and Yeates (1972) found they could reach as high as 10- to 20-fold increases. Echiniscus increased 100-fold at one Signy Island site (Jennings 1979).

Figure 59. Polytrichum strictum and Chorisodontium aciphyllumn in the Antarctic, where Jennings (1979) found 309x103 tardigrades per m2. Photo by Tim Hooker, with permission. Figure 56. Macrobiotus furciger. Photo by Smithsonian Institution through EOL Creative Commons.

Figure 57. Isohypsibius renaudi. Creative Commons.

In a different study on Wilkes Land, East Antarctica, Petz (1997) found tardigrades in more than 74% of the collections of fellfield mosses. These were the most abundant of the invertebrates, with 4,607 in just one gram of moss. Rotifers were the most abundant in other habitats. Ottesen and Meier (1990) likewise found that tardigrades were more abundant among mosses on South Georgia, compared to other habitats.

Photo through EOL

Figure 60. Chorisodontium aciphyllumn in the Antarctic. Photo by Tim Hooker, with permission.

Figure 58. Isohypsibius asper. Photo by Smithsonian Institution through EOL Creative Commons.

In their Antarctic study, Utsugi and Ohyama (1989) found five species of tardigrades in 15 out of 31 samples from Ongul Island, Langhovde, Skarvsnes, Einstoingen, and Rundvagshetta, including algae, lichens, and mosses. Hypsibius arcticus (Figure 45) was common in all their samples. The other four species were rare.

Figure 61. Polytrichum strictum, a moss habitat in the Antarctic and other cool, wet areas. Photo by Michael Lüth, with permission.

Chapter 5-5: Tardigrade Densities and Richness

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Seasonal Variation

Figure 62. Calliergon sarmentosum, of the Calliergidium austro-stramineum – Calliergon sarmentosum – Sanionia uncinatus association in the Antarctic. Photo by Michael Lüth, with permission.

Densities may vary with seasons (Figure 66). Hypsibius convergens (Figure 18) exhibits temporal variation in pool and meadow moss habitats (Marcus 1929). In city mosses, the numbers of individuals of Macrobiotus hufelandii (Figure 15) and Pseudechiniscus pseudoconifer correlated with meteorological factors during a 3-month winter/early spring study (Franceschi et al. 1962-63). It appears that Echiniscus (Figure 55) and its segregate genera may commonly have seasonal variations. Jennings (1979) found that Echiniscus (possibly considered a segregate genus now) was the only tardigrade with seasonal variation among the eight species in his Signy Island study. This is at least in part a reflection of changes in moisture. As already seen for Diphascon rugosum (Figure 22), Hypsibius dujardini (Figure 23), and Hypsibius cf. convergens (Figure 18), there were clear population peaks in winter in a carpet of the soil moss Rhytidiadelphus squarrosus (Figure 19) in the Black Forest, Germany (Schuster & Greven 2007). Species diversity and evenness was generally higher for the tardigrade communities in winter and least in summer (Figure 64). On the other hand, Macrobiotus hufelandi (Figure 15), Diphascon pingue (Figure 21), and to a lesser degree Paramacrobiotus richtersi (Figure 20), declined in winter, increasing in spring through fall (Figure 65). Macrobiotus hufelandi had its peaks in summer and lows in January (Schuster & Greven 2007), as shown for total tardigrades by Merrifield and Ingham (1998), but the other major species did not follow that pattern (Schuster & Greven 2007).

Figure 63. Sanionia uncinata, a cosmopolitan moss that provides tardigrade habitat in the Antarctic. Photo by Jan-Peter Frahm, with permission.

Figure 64. Seasonal changes in number of species of tardigrades found in Rhytidiadelphus squarrosus (Figure 19) clumps. (n = 108). Redrawn from Schuster & Greven 2007.

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Chapter 5-5: Tardigrade Densities and Richness

Figure 65. Seasonal changes in number of individuals of the dominant tardigrades found in Rhytidiadelphus squarrosus (Figure 19). Paramacrobiotus richtersi shows a trend of decline during the sampling years, as shown by the regression line. Modified from Schuster and Greven 2007.

Using a Baermann funnel (Merrifield & Ingham 1998), Merrifield (1992) reported 5 tardigrades per gram on Eurhynchium oreganum (Figure 67) in Oregon, USA, from April to August, with an increase to 15 in September and October, then a crash to 1 for winter months of November through March (Figure 66). Were the bears hibernating elsewhere, or were numbers crashing in the damp Oregon winter?

Figure 66. Seasonal changes in numbers of tardigrades on mosses at Mary's Peak, Oregon, USA. Redrawn from Merrifield & Ingham 1998.

Figure 67. Eurhynchium oreganum, a non-winter habitat for tardigrades. Photo from University of British Columbia bryophyte website, with permission.

Chapter 5-5: Tardigrade Densities and Richness

Romano et al. (2001) attempted to determine the seasonal effects on tardigrades among mosses along Choccolocco Creek, Alabama, USA. They surveyed mosses on three trees each in six sites for 18 months and found no correlation between occurrence and season. However, they did find seasonal differences in the number of species and abundance when they pooled samples.

Patchiness A number of studies suggest that the distribution of tardigrades within a given area or on a particular type of substrate is patchy. Degma et al. (2005) actually did both cluster analysis and CCA, demonstrating that most of the differences in species diversity were the result of randomly found species and that colonization of any given substrate is a random process. It would appear that the greatest determining factor in their specific location and species composition is their dispersal to that location, a process that is as random as it is for the mosses and liverworts they sit on. Further support for this randomness is their random distribution among populations of the moss Hypnum cupressiforme (Figure 68), supported by a Chi-square goodness of fit test.

Figure 68. Hypnum cupressiforme, a ubiquitous moss that seems suitable for many taxa of tardigrades. Photo by Michael Lüth, with permission.

Degma et al. (2009, 2011) found that the horizontal distribution of the tardigrades on a moss clump is aggregated, but that aggregation is not related to moisture in the moss cushion. They hypothesized that once a tardigrade arrives through random recruitment it is able to establish a micro-population. From that beginning slow radiation occurs. The result is that large substrates have more tardigrades but some parts of these larger patches will lack tardigrades while other parts will house aggregations. They continued their study (Degma et al. 2011) using Hypnum cupressiforme (Figure 68) with a 5x5 matrix of circular plots and determined that there was no significant moisture gradient along that moss slope. Nevertheless, the tardigrades existed in clumps or patches. With a large number of individuals (224) in seven species [Milnesium tardigradum (Figure 3), Hypsibius convergens (Figure 18), H. microps, Diphascon pingue (Figure 21), Astatumen trinacriae (Figure 36), Macrobiotus hufelandi (Figure 15), Minibiotus sp. (Figure 47)], they found that species number was random, but that species distribution

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was aggregated. That aggregated distribution was NOT related to moisture in the moss mat. They concluded, therefore, that the best hypothesis to explain the patchy distribution of the tardigrades within the moss cushion was that recruitment of eggs and specimens on the moss was random and that these recruits subsequently reproduced, creating micro-populations where density gradually increased over time. This hypothesis makes the assumption that tardigrades migrate little from the location of their birth. Following this reasoning on a larger scale would account for the patchy distribution observed on larger moss clumps. Larger patches of mosses are more likely to be the recipients of dispersed tardigrades or their eggs and hence are more likely to have tardigrades than would small patches. This would also account for the high degree of variation encountered in random sampling from various moss cushions in the same location. While the individuals are aggregated, the aggregations are random. Meyer (2006b) did a careful study on the spatial variability of tardigrade populations among moss patches on trees and rocks at three locations in the USA. He examined the fauna on patches ranging 0.1 to >5 cm2. He found very high variation among the patches. One interesting discovery was that very small patches rarely had tardigrades. Could it be that they did not retain moisture long enough, or was it a matter of dispersal, with small patches having endured too short a time for colonization to be common? Perhaps it is predictable that patchiness would characterize Antarctic moss dwellers. In the Antarctic, bryophytes, as well as lichens and algae, provide an important habitat for tardigrades, rotifers, and nematodes (Utsugi & Ohyama 1991; Sohlenius et al. 2004). Here tardigrades are also patchily distributed, nevertheless having the highest densities among these three groups of organisms. One might assume that bryophytes must arrive first, or that the tardigrades arrive with their bryophyte home. Hence, dispersal to the continent and its remote islands most likely plays a major role in their location. Studies by the Swedish Museum of Natural History (2009) likewise found patchy distribution of tardigrades on the nunataks of the Antarctic. These windswept peaks emerge above the ice sheets and provide the substrate needed for bryophytes, lichens, and inhabiting tardigrades. Moss cushions and humus enriched by bird colonies provided the greatest numbers of tardigrades, with 400 samples yielding only nine tardigrade taxa. Nevertheless, 32% of the samples had tardigrades (Sohlenius & Boström 2006). The importance of the stochastic process of colonization is supported by the presence of different developmental stages in various samples, suggesting that dispersal may be a dynamic, albeit random, process occurring constantly on the windy peaks. Further population control may exist through competition with the co-occurring nematodes, whereas it appears that the poor rotifers serve as dinner for at least some of these tardigrades. Bettis (2008) tested differences in tardigrade distribution on Grimmia (Figure 69) on exposed granitic outcrops vs protected seasonally riparian forms in California, USA. Again, the distribution was "very patchy" and did not support the hypothesis that more tardigrades would be on the more protected, more moist mosses.

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Chapter 5-5: Tardigrade Densities and Richness

UNT, and since then we have been collaborating. As a result, I “rearranged my lab” to add microscopes, dissecting scopes and cameras to mammal traps, camera traps, mist nests and binoculars. This triggered my enthusiasm to collect many bryophytes/moss samples throughout southern Chile from which I am “hunting” tardigrades to study their biogeography and habitat associations. I also have one graduate student working on tardigrade biotic homogenization.

Summary

Figure 69. Grimmia laevigata in Europe. Photo by Michael Lüth, with permission.

Both Meyer (2006b) (in the terrestrial system) and Romano et al. (2001) (in the aquatic system), emphasized the importance of accounting for this patchiness in designing a sampling strategy. Meyer suggested that the variability of a given location should be understood before determining the number of samples to take. Romano emphasized the need for a greater sampling effort. In short, it appears that the major factor accounting for tardigrade distribution and patchiness is dispersal. If the tardigrade lands in an appropriate habitat, it is able to withstand considerable environmental variation there, and the habitat itself seems to offer little to discriminate against any tardigrade species. Rather, factors like reproductive potential may play the greater role in determining the abundance, and possibly even the diversity, once the tardigrades arrive. I was excited to make new friends among newcomers to the bryophyte-tardigrade relationship. I hope this chapter has inspired more young researchers to include the bryophyte microcosm in their studies. Jaime Jiménez, a vertebrate zoologist, summarizes his conversion to tardigrade study in the box below:

Statement by Jaime Jiménez While working on the seedsnipe and geese feces with Nick [Nick Russo] and Mike [Robertson] on Navarino Island, examining for bryophytes fragments to cultivate, they found a few small water bears. Nick and Mike were my first cohort students from my IRES-NSF grant (Bernard [Goffinet] was the US co-mentor). We were all captivated to see these creature for the first time. It happened that Peter Convey (BAS that used to examine tardigrades from Antarctica) was at the lab during these days and he offered to bring these samples to Cambridge to the tardigrade world expert Sandra McInnes. She put me in contact with William Miller (KS, one of the US tardigrade experts), as she recently reviewed a paper of him on tardigrades found in bird nests. Simultaneously, with Mike, Nick and Sandra we presented a poster in Copenhagen and then in British Columbia (American Ornithological Societies conference). I invited William to

Tardigrades can range in numbers from none to 22,000 individuals per gram of dry moss. More than 2 million are known from a single square meter. On Vancouver Island in Canada, 39 species have been found among bryophytes. They do not seem to prefer any particular moss, and they often occur equally as frequently on lichens and liverworts as they do on mosses. In New Zealand, 30 species are known from liverworts. Macrobiotus and Echiniscus (and their more recent segregates) are among the most abundant tardigrades in the bryophyte fauna. Although most invertebrates decrease in numbers toward the poles, tardigrades actually increase. However, their numbers are highly variable from one place to another. Here, even more so than elsewhere, distribution of the tardigrades is patchy. Even adjacent trees in some localities are known to differ greatly in their tardigrade fauna. Yet, on Wilkes Land in the Antarctic, 74% of the fellfield mosses had tardigrades. They are known to increase up to 100-fold, but it appears that 3- to 4-fold is more typical. Their abundance can be seasonal, with some peaking in winter and others in summer or spring/fall. Some respond to the rainy season. Others don't seem to respond to season. Dispersal plays a large role in both geographic distribution and local patchiness. Within the cushions the tardigrades are often aggregated, but there appears to be no relationship with moisture. On the other hand, small patches seem to lack tardigrades, suggesting that moisture is important. But arrival is a major factor, and from that arrival of one tardigrade, a population develops. Since their movement is slow, they accumulate. But small patches of mosses indicate a short time in which arrival could have occurred.

Acknowledgments Like all of my chapters, this one is really the product of the efforts of many biologists. Roberto Bertolani provided an invaluable update to the tardigrade taxonomic names and offered several suggestions on the text to provide clarification or correct errors. Bryonetters have been wonderful in making their photographs available to me and seeking photographs from others. Paul Davison and Des Callahan have been helpful in providing suggestions and offering images. Bob Goldstein, Bjorn Sohlenius, Martin Mach, and Yuuji Tsukii have given permission to use images that illustrate the species and life

Chapter 5-5: Tardigrade Densities and Richness

cycle stages. Łukasz Kaczmarek has provided me with references and contact information. As always, a big thank you goes to Michael Lüth for permission to use his many images and to all those who have contributed their images to Wikimedia Commons and other public domain sources for all to use. I must admit that this chapter would have been far less interesting without the help of Google to locate the tardigrade stories. I fear I have forgotten some who have helped – I have worked on this chapter for too many years. Nomenclature follows Degma et al. (2010).

Literature Cited Bartels, P. J. and Nelson, D. R. 2007. An evaluation of species richness estimators for tardigrades of the Great Smoky Mountains National Park, Tennessee and North Carolina, USA. Proceedings of the Tenth International Symposium on Tardigrada. J. Limnol. 66(Suppl. 1): 104-110. Bartos, E. 1963. Die Tardigraden der chinesischen und javanischen Mossproben. Vìst. Èsl. Spol. Zool., Acta Soc. Zool. Bohemoslov. 27: 108-114. Beasley, C. W. and Miller, W. R. 2007. Tardigrada of Xinjiang Uygur Autonomous Region, China. Proceedings of the Tenth International Symposium on Tardigrada. J. Limnol. 66(suppl. 1): 49-55. Beasley, C. W., Kaczmarek, Ł. and Michalczyk, Ł. 2006. New records of tardigrades from China, with zoogeographical remarks. Biol. Lett. 2006, 43: 13-20. Bertolani, R. 1983. Tardigardi muscicoli delle dune costiere Italiane, con descrizione di una nuova specie. Atti Soc. Tosc. Sci. Nat. Mem. Ser. B 90: 139-148. Bettis, C. J. 2008. Distribution and abundance of the fauna living in two Grimmia moss morphotypes at the McKenzie Table Mountain Preserve, Fresno County, California. M.S. Thesis, California State University, Fresno, CA, 65 pp. Binda, M. G. 1984. Notizie sui tardigradi dell'Africa Meridionale con descrizione di una nuova specie di Apodibius (Eutardigrada). [Remarks on some species of tardigrades from Southern Africa and description of Apodibius nuntius n. sp.]. Animalia 11(1-3): 5-15. Briones, M. J. I., Ineson, P., and Piearce, T. G. 1997. Effects of climate change on soil fauna; responses of Enchytraeids, Diptera larvae and tardigrades in a transplant experiment. Appl. Soil Ecol. 6: 117-134. Brusca, R. C. and Brusca, G. J. 1990. Invertebrates. Sinauer Associates, Sunderland, Massachusetts. Claps, M., Rossi, G., and Rocha, A. 2009. Tardigrade fauna from two biogeographic regions of Patagonia (Río Negro province, Argentina). In: Greven, H., Hohberg, K., and Schill, R. O. 11th International Symposium on Tardigrada. Conference Guide. Tübingen, Germany, 3-6 August 2009. Degma, P. 2003. First records of six Hypsibiidae species (Tardigrada, Eutardigrada) in Slovakia. Biologia, Bratislava 58: 1003-1005. Degma, P. 2006. First records of two Heterotardigrada (Tardigrada) species in Slovakia. Biologia, Bratislava 61: 501-502. Degma, P. and Pecalková, M. 2003. First records of Tardigrada in Slovakia from Stuzica National Nature Reserve (Bukovské vrchy Mts, NE Slovakia). Biologia, Bratislava 58: 274, 286. Degma, P., Bertolani, R., and Guidetti, R. 2010. Actual checklist of Tardigrada species (Ver. 10: 15-12-2009).

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Accessed on 23 January 2010 at . Degma, P., Gulánová, S., and Simurka, M. 2004. First records of three Hypsibiidae species (Tardigrada: Eutardigrada) in Slovakia. Biologia, Bratislava 59(Suppl 15): 29-11. Degma, P., Katina, S., and Sabatovicová, L. 2009. Horizontal distribution of moisture and Tardigrada in a single moss cushion. In: Greven, H., Hohberg, K., and Schill, R. O. 11th International Symposium on Tardigrada. Conference Guide. Tübingen, Germany, 3-6 August 2009. Degma, P., Simurka, M., and Gulánová, S. 2005. Community structure and ecological macrodistribution of moss-dwelling water bears (Tardigrada) in central European oak-hornbeam forests (SW Slovakia). Ekológia (Bratislava) 24(Suppl 2): 59-75. Degma, P., Stanislav, K., and Sabatovicova, L. 2011. Horizontal distribution of moisture and Tardigrada in a single moss cushion. J. Zool. Syst. Evol. Res. 49 (Suppl 1): 71–77. Felföldy, L. and Iharos, A. 1947. Relation between mossassociation and Tardigrada-fauna on the northern shores of the Tihany Peninsula. Borbasia 7(1-6): 31-38. Franceschi, T., Loi, M. L., and Pierantoni, R. 1962-1963. Risultati di una prima indagine ecologica condotta su popolazioni di Tardigradi. Bollettino dei Musei e degli Istituti Biologici dell'Universita di Genova 32: 69-93. Hallas, T. E. and Yeates, G. W. 1972. Tardigrada of the soil and litter of a Danish beech forest. Pedobiologia 12: 287-304. Hofmann, I. and Eichelberg, D. 1987. Faunistisch-oekologische Untersuchungen zur Habitat-praeferenz moosbewohnender Tardigraden. [Ecological investigations of the habitat preference of moss-inhabiting tardigrades.]. Zool. Beitr. 31(1): 61-76. Horikawa, D. D. 2008. The tardigrade Ramazzottius varieornatus as a model animal for astrobiological studies. Biological Sciences in Space 22: 93-98. Horikawa, D. D. and Higashi, S. 2004. Desiccation tolerance of the tardigrade Milnesium tardigradum collected in Sapporo, Japan, and Bogor, Indonesia. Zool. Sci. 21: 813-816. Horning, D. S., Schuster, R. O., and Grigarick, A. A. 1978. Tardigrada of New Zealand. N. Zeal. J. Zool. 5: 185-280. Iharos, G. 1966. Neue tardigraden-arten aus Ungarn (Neuere beiträge zur kenntnis der tardigraden-fauna Ungarns, VI.). Acta Zool. Acad. Scien. Hung. 12: 111–122. Ito, M. 1999. Ecological distribution, abundance and habitat preference of terrestrial tardigrades in various forests on the northern slope of Mt. Fuji, central Japan. Zool. Anz. 238: 225-234. Jennings, P. G. 1979. The Signy Island terrestrial reference sites: X. Population dynamics of Tardigrada and Rotifera. Brit. Antarct Surv. Bull. 47: 89-105. Jerez Jaimes, J. H., Narváez, E. X., and Restrepo, R. 2002. Tardígrados en musgos de la Reserva el Diviso (Santander, Colombia). Revista Colombiana de Entomología 28(2): 199206. Jönsson, K. I. 2003. Population density and species composition of moss-living tardigrades in a boreo-nemoral forest. Ecography 26: 356-364. Kaczmarek, Ł., Michalczyk, Ł., Gołdyn, B., and Welnicz, W. 2009. Ecological factors determining distribution of Tardigrada in Costa Rican rain forests. In: Greven, H., Hohberg, K., and Schill, R. O. 11th International Symposium on Tardigrada. Conference Guide. Tübingen, Germany, 3-6 August 2009, p. 49.

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Kaczmarek, Ł., Gołdyn, B., Wełnicz, W., and Michalczyk, Ł. 2011. Ecological factors determining Tardigrada distribution in Costa Rica. J. Zool. Syst. Evol. Res. 49 (Suppl 1): 78-83. Kathman, R. D. and Cross, S. F. 1991. Ecological distribution of moss-dwelling tardigrades on Vancouver Island, British Columbia, Canada. Can. J. Zool. 69: 122-129. Kinchin, I. M. 1994. The Biology of Tardigrades. Blackwell Publishing Co., London. 186 pp. Kristensen, R. M., Michalczyk, Ł., and Kaczmarek, Ł. 2009. Tardigrades from Easter Island, Rapa Nui (Pacific Ocean, Chile) with the description of a new species of Bryodelphax (Heterotardigrada: Echiniscidae). In: Greven, H., Hohberg, K., and Schill, R. O. 11th International Symposium on Tardigrada. Conference Guide. Tübingen, Germany, 3-6 August 2009. Marcus, E. 1928a. Spinnentiere oder Arachnoidea IV: Bärtierchen (Tardigrada). Springer, Jena. Marcus, E. 1928b. Zur Ökologie und Physiologie der Tardigraden. Zool. Jahrb. Abt. Allg. Zool. Physiol. Tiere 44: 323-370. Marcus, E. 1929. Tardigrada. In: Bronn, H. G. (ed.). Klassen und Ordnungen des Tierreichs. Section 4, part 3, 5. Akademische Verlagsgesellschaft, pp. 1-608. Mathews, G. B. 1936/1937. Tardigrada from Japan. Pek. Nat. Hist. Bull. 2: 411-412. Mathews, G. B. 1937a. The Tardigrada or water bears. China J. 26: 97-105. Mathews, G. B. 1937b. More tardigrades from the Far East. China J. 27: 32-35. Mathews, G. B. 1938. Tardigrada of North America. Amer. Midl. Nat. 19: 619-627. Maucci, W. 1980. Analisi preliminare di alcuni dati statistici sulla ecologia dei tardigradi muscicoli. [Some statistical data about ecology of moss-dwelling Tardigrada.]. Boll. Mus. Civ. Stor. Nat. Verona 7: 1-47. McInnes, S. J. 1994. Zoogeographic distribution of terrestrial/freshwater tardigrades from current literature. J. Nat. Hist. 28: 257-352. McInnes, S. J., Chown, S. L., Dartnall, H. J. G., and Pugh, P. J. A. 2001. Milnesium cfr. tardigradum (Milnesiidae, Apochela, Tardigrada): A monitor of high altitude meiofauna on subAntarctic Marion Island. Zool. Anz. 240: 461-465. Merrifield, K. 1992. Population dynamics of forest floor mossdwelling nematodes and tardigrades. J. Nematol. 24: 607. Merrifield, K. and Ingham, R. E. 1998. Nematodes and other aquatic invertebrates in Eurhynchium oreganum (Sull.) Jaeg., from Mary's Peak, Oregon Coast Range. Bryologist 101: 505-511. Meyer, H. A. 2006a. Interspecific association and substrate specificity in tardigrades from Florida, southeastern United States. Hydrobiologia 558: 129-132. Meyer, H. A . 2006b. Small-scale spatial distribution variability in terrestrial tardigrade populations. Hydrobiologia 558: 133-139. Meyer, H. A. and Hinton, J. G. 2009. The Tardigrada of Southern Africa, with the description of Minibiotus harrylewisi, a new species from KwaZulu-Natal, South Africa (Eutardigrada: Macrobiotidae). African Invertebrates 50: 255-268. Meyer, H. A., Hinton, J. G., and Trahan, K. 2003. Spatial variability of tardigrade populations in leaf litter, moss, and lichens. Abstracts of the 88th meeting of the Ecological Society of America, Savannah, GA, 3-8 August 2003. Accessed on 3 July 2004, at . Meyer, H. A., Lyons, A. M., Nelson, D. R., and Hinton, J. G. 2011. Tardigrada of Michigan, Northern USA, with the description of Minibiotus jonesorum sp. n. (Eutardigrada: Macrobiotidae). J. Zool. Syst. Evol. Res. 49 (Suppl. 1): 4047. Mitra, B. 1999. Studies on moss inhabiting invertebrate fauna of Schirmacher Oasis. Fifteenth Indian Expedition to Antarctica, Scientific Report, 1999, Department of Ocean Development, Technical Publication No. 13: 93-108. Morgan, C. I. 1974. Studies on the Biology of Tardigrades. Unpublished Ph. D. thesis, University College, Swansea. 220 pp. 2287x103 m-2 in Welsh mosses Morgan, C. I. 1976. Studies on the British tardigrade fauna. Some zoogeographical and ecological notes. J. Nat. Hist. 10: 607-632. Morgan, C. I. 1977. Population dynamics of two species of Tardigrada, Macrobiotus hufelandii (Schultze) and Echiniscus (Echiniscus) testudo (Doyère), in roof moss from Swansea. J. Anim. Ecol. 46: 263-279. Nelson, D. R. 2002. Current status of the Tardigrada: Evolution and ecology. Integrat. Compar. Biol. 42: 652-659. Nelson, J. and Hauser, D. 2012. A survey of invertebrate communities on epiphytic mosses and liverworts in Tryon Creek State Natural Area. Unpublished Undergraduate Report, Lewis & Clark College, Portland, OR. Ono, F., Sigusa, M., Uozumi, T., Matsushima, Y., Ikeda, H., Sainic, N. L., and Yamashita, M. 2008. Effect of high hydrostatic pressure on to life of the tiny animal tardigrade. J. Phys. Chem. Solids 69: 2297-2300. Ottesen, P. S. and Meier, T. 1990. Tardigrada from the Husvik area, South Georgia, sub-Antarctic. Polar Res. 8: 291-294. Peters, T. and Dumjahn, P. 1999. Ecological aspects of tardigrade distribution on Disko Island, West Greenland. In: Brandt, A., Thomsen, H. A., Heide-Jørgensen, H., Kristensen, R. M., and Ruhberg, H. (eds.). The 1998 Danish-German Excursion to Disko Island, West Greenland. Ber. Polarforsch. 330: 64-75. Petz, W. 1997. Ecology of the active soil microfauna (Protozoa, Metazoa) of Wilkes Land, East Antarctica. Polar Biol. 18: 33-44. Pilato, G. 1974. Tre nuove specie di Tardigradi muscicoli di Cina. [Three new species of Tardigrada from mosses in China.]. Animalia 1(1-3): 59-68. Pilato, G. and Pennisi, G. 1976. Prime notizie sui Tardigradi di Cirenaica. [First account on Tardigrada of Cyrenaica.]. Animalia 3(1-3): 243-258. Pilato, G. and Sperlinga, G. 1975. Tardigradi muscicoli di Sardegna. [Moss-inhabiting Tardigrada from Sardinia.]. Animalia 2(1-3): 79-90. Ramazzotti, G. and Maucci, W. 1983. II. Phylum Tardigrada. 3rd ed. Mem. 1st. Ital. Idrobiol. Dott. Marco di Marchi, Vol. 41. Riggin, G. T. Jr. 1962. Tardigrada of Southwest Virginia: With the Addition of a Description of a New Marine Species from Florida. Technical Bull. 152, Virginia Agricultural Experiment Station, Blacksburg, VA. 145 pp. Romano, F. A. III., Barreras-Borrero, B., and Nelson, D. R. 2001. Ecological distribution and community analysis of Tardigrada from Choccolocco Creek, Alabama. Zool. Anzeiger 240: 535-541. Schuster, R. and Greven, H. 2007. A long-term study of population dynamics of tardigrades in the moss

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Rhytidiadelphus squarrosus (Hedw.) Warnst. J. Limnol, 66(Suppl. 1): 141-151. Sohlenius, B. and Boström, S. 2006. Patch-dynamics and population structure of nematodes and tardigrades on Antarctic nunataks. Eur. J. Soil Biol. 42(suppl. 1): S321S325. Sohlenius, B., Boström, S., and Jönsson, K. I. 2004. Occurrence of nematodes, tardigrades and rotifers on ice-free areas in East Antarctica. Pedobiologia 48: 395-408. Steiner, W. 1994a. The influence of air pollution on mossdwelling animals: 1. Methodology and composition of flora and fauna. Rev. Suisse Zool. 101: 533-556. Steiner, W. A. 1994b. The influence of air pollution on mossdwelling animals: 2. Aquatic fauna with emphasis on Nematoda and Tardigrada. Rev. Suisse Zool. 101: 699-724. Steiner, W. A. 1994c. The influence of air pollution on mossdwelling animals: 4. Seasonal and long-term fluctuations of rotifer, nematode and tardigrade populations. Rev. Suisse Zool. 101: 1017-1031.

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Steiner, W. A. 1995. The influence of air pollution on moss dwelling animals: 5. Fumigation experiments with SO2 and exposure experiments. Rev Suisse Zool. 102: 13-40. Swedish Museum of Natural History. 2009. Microfauna on Antarctic nunataks. Accessed on 14 February 2010 at . Utsugi, K. and Ohyama, Y. 1989. Antarctic Tardigrada. Proc. NIPR Symp. Polar Biol. 2: 190-197. Utsugi, K. and Ohyama, Y. 1991. Tardigrades in King George Island (Antarctica). Zool. Sci. 8: 1198. Yang, T. 2002. The tardigrades from some mosses of Lijiang County in Yunnan Province (Heterotardigrada: Echiniscidae; Eutardigrada: Parachela: Macrobiotidae, Hypsibiidae). A. Zootaxon. Sin. 27: 53-64.

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Glime, J. M. 2017. Tardigrade Ecology. Chapt. 5-6. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. Ebook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 18 July 2020 and available at .

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CHAPTER 5-6 TARDIGRADE ECOLOGY TABLE OF CONTENTS Dispersal.............................................................................................................................................................. 5-6-2 Peninsula Effect ........................................................................................................................................... 5-6-3 Distribution ......................................................................................................................................................... 5-6-4 Common Species ................................................................................................................................................. 5-6-6 Communities ....................................................................................................................................................... 5-6-7 Unique Partnerships? .......................................................................................................................................... 5-6-8 Bryophyte Dangers – Fungal Parasites ............................................................................................................... 5-6-9 Role of Bryophytes in Fungal Interactions ................................................................................................ 5-6-12 Pollution ............................................................................................................................................................ 5-6-12 Acid Rain, SO2, and NO2 ........................................................................................................................... 5-6-13 Urban Environment .................................................................................................................................... 5-6-13 Tardigrades in Space ......................................................................................................................................... 5-6-14 Evolutionary Similarities to Bryophytes ........................................................................................................... 5-6-15 Sampling and Extraction ................................................................................................................................... 5-6-16 Checklist of Bryophyte Dwellers ...................................................................................................................... 5-6-17 Heterotardigrada (armored tardigrades) ..................................................................................................... 5-6-17 Eutardigrada (unarmored/naked tardigrades) ............................................................................................. 5-6-17 Summary ........................................................................................................................................................... 5-6-19 Acknowledgments ............................................................................................................................................. 5-6-19 Literature Cited ................................................................................................................................................. 5-6-19

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Chapter 5-6: Tardigrade Ecology

CHAPTER 5-6 TARDIGRADE ECOLOGY

Figure 1. Echiniscus, the genus with the most species among mosses. with permission.

Photo by Łukasz Kaczmarek and Łukasz Michalczyk,

Dispersal As already discussed, one suggested reason for patchy distribution of tardigrades is the difficulty of dispersal for this small organism. Miller et al. (1994) concluded that tardigrade distribution in the Antarctic is influenced more strongly by dispersal limitations than it is by climate or habitat interactions. McInnes and Convey (2005) found a low species richness of tardigrades (6 taxa) in the South Sandwich Islands in the sub-Antarctic. They found indications that the tardigrades originated from both subAntarctic and maritime Antarctic populations. Wind dispersal is considered the major means by which tardigrades move to new locations (Christenberry & Higgins 1979; Bertolani et al. 2009). The anhydrobiotic state is very light weight and can easily survive the various dangers of space (see below). Faurby et al. (2008) suggested dispersal rate may be coupled with survival in the anhydrobiotic state. Based on these assumptions, Bromley (2009) has considered the possibilities that habitats such as rooftops with mosses serve as islands for tardigrades. Sudzuki (1972) experimented with wind dispersal to moss mats and found that smaller micro-organisms (. West, C. C. 1984. Micro-arthropod and plant species associations in two subAntarctic terrestrial communities. Oikos 42: 66-73. Wiggins, G. B., Mackay, R. J., and Smith, I. M. 1980. Evolutionary and ecological strategies of animals in annual temporary pools. Arch. Hydrobiol. Suppl. 58: 97-206. Wikipedia: Chilopoda 2010. Updated 5 December 2010. Accessed 13 December 2010 at . Wikipedia: Diplopoda. 2010. Updated 2 December 2010. Accessed on 13 December 2010 at . Wikipedia: Myriapoda. 2010. Updated 2 December 2010. Accessed 13 December 2010 at . Wikipedia: Tachopodoiulus niger. 2012. Updated 10 June 2012. Accessed 18 September 2012 at . Willmann, C. 1928. Die Oribatidenfauna nordwestdeutscher und einiger suddeutscher Moore. Abh. Naturwiss. Bremen 27: 143-176. Willmann, C. 1931a. Oribatiden aus dem Moosebruch. Arch. Hydrobiol. 23: 333-347. Willmann, C. 1931b. Moosmilben oder Oribatiden (Oribatei). In: Dahl, F., Gischer, V. G., and Jena. (eds.). Die Tierwelt Deutschlands 22: 77-200. Willmann, C. 1933. Acari aus dem Moosebruch. Zeit. Morphol. Ökol. Tiere 27: 373-383. Winchester, N., Behan-Pelletier, V., and Ring, R. A. 1999. Arboreal specificity, diversity and abundance of canopydwelling oribatid mites (Acari: Oribatida). Pedobiologia 43: 391-400.

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Witte, H. 1991. Indirect sperm transfer in prostigmatic mites from a phylogenetic viewpoint. In: The Acari. Springer Netherlands, pp. 137-176. Wohltmann, A. 1998. Water vapour uptake and drought resistance in immobile instars of Parasitengona (Acari: Prostigmata). Can. J. Zool. 76: 1741-1754. Wohltmann, A. 1999. Life-history evolution in Parasitengonae (Acari: Prostigmata): Constraints on number and size of offspring. In: Ecology and Evolution of the Acari. Springer Netherlands, pp. 137-148. Wohltmann, A. 2004. No place for generalists? Parasitengona (Acari: Prostigmata) inhabiting amphibious biotopes. In: Weigmann, G., Alberti, G., Wohltmann, A., and Ragusa, S. (eds.). Acarine Biodiversity in the Natural and Human Sphere. Phytophaga 14: 185-200. Wohltmann, A. and Wendt, F. E. 1996. Observations on the biology of two hygrobiotic trombidioid mites (Acari: Prostigmata: Parasitengonae), with special regard to host recognition and tactics parasitism. Acarologia 37: 31-44. Womersley, H. 1961. Studies of the Acarina fauna of leaflitter and moss from Australia. 2. A new trachytid mite, Polyaspinus tuberculatus, from Queensland (Acarina, Trachytina). Rec. S. Austral. Mus. (Adelaide) 14(1): 115123. Woolley, T. A. 1968. North American Liacaridae, II – Liacarus (Acari: Cryptostigmata). J. Entomol. Soc. Kans. 41: 350-366. Yanoviak, S. P., Nadkarni, N. M., and Gering, J. 2003. Arthropods in epiphytes: A diversity component not effectively sampled by canopy fogging. Biodiv. Conserv. 12: 731-741. Yanoviak, S. P., Walker, H., and Nadkarni, N. M. 2004. Arthropod assemblages in vegetative vs humic portions of epiphyte mats in a neotropical cloud forest. Pedobiologia 48: 51-58. Yanoviak, S. P., Nadkarni, N. M., and Solano J., R. 2006. Arthropod assemblages in epiphyte mats of Costa Rican cloud forests. Biotropica 36: 202-210. Zhang, L., Paul, P., But, H., and Ma, P. 2002. Gemmae of the moss Octoblepharum albidum taken as food by spider mites. Porcupine 27 (Dec 2002): 15 accessed on 19 August 2005 at , 2 pp.

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Chapter 9-2: Arthropods: Mite Habitats, Minor Arachnids, and Myriapods

Glime, J. M. 2017. Arthropods: Crustacea – Copepoda and Cladocera. Chapt. 10-1. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. Ebook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 19 July 2020 and available at .

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CHAPTER 10-1 ARTHROPODS: CRUSTACEA – COPEPODA AND CLADOCERA TABLE OF CONTENTS SUBPHYLUM CRUSTACEA ......................................................................................................................... 10-1-2 Reproduction .............................................................................................................................................. 10-1-3 Dispersal .................................................................................................................................................... 10-1-3 Habitat Fragmentation................................................................................................................................ 10-1-3 Habitat Importance..................................................................................................................................... 10-1-3 Terrestrial ............................................................................................................................................ 10-1-3 Peatlands ............................................................................................................................................. 10-1-4 Springs ................................................................................................................................................ 10-1-4 Streams ............................................................................................................................................... 10-1-5 Collection Methods .................................................................................................................................... 10-1-5 CLASS BRANCHIOPODA, ORDER CLADOCERA ..................................................................................... 10-1-6 Adaptations ................................................................................................................................................ 10-1-6 Structural............................................................................................................................................. 10-1-6 Life Cycle Strategies ........................................................................................................................... 10-1-6 Habitats ...................................................................................................................................................... 10-1-6 Terrestrial ............................................................................................................................................ 10-1-6 Peat Bogs ............................................................................................................................................ 10-1-7 Aquatic ................................................................................................................................................ 10-1-9 Lakes............................................................................................................................................ 10-1-9 Streams ...................................................................................................................................... 10-1-10 CLASS MAXILLOPODA, SUBCLASS COPEPODA .................................................................................. 10-1-10 Adaptations .............................................................................................................................................. 10-1-11 Structure ............................................................................................................................................ 10-1-11 Life Cycle Strategies ......................................................................................................................... 10-1-11 Feeding ............................................................................................................................................. 10-1-12 Habitats .................................................................................................................................................... 10-1-12 Terrestrial .......................................................................................................................................... 10-1-12 Antarctic .................................................................................................................................... 10-1-14 Peat Bogs and Sphagnum .................................................................................................................. 10-1-14 Aquatic .............................................................................................................................................. 10-1-17 Mossy Tarns .............................................................................................................................. 10-1-17 Springs ....................................................................................................................................... 10-1-17 Rivulets ...................................................................................................................................... 10-1-18 Streams ...................................................................................................................................... 10-1-18 Splash Zones .............................................................................................................................. 10-1-19 Cave Pool ................................................................................................................................... 10-1-19 Summary ......................................................................................................................................................... 10-1-20 Acknowledgments ........................................................................................................................................... 10-1-20 Literature Cited ............................................................................................................................................... 10-1-20

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Chapter 10-1: Arthropods: Crustacea – Copepoda and Cladocera

CHAPTER 10-1 ARTHROPODS: CRUSTACEA – COPEPODA AND CLADOCERA

Figure 1. Simocephalus sp. with eggs in the carapace. Note the white Vorticella on the lower left edge of the carapace and near the base of the antennae. Photo by Jasper Nance through Creative Commons.

SUBPHYLUM CRUSTACEA Crustaceans (Figure 1) are those tiny arthropods that most of us have never noticed on the bryophytes. But in some habitats, and some parts of the world, the bryophytes – invaders of land – are home for such terrestrialized arthropods. This large subphylum is mostly marine or aquatic, including such familiar animals as barnacles, crabs, crayfish, krill, lobsters, and shrimp (Wikipedia: Crustacean 2011). But it is mostly the smaller animals, the microcrustacea, that inhabit the bryophytes. The Crustacea are distinguished from other arthropods by their two-parted limbs (biramous; e.g. the pincers on the end of a crab claw or divided antenna of Daphnia or Simocephalus – Figure 13) and a life cycle that includes a nauplius larva stage (first larval stage of many crustaceans, having an unsegmented body and usually a single eye, Figure 2), although most have additional larval stages after that. Almost all of them have a chitinous exoskeleton.

Figure 2. Nauplius of copepod. Creative Commons.

Photo from Wikipedia

Chapter 10-1: Arthropods: Crustacea – Copepoda and Cladocera

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Reproduction

Habitat Importance

Most crustaceans have separate sexes, but some change sex and many are parthenogenetic, with females producing viable eggs that develop into new organisms in the absence of fertilization (Wikipedia: Crustacean 2011). Eggs are generally released into the water column, but some isopods form a brood pouch and carry their eggs and young around with them. Many copepods form egg sacs that hang from the body until the young hatch. Decapods typically carry their eggs attached to their swimmerets. The meiofauna [small metazoans that pass through 500µm or greater sieves, but are retained on 40 or 62 or 40 μm sieves (Dražina et al. 2011)] of springs typically have shorter life cycles, permitting such groups as cyclopoid copepods to have a rapid recruitment ability (Robertson 2002) and other copepods and ostracods to develop rapidly compared to insects, completing their development in only a few months (Dole-Olivier et al. 2000).

Krebs (2001) reminded us that habitat heterogeneity is related to the creation of more ecological niches. Bryophytes can create many niches, providing protected space for the small microcrustaceans. Srivastava et al. (2004) contend that moss-arthropod ecosystems form natural microcosms that are useful for testing such concepts as fragmentation, metacommunity theory, and connections between biodiversity and ecosystem processes. Their small size, short generation times, hierarchical spatial structure, and contained, definable systems provide advantages in conducting field experiments that are subject to natural conditions and interactions with neighboring communities. The authors argue that "natural microcosms are as versatile as artificial microcosms, but as complex and biologically realistic as other [larger] natural systems."

Dispersal

Acosta-Mercado et al. (2012) found strong support for the hypothesis that abiotic factors (especially water chemistry of the bryophytes and pH) are important determinants of terrestrial microcrustacean diversity. They added that water-holding capacity is correlated with the morphology and canopy structure of the bryophytes. Roughness of the bryophyte canopy in the Bahoruco Cloud Forest, Cachote, Dominican Republic, was important in determining differences in species composition. For amoebae, the lowest species richness was on Acroporium pungens (Figure 3), a species with low roughness and faunal density, whereas Thuidium urceolatum had the highest roughness index, highest faunal richness, and highest species density. But for the 26 microcrustacean morphospecies among 11 bryophyte species, there was no detectable canopy effect on faunal richness or density. The lowest density of 1 individual per 50 cm2 was on the cushions of Leucobryum (Figure 4) with a maximum of 6±3.37 on the same area of the thallose liverwort Monoclea (Figure 5), suggesting that openness of the community might play a role in diversity.

As with mites and other bryophyte dwellers, microcrustacea might be dispersed on a "magic carpet" – bryophyte fragments on which they are living. Sudzuki (1972) tested this hypothesis by exposing moss-soil samples to wind velocities of 2.9 m s-1. Sampling at distances of 100-400 cm from the "wind" source, they determined that even after 2 months, wind velocities up to 2 m s-1 failed to disperse the Crustacea. Those animals dispersed were primarily protozoa. Nevertheless, encysted animals could get dispersed with bryophyte fragments or even with moss clumps that get carried by small mammals or wind.

Habitat Fragmentation Microarthropods must move from one leaf patch to another, or from hiding places to food sources. During this time, especially if disturbed during the daytime, they are vulnerable to desiccation. Gonzalez et al. (1998) experimented with such fragmented microcosms to determine parameters that led to success of the inhabitants. They found that when microecosystems were fragmented, species declines occurred. But when the patches were connected by habitat corridors, much as has been shown for large mammals, both abundance and distribution of the fauna experienced a rescue effect through immigration. Bryophytes can often serve as such corridors, providing places to replenish lost moisture and to hide from predators. Gonzalez and Chaneton (2002) used bryophyte habitats for experimentation. They fragmented the bryophyte communities and found that this system likewise experienced loss of both faunal species richness and community biomass. Rare species were more likely to become extinct. Moss habitat corridors that connected fragments to a larger "mainland" of bryophytes permitted immigration and maintained microarthropod richness, abundance, and biomass in the fragments. While we tend to view corridors as continuous suitable habitats, such continuity is probably not necessary for the larger arthropods like isopods. They can use the bryophyte clumps as islands of safety between larger suitable habitats such as leaf litter.

Terrestrial

Figure 3. Acroporium pungens in the Neotropics, a species with low roughness and low faunal density. Photo by Michael Lüth, with permission.

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Chapter 10-1: Arthropods: Crustacea – Copepoda and Cladocera

Figure 4. Leucobryum glaucum cushion, a species with low faunal density. Photo by Janice Glime.

Figure 6. Bohemia bog with Sphagnum cuspidatum, S. denticulatum, and others, showing the varied habitats of hummocks, hollows, and small pools available to bog fauna. Photo by Jonathan Sleath, with permission.

Figure 5. Monoclea forsteri, a liverwort that harbors a relatively high microcrustacean diversity. Photo by Jan-Peter Frahm, with permission.

Peatlands Peatlands, for our purposes those habitats dominated by Sphagnum and not including other types of peatlands (Figure 6), provide a mix of moist and dry mosses and pools influenced by those mosses. The "terrestrial plankton" are often sensitive to high CO2 concentrations and low O2 tensions such as those found among rotting leaves and other areas with high rates of decomposition (Stout 1963). For these organisms with good tolerance for low pH (sometimes below 4.0), Sphagnum provides a suitable habitat. Krebs (2001) found that the center of the Sphagnum moss mat had a higher species diversity than the edges, perhaps due to additional niches (habitat heterogeneity) resulting from the plant-associated species dwelling there. On the other hand, the low pH created through cation exchange and organic acids produced by mosses in the genus Sphagnum (Figure 6) is detrimental to many organisms. Hillbricht-Ilkowska et al. (1998) examined the role of pH on Crustacea and other organisms by providing powdered lime to the system. Measurements after 1-4 years and 20-23 years indicated that both the water Ca and that of the sediment were permanently raised. This change coincided with a significantly increased rate of decomposition and an increase in species richness and diversity of crustaceans, among others. Overall diversity was doubled. The treatment eliminated peatmosses from encroaching on the lake but had no effect on those of the surrounding area.

To add to this image of Sphagnum (Figure 6) as an unfriendly substrate, Smirnov (1961) stated that few animals were specialized to gain their nutrition by consuming emersed Sphagnum. He cited only one species of flies whose larvae are known to feed directly on Sphagnum. On the other hand, in such Sphagnum lakes the bladderwort, an insectivorous plant, traps and digests Crustacea such as Daphnia (Cladocera) – a not so friendly place for many. But Sphagnum (Figure 6) may play a more positive role in the lives of these fauna. Sphagnum has long been known for its antibiotic properties; it was used as a wound dressing in WWI. Could it protect the crustaceans from fungal or bacterial attacks? Furthermore, for these invertebrates it may serve as a refugium – a place to escape predators (Kuczyńska-Kippen 2008), possibly due to its antifeedant properties as well as small hiding places. Springs Among the favored habitats of limnoterrestrial (living in wet films on land) Crustacea are mosses of springs, i.e. these Crustacea are crenophilous, where temperature and pH were important determinants of community composition in four Northern Apennine springs (Bottazzi et al. 2011). Mosses in these springs usually had harpacticoid copepods and ostracods representing the Crustacea. The moss inhabitants had a seasonality, whereas drift assemblages did not. Bottazzi et al. suggest that the mosses were important in increasing the species diversity in these springs. Springs are often a transitional habitat between aquatic and terrestrial systems. Even within the spring habitat, such a transition is typical, and moisture zones within the habitat can change as the seasons and weather change. Thus, the bryophytes of this habitat provide not only a refuge, but an avenue (more like a labyrinth) where macroinvertebrates can travel to escape the receding preferred moisture level.

Chapter 10-1: Arthropods: Crustacea – Copepoda and Cladocera

Crustacea are not usually seen among bryophytes, but in some areas they can be quite abundant. For example, Michaelis (1977) reported that at Pupu Springs in New Zealand, there were ten species of bryophytes. The fauna included Crustacea among the most abundant groups. Suren (1993) suggests that the abundance of crustaceans in the New Zealand bryofauna may be due to the absence of some of the bryophyte dwellers found elsewhere, i.e. some families of Trichoptera (caddisflies), Plecoptera (stoneflies), and Ephemeroptera (mayflies). Bottazzi et al. (2011) reported the ostracods and Harpacticoida (an order of copepods) among the three most abundant taxon groups among mosses in northern Apennine rheocrene springs (springs that become streams immediately upon emerging from the ground). Like Michaelis (1977) and Suren (1993), they suggested that favorable habitats, including mosses, accounted for the high diversity and the large numbers of these two crustacean groups. Bottazzi et al. (2011) concluded that emergent mosses were important in increasing species diversity of these springs (see also Barquín & Death 2009; Ilmonen & Paasivirta 2005). Bryophytes act as an ecotone between the aquatic and terrestrial habitat by creating a range of microhabitats that vary both horizontally and vertically (Lindegaard et al. 1975; Thorup & Lindegaard 1977), including the madicolous zone (having thin sheets of water flowing over rock surfaces). These provide a range of moisture conditions that permit the meiofauna to migrate to a more suitable location as moisture conditions change. While providing a refuge from rapid flow (Madaliński 1961; Elliot 1967; Gurtz & Wallace 1984; Suren 1992; Glime 1994), bryophytes provide a variety of food sizes in trapped particulate matter (Habdija et al. 2004). Linhart et al. (2002c) demonstrated a direct association between harpacticoid copepods, including nauplii, and trapped organic and mineral matter among the mosses. Lindegaard et al. (1975) found that in the Danish spring at Ravnkilde these vertical and horizontal differences among the bryophytes provided a source of diversity among the macroinvertebrates. They found that whereas the horizontal zonation sported different assemblages of species, the fauna of the neighboring stones had little influence on the moss fauna. More importantly, the flow rate and available detritus as a food source could account for the horizontal differences. Lindegaard et al. (1975) found that the numbers of individuals fluctuated throughout the year, corresponding with changes in the life cycle stages of the dominate species. Bryophyte habitation is also seasonal in Northern Apennine springs, with a maximum in the spring and minimum in winter, whereas seasonal habitation is nearly constant in non-bryophyte areas sampled by the traps Bottazzi et al. (2011). On the other hand, permanent meiofauna had its minimum in autumn; temporary meiofauna of the mosses peaked in spring, then decreased thereafter. Streams Bryophytes in streams create a rich source of invertebrate fauna, so much so that the aquatic moss Fontinalis antipyretica (Figure 7) was transplanted to streams in South Africa to increase the food source for

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trout (Richards 1947). The bryophytes are able to provide a refuge from fast-flowing water and to increase stream heterogeneity (Tada & Satake 1994; Wulfhorst 1994; Dražina et al. 2011).

Figure 7. Fontinalis antipyretica. Frahm, with permission.

Photo by Jan-Peter

Despite their seeming rarity among bryophytes, Amos (1999) included ostracods, cladocerans, copepods, and amphipods as "life in the torrent" in the UK – a description of the inhabitants of Fontinalis (Figure 7). His point was that "all was quiet" at the bottom of the moss clump despite the torrent occurring at the surface. Linhart et al. (2002a), in Europe, found that regulated channels had a much greater meiofauna, including Cladocera and Harpacticoida (copepods), when the channel was overgrown by aquatic bryophytes, in this case Fontinalis antipyretica (Figure 7). In a different stream, the meiofauna of mosses was an order of magnitude higher than that in the surrounding mineral substrate (Linhart et al. 2000), but the crustaceans were not a significant part of this fauna. Rather, the density of the Harpacticoida was the second most abundant group in the gravel, where the fine particulate matter was also highest compared to that among the mosses. They further determined that high flow rates approaching the mosses had a negative impact on the crustaceans [Cladocera, Ostracoda, and Cyclopoida (an order of copepods)], although the velocity seemed to have no effect on the Harpacticoida (Linhart et al. 2002b, c). They suggested that fine detritus trapped by the F. antipyretica provided food for the harpacticoid copepods. It is interesting that in their 2000 study Linhart et al. suggested that it is "questionable whether F. antipyretica can serve as a refuge from the current for stream meiobenthos," a seeming contradiction to their conclusions in a different stream. It appears that food is the primary factor in distribution of the microcrustacea, but that does not rule out the role of the mosses as a refuge when sufficient food is present.

Collection Methods Methods of collection can have a biasing effect on the relative numbers of taxa collected. Copepods and other Crustacea in aquatic habitats can be collected by squeezing mosses into a collection bottle or squeezing the mosses in place and collecting the crustaceans downstream from the mosses with a plankton net (Gerecke et al. 1998; Reid 2001; Stoch 2007). Copepods, ostracods, and

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Chapter 10-1: Arthropods: Crustacea – Copepoda and Cladocera

amphipods may all be extracted from forest litter by the Berlese funnel, but as the litter dries out many will perish before they can escape (Stout 1963). Heat extraction can present the same problem. Chapman (1960) was successful in extracting terrestrial ostracods alive by slowly drying out the leaf litter (but it would work for bryophytes as well) in a Berlese funnel, using a water-jacket at 40° C to avoid overheating, in which case the ostracods close their valves and stop moving. The end of the funnel led to water rather than alcohol.

haven't found them yet. There are indications that appendage reduction is a terrestrial adaptation in this group. After all, why waste energy to make appendages that are not useful. Frey (1980) describes the non-swimming chydorid Bryospilus (Figure 9) from wet cloud forests as lacking a compound eye, a change that still requires explanation. The genus resembles the limnoterrestrial genus Monospilus, possibly through convergence. They exhibit reduced setation on their antennae and trunk limb, perhaps facilitating their slow crawl among wet bryophytes as high as 3-5 m above the forest floor.

CLASS BRANCHIOPODA, ORDER CLADOCERA The class name of Branchiopoda literally means gill feet and refers to the pereiopods by which the aquatic species can swim. The order name Cladocera derives from the Ancient Greek κλάδος (kládos, "branch") and κέρας (kéras, "horn").

Adaptations Structural Cladocera are a predominately aquatic group of small individuals known as water fleas (no relationship to the insect group of fleas). They swim using their antennae, using a series of jerks similar to the hops of a flea. Some have adapted to terrestrial habitats with free water, such as bromeliad basins. Others are able to use the film of water from the capillary spaces and leaf surfaces of bryophytes. Not only are the antennae important for swimming, but they are also powerful chemical sensory organs (Ecomare 2014). They can use these not only to find food, but also to detect the presence of enemies. The body of a cladoceran is a valve-like carapace that covers an unsegmented thorax and abdomen. Adults have a single compound eye. Life Cycle Strategies

Figure 8. Daphnia pulex with three eggs shown here to the right of the digestive tract. Photo by Paul Hebert, through Wikimedia Commons.

Cladocerans spend most of their lives as a female population that reproduces multiple times asexually by cyclical parthenogenesis. When conditions become unfavorable, they produce male offspring and subsequently reproduce sexually, producing resting eggs that remain within the carapace (Daphnia; Figure 8). In this state, they can dry out and travel long distances on wind currents or as hitch hikers on other travelling animals or even moss fragments. In fact, some of these dormant eggs are known to remain viable for 70-80 years in Lake Superior sediments (Kerfoot & Weider 2004) and can even survive the digestive tracts of birds (Figuerola & Green 2002).

Habitats Cladocera are primarily aquatic and marine, but a few are adapted to terrestrial living, taking advantage of films of water, pools in bromeliads, and other surfaces where they have easy access to water when they are active.

Figure 9. Bryospilus repens, a chydorid cladoceran that lives mostly in wet moss.. Photo by Francisco D. R. Sousa , with permission.

Terrestrial Since Cladocera live primarily in fresh or marine water, living on land requires special adaptations for both water conservation and locomotion. It seems that few cladoceran species have accomplished this, or we simply

Existing 3-5 meters above the rainforest floor are Cladocera that crawl from place to place, unable to swim. Frey (1980) reported the cladoceran Bryospilus repens (Figure 9), a semiterrestrial species known from wet mosses in Puerto Rico, Venezuela, and New Zealand, and

Chapter 10-1: Arthropods: Crustacea – Copepoda and Cladocera

Bryospilus bifidus from New Zealand, both in the same subfamily of Chydoridae as Alona (Figure 10-Figure 11), a common cladoceran from springs. Resting eggs are often buried in deep masses of vegetation (Powers & Bliss 1983) where they are protected from water loss. Dispersal of fragments of mosses they inhabit can aid in dispersal of both eggs and adults to new sites. Frey suggested that the mossy habitat in the rainforest exhibited the same continuity through time as ancient lakes, thus being a likely site for even more endemic species. Van Damme et al. (2011) consider B. repens (Figure 9) to be a "well known" species that lives in wet moss. They consider its occasional presence in river samples to be the result of individuals that got washed into the river from these mossy homes. There may be more species of these tiny cladocerans hiding among bryophytes in terrestrial habitats. These organisms are typically studied by aquatic biologists who spend their time looking at plankton. Terrestrial bryophyte habitats are rarely studied with the aim of locating Cladocera. I have to wonder if somewhere there might be some Cladoceran species living in liverwort lobules.

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Figure 11. Alona cf affinis, a common species in bog lakes. Photo by Yuuji Tsukii, with permission.

Peat Bogs Living among Sphagnum (Figure 6) or in the bog pools requires a tolerance of low pH. Nevertheless, Sphagnum can increase the abundance of Cladocera by as much as tenfold in Swedish peatlands (Henrickson 1993). The heterogeneity of the Sphagnum habitat illustrated in Figure 6 provides shelter and refuge against predation while being a suitable foraging site. The bryophytes further contribute to this habitat through their production of antibiotics, organic acids, and cation exchange. Bog lakes can support a number of species of Cladocera. Minelli (2004) listed Alona quadrangularis (Figure 10), Alona affinis (Figure 11), Simocephalus exspinosus (Figure 12), S. vetulus (Figure 13), and Ceriodaphnia pulchella as being among the common species in bog lakes in Italy. Hingley (1993) reported Streblocerus serricaudatus (Figure 14) and Acantholeberis curvirostris (Figure 15) swimming in UK peat pools. Macan (1974) likewise reported the latter species in Sphagnum (Figure 6). Chydorus piger (Figure 16) is typical of bare substrates such as rock or sand, but including Sphagnum, and is known from acidic pools in peatlands in Europe (Duigan & Birks 2000).

Figure 10. Alona quadrangularis, a common species in bog lakes. Photo by Ralf Wagner , with permission.

Figure 12. Simocephalus exspinosus, a common species in bog lakes. Photo by Malcolm Storey through , through online license.

Figure 13. Simocephalus vetulus, a common species in bog lakes. Note the divided (biramous) antenna (arrow). Photo by Ralf Wagner , with permission.

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Chapter 10-1: Arthropods: Crustacea – Copepoda and Cladocera

Kairesalo et al. (1992) considers the peatland habitat to be unsuitable for Daphnia (Figure 17) because the available food is "recalcitrant." In a lake in southern Finland that was bordered by the mosses Warnstorfia (Figure 18) and Sphagnum (Figure 6), the organic carbon excreted by Warnstorfia suppressed the growth of planktonic algae and provided little contribution to bacterial productivity. This meant that bacterial productivity was necessarily dependent on humic acids for their carbon source, resulting in decreased availability of this food source for the Daphnia. The predominantly particulate matter in the water was largely useless for the Daphnia as a food source.

Figure 14. Streblocerus serricaudatus, a cladoceran that inhabits peatland pools. Photo from Haney, J. F. et al. 2013. AnImage-based Key to the Zooplankton of North America, version 5.0 released 2013. University of New Hampshire Center for Freshwater Biology. Accessed 21 March 2014 at , with permission.

Figure 17. Daphnia. Creative Commons.

Photo by Gerard Visser through

Figure 15. Acantholeberis curvirostris, a cladoceran of peatland pools. Photo from Haney, J. F. et al. 2013. An-Imagebased Key to the Zooplankton of North America, version 5.0 released 2013. University of New Hampshire Center for Freshwater Biology. Accessed 21 March 2014 at , with permission.

Figure 18. Warnstorfia exannulata, a peatland moss that seems to be "recalcitrant," unable to provide food for the Cladocera living there. Photo from Biopix through Creative Commons.

Figure 16. Chydorus piger, a cladoceran from peatland pools. Photo by Angie Opitz, through online permission.

Cladocera have played a role in reconstructing the history of some peatlands. Duigan and Birks (2000) report on Sphagnum (Figure 6) and other bryophytes from 9200 BP microfossils in western Norway with Alonella nana (Figure 19), Alonella excisa (Figure 20), and Alona rustica (Figure 21). Alona rustica is also known in peat bogs among mosses in Italy (Minelli 2004).

Chapter 10-1: Arthropods: Crustacea – Copepoda and Cladocera

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Aquatic Lakes Typically, cladocerans are rare among aquatic mosses, being adapted for planktonic life. However, in the subAntarctic lakes of South Georgia, the most common invertebrate was the cladoceran genus Alona (Figure 22), with 2544 individuals in a liter of water (Hansson et al. 1996). Several species in this genus were present, with the greatest numbers among mosses that extended into shallow lakes. In fact, the littoral mosses had the highest number of invertebrate species (20) and abundance (1539 individuals) of invertebrates in those lakes. With increasing UV levels reaching the shallow Antarctic lakes, mosses may provide refugia that protect these invertebrates from UV damage.

Figure 19. Alonella nana, a cladoceran from peat deposits in ~9200 BP. Photo from Great Lakes Research Laboratory, through public domain.

Figure 20. Alonella excisa, a cladoceran that occurs in peat deposits in ~9200 BP. Photo by Manuel Elias, ECOSUR, through Creative Commons.

Figure 22. Alona sp., a genus with a number of terrestrial bryophyte-dwelling species. Photo by Yuuji Tsukii, with permission.

Figure 21. Alona rustica, a cladoceran that lives among bryophytes on stream banks. Photo from Haney, J. F. et al. 2013. An-Image-based Key to the Zooplankton of North America, version 5.0 released 2013. University of New Hampshire Center for Freshwater Biology. Accessed 21 March 2014 at , with permission.

Van Damme et al. (2011) explain the absence of Alona karelica in littoral samples of European lakes by suggesting that it may actually be a terrestrial cladoceran that is normally associated with moss. This species has been reported twice from Sphagnum (Figure 6) in Europe (Flößner 2000; Kuczyńska-Kippen 2008) and its European distribution coincides with that of regions of high Sphagnum diversity (see Séneca & Söderström 2008; Van Damme et al. 2011). Another species of Alona, A. bromelicola, is from Nicaragua and lives in the basins of bromeliads (Van Damme et al. 2011). Yet another species, Alona rustica (Figure 21), is present in collections of bryophytes from stream banks in Italy (Margaritora et al. 2002), another transitional habitat. Such transitional habitats often have both higher diversity and density of organisms, a phenomenon known as the edge effect (Leopold 1933; Lay 1938; Good & Dambach 1943; Bider 1968; Wiens 1976). Kuczyńska-Kippen (2008) examined the role of Sphagnum (Figure 6) compared to open water for zooplankton in a lake in Poland. The highest species diversity values occurred in the peat mat (mean = 0.67 for crustaceans compared to 1.76 for rotifers), whereas the

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Chapter 10-1: Arthropods: Crustacea – Copepoda and Cladocera

lowest values occurred in open water (0.36 and 0.99 respectively). The cladocerans present in the transition zone between the peat mat and the open water seem to relate to the presence of both invertebrate and vertebrate predators, and competition between the large cladocerans and smaller rotifers. For the cladocerans, Sphagnum (Figure 23) can serve as a refugium to protect them from other invertebrate predators.

expect to be frequent in moss communities. The entire group comprises about 13,000 species with three of its ten orders being the most common (Harpacticoida, Cyclopoida, Calanoida) and containing the ones known from bryophytes (Wikipedia: Copepod 2014). Copepods have two pairs of antennae and a single red compound eye (in most). They are perhaps the fastest organisms alive, swimming in irregular spurts (Kiørboe et al. 2010). Some of the meiofauna taxa have switched to direct development (lacking the larval stage) and care of their young (Dahms & Qian 2004), traits that are absent in most copepods but that are beneficial in a terrestrial environment. The Harpacticoida (Figure 24) have a short pair of first antennae (Figure 25), often a somewhat wormlike body, and are mostly benthic (living on the bottom) (Wikipedia: Harpacticoida 2013). Nevertheless, Dumont and Maas (1988) consider the harpacticoid copepods to be widespread in wet habitats such as wet mosses. The harpacticoid copepods include crawlers, walkers, and burrowers (Dole-Olivier et al. 2000), pre-adapting the crawlers and walkers to mobility in the water film of bryophytes.

Figure 23. Sphagnum cuspidatum mat (foreground) and nearby hummock (upper left), habitats where one can find more Cladocera than in the open water (upper left). Photo by Michael Lüth, with permission.

Cammaerts and Mertens (1999) discovered Bryospilus repens (Figure 9) in the Palaeotropics (tropical areas of Africa, Asia, and Oceania, excluding Australia) of western Africa, where it occurred in vernal pools of forests. This dispels the notion that this genus is strictly a moss dweller. One problem in sorting out the Cladocera-bryophyte relationship is that species descriptions frequently fail to include the substrate, reporting only the general habitat, if even that. Streams Stream drift, a popular topic in the 60's and 70's, is generally a phenomenon we relate to the insects and other macroinvertebrates. But microcrustacea can be part of this as well. For moss-dwelling Cladocera, this is a means to get from one moss clump to another in an unfriendly moving environment. Peric et al. (2014) found that of 60 invertebrate taxa in a moss-rich karst system in Croatia, six were annelids and arthropods from the meiofauna, representing 35% of the total drift, but among the most abundant drift organisms were several species of Alona (26.7%) (Figure 22), a cladoceran known for being a mossdweller (Hansson et al. 1996; Van Damme et al. 2011). The drift was lowest in winter and highest in autumn and late spring to early summer.

Figure 24. Terrestrial Canthocamptidae male, a harpacticoid copepod. Photo by Walter Pfliegler, with permission.

CLASS MAXILLOPODA, SUBCLASS COPEPODA The name Copepoda comes from the Greek word koʊpɪpɒd, which literally means oar-feet (Wikipedia: Copepod 2014). Copepods are microcrustacea, mostly 0.52 mm (Encyclopaedia Britannica 2012), usually occurring as planktonic or benthic organisms and not ones we would

Figure 25. Canthocamptus, a harpacticoid copepod showing antennae. Photo by Yuuji Tsukii, with permission.

Chapter 10-1: Arthropods: Crustacea – Copepoda and Cladocera

The Cyclopoida (Figure 26) are mostly planktonic (live in water column and float or drift – can't swim against a current) (Wikipedia: Cyclopoida 2013). Their antennae are longer than those of Harpacticoida but shorter than those of Calanoida, reaching no farther than the thorax. They are capable of rapid movement.

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Adaptations Copepods, like several other crustacean groups, have evolved to a terrestrial life style, but still live where water is generally available (Stout 1963). Bryophytes provide such a habitat. Stout suggests that through evolutionary time both copepods and ostracods moved from streams to adjoining moss carpets and currently are able to live among Sphagnum (Figure 23) as well as forest litter (Harding 1953, 1955). Bryophyte-dwelling copepods are not very numerous, which probably explains, in part, the absence of descriptions of adaptations to the bryophytic habitat. Nevertheless, one might consider the adaptations to a terrestrial life style as exemplary of bryophytic adaptations. One such adaptation is the absence of hemoglobin (Green 1959). This is a stretch, because it appears that this pigment has evolved primarily in those species with a parasitic life style and a limited number of mud-dwelling taxa. Nevertheless, it suggests that oxygen is in adequate supply in the bryophytic habitat, so energy-requiring pigment development is not necessary. Structure

Figure 26. Cyclops vicinus, a cyclopoid copepod carrying egg sacs. Photo by Ralf Wagner , with permission.

The Calanoida (Figure 27) are also mostly planktonic species (Wikipedia: Calanoida 2013). Unlike the short antennae of the Harpacticoida, the first antennae of the Calanoida extend about half the length of the body or more.

Figure 27. Neocalanus cristatus, a calanoid copepod showing the long antennae. Photo by Seward Line , with online permission for educational use.

Copepods are known for their egg longevity, with some surviving as much as 322 years (Hairston et al. 1995).

The moss-dwelling nauplius (larval stage; Figure 28) of the copepod uses its antennae for swimming and possesses a single eye that can disappear in some species in later developmental stages. The copepod eye, in at least some species, senses the direction of light and permits the copepod, by moving its tail, to keep its back oriented toward the light (Land 1988). This behavior furthermore permits the copepod to distinguish its own species from other species by the movement patterns. Directed movement in response to light seems to be useful in minimizing exposure to UV light in tidal areas (Martin et al. 2000). These light avoidance behaviors are probably less useful among bryophytes.

Figure 28. Copepoda nauplius, the immature state. Photo by Graham Matthews , with permission.

Life Cycle Strategies Whether living in water that freezes, pools that dry up, or among mosses and other terrestrial habitats, life cycle strategies are important in enduring unfavorable seasons

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Chapter 10-1: Arthropods: Crustacea – Copepoda and Cladocera

(Santer 1998). Terrestrial habitats are all unstable. Evolution favors traits that help the copepods sense and prepare for these potentially deadly periods. These strategies include dormancy and migration as escape mechanisms, but also include synchronizing growth and reproduction with favorable periods. Dormancy is a common trait among copepods, particularly in higher and temperate latitudes (Dahms 1995; Williams-Howze 1997). It permits them to survive periods of desiccation and other unfavorable conditions. The timing of dormancy varies with the species and can occur in different forms in multiple life cycle stages, including desiccation-resistant resting eggs, arrested larval development, encystment of juveniles and adults (Deevey 1941; Dahms 1995), and arrested development of adults (Dahms 1995; Williams-Howze 1997). Dormancy saves energy during a time when living conditions are unfavorable. In addition to facilitating copepod survival during desiccation, dormancy helps copepods escape unfavorable temperatures, insufficient oxygen availability, limited food availability, and predation. Among these dormancy strategies, one potential adaptation is encystment. Canthocamptus staphylinoides (Figure 29) is a harpacticoid copepod that encysts (Deevey 1941). Some members of this genus are known from mosses in the aquatic environment and peat bogs, where encystment can permit them to survive not only desiccation but also unfavorable temperatures.

Figure 29. Canthocamptus staphylinoides. Photo from US Geological Survey, through public domain.

Diapause can be defined as a delay in development in response to regular and recurring periods of adverse environmental conditions. In its narrow sense, it is initiated and terminated by triggers such as photoperiod, temperature, chemical cues, population density, and physiological factors (Dahms 1995). Feeding Fryer (1957a, b) considered chance encounter to be a primary mechanism in finding food for the mostly planktonic copepods. Nevertheless, chemoreceptors help them to distinguish edible from inedible food particles and thus may help somewhat in locating food. The carnivorous diet appears to be the primitive condition, with the change to an algal diet facilitating adaptive radiation.

Habitats Reid (1986, 1987, 1999, 2011) has contributed considerably to our knowledge of bryophyte-dwelling

copepods. She reported them from such overlooked habitats as mosses (including Sphagnum – Figure 23) and liverworts, as well as from tree holes (Reid 1986). She described the new species Muscocyclops therasiae from Brazil, primarily from soils, but also from mosses. Reid (2001) considered the publications on the harpacticoids and small cyclopoids from mosses in humid climates to be so numerous that they were almost impossible to review. She found that such "aquatic" mosses as Sphagnum (Figure 23) and Hypnum (Figure 30) as well as those bryophytes from more humid habitats provide homes for their own unique communities of copepods. Stoch (2007) attributes the copepod abundance to the complex spatial structure and high availability of food resources among bryophytes. In their study on Fontinalis antipyretica (Figure 7) meiofauna in Central Europe, Vlčková et al. (2002) found that harpacticoid copepods were able to feed on organic matter in the size range of 30-100 µm trapped within the moss clumps.

Figure 30. Calliergonella lindbergii (=Hypnum lindbergii), a moss genus where copepods are known to live. Photo by JanPeter Frahm, with permission.

Terrestrial One would not expect a plankton organism like the copepods to occur on mosses on land, but a few have managed to venture into that habitat. Paul Davison (pers. comm. 9 November 2011) reported to me that harpacticoid copepods are well known from terrestrial mosses, but finding documentation of that has been challenging. Menzel (1921, 1925) reported both cyclopoid and harpacticoid copepods as moss dwellers. Bryophytes do not harbor a rich fauna, so they have not attracted much attention from the copepodologists. Nevertheless, those copepods that live among mosses can, at times, be important to ecosystem functioning. For example, the harpacticoid copepods are a first food source for the young salamanders living near and among the mosses (Paul Davison, pers. comm. 9 November 2011) (See Epiphytes below). Scattered reports of terrestrial bryophyte-dwelling copepods, especially harpacticoids, occur in the literature (e.g. Olofsson 1918; Lang 1931), including mosses

Chapter 10-1: Arthropods: Crustacea – Copepoda and Cladocera

(including Sphagnum – Figure 23) and liverworts as habitat. The genus Bryocamptus seems to be among the more common taxa in the Eastern Hemisphere. Bryocamptus pygmaeus and B. zschokkei (Figure 31) occur primarily among mosses in Central Europe (Illies 1952). Harding (1958) reported Bryocamptus stouti from mosses in New Zealand.

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Bryocyclops bogoriensis from the Fiji Islands among mosses and in tree holes. More recently, Watiroyram et al. (2012) listed ten additional wet moss dwellers in the genus Bryocyclops in Thailand, mostly near springs and waterfalls. Harding (1953) reported that Epactophanes (Figure 54) and Maraenobiotus live in damp mosses in Europe. Epactophanes muscicola (in UK) avoids mosses that are very wet. Michailova-Neikova (1973) found that of the nine harpacticoid copepods living among wet mosses near water bodies on a mountain in Bulgaria, eight also lived among leaf litter. In an apparently rare Western Hemisphere record of bryophyte dwellers, Rocha (1994) described Metacyclops oraemaris as a new species from moist moss in São Paulo, Brazil. In neighboring Suriname, Menzel (1916) found Parastenocaris staheli (see Figure 33) among mosses in the old leaf axils of the palm Livingstonia.

Figure 31. Bryocamptus zschokkei female, a moss dweller. Photo from US Geological Survey, through public domain.

Lewis (1984) reported twelve species of harpacticoid copepods from terrestrial mosses in forests and open areas in New Zealand. Lewis (1972a) found copepods in New Zealand among forest mosses that remained moist most of the year. These included Elaphoidella silvestris (see Figure 32), a copepod among damp mosses on the forest floor or nearby, but this species is limited to the damp conditions of higher altitude bush areas of North Island and dripping wet forests of the West Coast of South Island.

Figure 32. Elaphoidella bidens. Members of this genus live among damp mosses on the forest floor of New Zealand. Photo through Creative Commons.

Mrázek (1893) found the harpacticoid copepod Maraenobiotus vejdovski among mosses in Bohemia, and Harding (1953) reported them from woodland mosses in Scotland. These copepods are small and slender, permitting them to live an aquatic life in the water film among mosses (Harding 1953). Scourfield (1939) reported Bryocyclops and Muscocyclops as living among mosses in Wales. With a name like Bryocyclops muscicola, one expects to find a moss-dweller. Reid (1999) reported this species, originally described from Indonesia, from a plant nursery in Florida, USA, apparently introduced with some of the plants, perhaps mosses. This is the only species of Bryocyclops known from continental US, although Bryocyclops caroli is known from Puerto Rico. In the Eastern Hemisphere the genus seems to be more common than in the Western Hemisphere, or perhaps just better known. Menzel (1926) described the new species Bryocyclops anninae from moist mosses in Java and reported

Figure 33. Parastenocaris lacustris female, member of a genus with species that live among epiphytic mosses. Photo from US Geological Survey, through public domain.

North American records seem to be almost nonexistent. Nevertheless, Margaret (Maggie) Ray (pers. comm. 9 November 2011) told me that she found copepods in many of her bryophyte samples across North Carolina, USA. Paul Davison (pers. comm. 9 November 2011) likewise has often found them among bryophytic epiphytes in Alabama. Others have reported on them as a group (Camann 2011; Camann et al. 2011). Seepage Areas – Seepage areas, typically with bryophytes, seem like a logical place to look for limnoterrestrial copepods. Scourfield (1932) found Bryocyclops pygmaeus, a common species, and Speocyclops dimentiensis among mosses of seeps on rock outcrops at Tenby in Wales. In New Caledonia, Hamond (1987) found Fibulacamptus among wet mosses as well as other wet terrestrial substrata. Fiers and Ghenne (2000) suggested an interesting role for mosses in forests. They provide epigean highways, especially for the tiny (~0.5 mm long) species, that help to connect the various patches of leaf litter and moist soils while also serving as a temporary or permanent habitat. Epiphytes – It is interesting that one can see canopy food webs similar to those in the water, with bryophytes forming the habitat structure. In a (regrettably) rare North American study, Camann and coworkers (Camann 2011; Camann et al. 2011) report communities at 84 m above the forest floor in the redwood forest of California, USA. In these humus moss patches harpacticoid copepods dwell, encysting when conditions get dry. And further up the food web are Wandering Salamanders (Aneides vagrans; Figure 34), likewise bryophyte dwellers, that use the copepods as food. Most likely there are birds or other vertebrates that prey on the salamanders.

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Chapter 10-1: Arthropods: Crustacea – Copepoda and Cladocera

America; in Austria it is commonly associated with salt pools (Kipp et al. 2012). The most common species in high-altitude peat bogs of Europe is Acanthocyclops vernalis (Figure 42), reaching an altitude of 2800 m in the Alps.

Figure 34. Aneides vagrans, a salamander whose larvae feed on terrestrial copepods. Photo by John P. Clare, through Creative Commons..

Antarctic Pesta (1928) described the harpacticoid copepod Attheyella koenigi (Harpacticoida: Canthocamptidae; see Figure 35) from mosses in a stream on the island of South Georgia in the Antarctic. Also on the island of South Georgia, it is likewise the family Canthocamptidae that has the only known copepod species living among mosses at the edges of shallow lakes (Hansson et al. 1996). Although only three larval forms were found, the mosses were the only location where these copepods appeared in that study of Antarctic lakes. Also among these Antarctic dwellers is the harpacticoid copepod Marionobiotus jeanneli (family Thalestridae) living among wet mosses (Pugh et al. 2002).

Figure 36. Megacyclops viridis, a widespread species whose habitats include peatlands. Photo by R. M. Kipp et al. at USGS, with permission.

Figure 37. Macrocyclops albidus female with egg sacs. Photo by Ralf Wagner at , with permission.

Figure 35. Attheyella americana immature. This genus has several bryophyte-dwelling species. Photo by US Geological Survey, through public domain.

Peat Bogs and Sphagnum Bog lakes and pools in peat bogs are often rich in copepod species (Minelli 2004). In the Italian bog pools and lakes (and likely throughout most of Europe as well), the copepods are represented by the orders Cyclopoida and Harpacticoida. The most abundant species are typically widespread predators, including Megacyclops viridis (Figure 36), Macrocyclops albidus (Figure 37-Figure 38), and Diacyclops bicuspidatus (Figure 39), and algal or detritus feeders including Paracyclops fimbratus (see Figure 48), Eucyclops serrulatus (Figure 55), Thermocyclops dybowskii (see Figure 40), and Tropocyclops prasinus (Figure 41). Megacyclops viridis seems to have been introduced to the Great Lakes of North

Figure 38. Macrocyclops albidus nauplius. Photo by Ralf Wagner at , with permission.

Chapter 10-1: Arthropods: Crustacea – Copepoda and Cladocera

Figure 39. Diacyclops bicuspidatus with egg sacs, a widespread predator that can be found on Antarctic bryophytes. Photo from Haney, J. F. et al. 2013. An-Image-based Key to the Zooplankton of North America, version 5.0 released 2013. University of New Hampshire Center for Freshwater Biology. Accessed 21 March 2014 at , with permission.

Figure 40. Thermocyclops sp. with egg sacs. Photo through Creative Commons.

Figure 41. Tropocyclops prasinus with egg sacs. Photo from Haney, J. F. et al. 2013. An-Image-based Key to the Zooplankton of North America, version 5.0 released 2013. University of New Hampshire Center for Freshwater Biology. Accessed 21 March 2014 at , with permission.

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Figure 42. Acanthocyclops vernalis female with egg sacs. Photo from Haney, J. F. et al. 2013. An-Image-based Key to the Zooplankton of North America, version 5.0 released 2013. University of New Hampshire Center for Freshwater Biology. Accessed 21 March 2014 at , with permission.

Peat bogs, with a ground cover of Sphagnum species (Figure 43), provide the film of water needed by limnoterrestrial copepods. Diacyclops languidus and D. hypnicola (see Figure 44) are small species adapted to living in the water film on the mosses and characteristic of peat bogs in the Alps, Apennines, and central and northern Europe (Minelli 2004). Among European alpine Sphagnum and other moss cushions one can find Bryocamptus pygmaeus, Epactophanes richardi (Figure 54), and Phyllognathopus viguieri. Barclay (1969) found the latter species in New Zealand among mosses at the base of gravel piles in the winter when the mosses become quite soggy. A species of Bryocyclops is common in this same habitat.

Figure 43. Sphagnum blanket bog. Photo through Creative Commons.

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Chapter 10-1: Arthropods: Crustacea – Copepoda and Cladocera

Figure 44. Diacyclops sp., a genus of small copepods with some species adapted for living in the water film of bog mosses. Photo from USGS, through public domain.

Stoch (1998b) originally described the new species Moraria alpina and re-described M. radovnae (see Figure 45) from the Alps of Italy and Slovenia, where they occurred among mosses, in bogs, and in interstitial spaces in brooks. Additional European alpine species, for example Bryocamptus veidovskji, Elaphoidella gracilis, Moraria mrazeki, M. alpina, Maraenobiotus veidovskji, and Hypocamptus brehmi, live only in peat bogs and interstitial mountain habitats (Minelli 2004). In Britain, one can find Moraria arboricola among Sphagnum (Figure 43), as well as in leaf litter and tree hole pools (Fryer 1993). It seems none of these are strict tyrphobionts (living only in peat bogs and mires).

Figure 45. Moraria laurentica female, member of a genus including moss dwellers in the Antarctic South Georgia Island and known from mossy swamps and wet mosses on stream banks in the Great Lakes area, USA. Photo from US Geological Survey, through public domain.

Figure 47. Canthocamptus sp. on the alga Spirogyra. Photo by Gerard Visser through Creative Commons.

In peatlands, the mosses can have an indirect influence on the fauna due to the tracheophytes they support. The rare North American copepod Paracyclops canadensis (Figure 48) is common in the pool of water in the leaves of the pitcher plant (Sarracenia purpurea, Figure 49) (Hamilton et al. 2000). In Sphagnum (Figure 43) peatlands, the mosses are a necessary habitat element to support the growth of pitcher plants.

Figure 48. Paracyclops canadensis, an inhabitant of pitcher plants. Photo from US Geological Survey, through public domain.

Herbst (1959) reported Metacyclops paludicola and Ectocyclops herbsti (see Figure 46) from a Sphagnum bog in São Paulo, Brazil. Hingley (1993) reported Moraria sphagnicola (see Figure 45) and Canthocamptus weberi (see Figure 47) as associated with Sphagnum (Figure 43) in Europe. In addition to living in mossy tarns, Attheyella (Delachauxiella) brehmi and Attheyella (Chappuisiella) maorica (see Figure 35) occur among Sphagnum in New Zealand (Lewis 1972a).

Figure 46. Ectocyclops phaleratus with egg sacs, member of a genus in which some species occur in peat bogs. Photo from Haney et al. 2013, with permission

Figure 49. Sarracenia purpurea leaf amid Sphagnum where copepods can live in the pool formed within the leaf. Photo by Janice Glime.

Chapter 10-1: Arthropods: Crustacea – Copepoda and Cladocera

Aquatic Reid (2001) reported that squeezing aquatic mosses would reveal small copepods such as members of Acanthocyclops (Figure 50-Figure 51), Diacyclops (Figure 52), and other small cyclopoid genera (Gurney 1932; Scourfield 1932, 1939). Aquatic bryophytes can provide cyclopoid genera with safe sites from strong flow, hide them from predators, and trap particulate matter that serves as food.

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Mossy Tarns Tarns (Figure 53) are small mountain lakes. For the crustaceans, the mossy tarn habitat is similar in many ways to peatland pools, but it may differ in its pH and does not necessarily have Sphagnum (Figure 43) or may have different Sphagnum species. Several copepod species seem to prefer mossy tarns in New Zealand (Lewis 1972a). Among these are Attheyella (Delachauxiella) brehmi and Attheyella (Chappuisiella) maorica (species known to occur among Sphagnum; see Figure 35) and Attheyella (Delachauxiella) bennetti, genera known also from peatlands.

Figure 50. Acanthocyclops venustoides, genus of the small copepods that live among aquatic mosses. Photo by US Geological Survey, through public domain.

Figure 53. Tarn in Siskiyou Wilderness, CA, USA. Photo by Miguel Vieira, through Creative Commons.

Springs

Figure 51. Acanthocyclops robustus, member of a genus of small copepods that live among bryophytes. Photo from Haney, J. F. et al. 2013. An Image-Based Key to the Zooplankton of North America, version 5.0 released 2013. University of New Hampshire Center for Freshwater Biology. Accessed 21 March 2014 at , with permission.

Figure 52. Diacyclops navus, genus of the small copepods that live among aquatic mosses. Photo from US Geological Survey, through public domain.

Stoch (2007) found that mosses in springs in Italy were particularly good habitats for copepods, supporting large numbers. This may be due to their complex structure and highly available food sources. At the same time, the spring-dwelling species are often not true crenobionts (occurring only in springs and spring brooks) (Stoch 1998a), also occurring in other damp or aquatic habitats such as the littoral zone of lakes, moist mosses elsewhere, in groundwater, and in the epirithral region (upstream stream region suitable for trout) (Gerecke et al. 1998; Jersabek et al. 2001; Galassi et al. 2002; Stoch 1998a, 2003, 2006, 2007). Within the springs, species often segregate into microhabitats that supply their needs, including hygropetric rivulets, mosses, and patches of sediments with different characteristics (Stoch 2003; Fiasca et al. 2005). Bottazzi et al. (2011) reported crenophilous ("loving" springs and spring brooks) crustaceans from mosses in the Northern Apennine rheocrene springs (springs that flow to surface from underground), with pH and temperature best explaining their distribution and diversity pattern. In fact, the harpacticoid copepods and ostracods dominated the moss fauna, along with stoneflies and Chironomidae. The mosses were important contributors to the biodiversity. We know that the copepod genera Moraria (Figure 45) and Bryocamptus are associated with wet or submerged mosses in Europe, including springs (Harding 1953). In their Italian study, Bottazzi et al. (2008) used traps, tubes, and moss samples to determine the copepod fauna of rheocrene springs (those that exhibit flow immediately after emerging from the substrate). They found 63% of the copepod taxa in these springs were represented among the

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Chapter 10-1: Arthropods: Crustacea – Copepoda and Cladocera

mosses, including a species of Moraria, in this case, M. poppei. Some of the copepod taxa occurred only in the moss habitat (i.e., were not collected in traps). These were the harpacticoid copepods Bryocamptus tatrensis, B. alpestris (see Figure 31), Moraria vejdovski, M. vejdovski truncatus, M. poppei, Epactophanes richardi (Figure 54), Attheyella crassa (see Figure 35), and the cyclopoid Eucyclops serrulatus (Figure 55). Bryocamptus species were evenly recorded from both moss and trap samples.

Streams It appears that copepods are important bryophyte inhabitants in mountain streams of New Zealand. In unshaded areas of the streams, Suren (1992) found Canthocamptus howardorum, C. maoricus (see Figure 56), Attheyella stillicidarum, A. cf. brehmi (see Figure 35), Antarctobiotus elongatus, and A. cf. diversus, all in the Harpacticoida (Figure 57). In 1992, Suren suggested that the large numbers of Copepoda found in association with bryophytes there may relate to the high food value of abundant periphyton that grow on the surfaces and the ability of the bryophytes to serve as safe sites against fast water currents. But in 1993, he refined his assessment to suggest that the copepods are especially important on bryophytes that are covered with detritus rather than periphyton (Suren 1993).

Figure 54. Epactophanes richardi female, a harpacticoid copepod of rheocrene springs that seems to prefer mosses. Photo from US Geological Survey, through public domain.

Figure 56. Canthocamptus from moss; note nauplius in insert. Photo by Graham Matthews , with permission. Figure 55. Eucyclops serrulatus, a harpacticoid copepod that lives among mosses of rheocrene springs. Photo by Fausto at , with permission.

Bottazzi et al. (2011) also reported that the taxa most represented in the Northern Apennine rheocrene springs were the harpacticoid copepods: Bryocamptus zschokkei (Figure 31) (mean number of individuals per sample = 2 for traps, 14 for mosses) and B. pygmaeus (1 individual/sample for traps, 5 for mosses). Out of their total of 3,284 invertebrates collected, Ostracoda, harpacticoid Copepoda, and Diptera were the most abundant among the 54 taxa. Bottazzi and coworkers considered the mosses to be a favorable habitat that contributed to the high species diversity. Rivulets Rivulets, often as outflow from springs, often have mosses that serve as copepod habitats. Stoch (2003, 2007) reported copepods from mosses in hygropetric rivulets (having water forming a surface film on rocks). Genera such as Moraria (Figure 45), Epactophanes (Figure 54), Arcticocamptus, Nitocrella, Parastenocaris (see Figure 33), Speocyclops, and Diacyclops (Figure 52) occur among hygropetric rivulet mosses (Fiasca et al. 2005).

Figure 57. Harpacticoid copepod on leaf of Fontinalis antipyretica, demonstrating how tiny it is. Photo by Dan Spitale, with permission.

Leaf axils of bryophytes can be particularly protective against the current, but they also serve as collection sites for detritus. The differences in periphyton vs organic detritus may relate to location in sun vs shade. Cox (1988) found that bryophytes from an unshaded location had predominantly periphyton associated with them, whereas

Chapter 10-1: Arthropods: Crustacea – Copepoda and Cladocera

those from the shaded site had predominately fine amorphous detritus associated with them. This is reasonable, as more light would promote greater algal growth. But flow rates will play into this as well, and oxygen content will differ with both flow rate and periphyton vs detrital matter. Chironomidae (midges) are typically the dominant group on stream bryophytes [see, for example Williams (1989) in southern Ontario, Canada, and Nolte (1991) in Germany, who found that chironomids dominated on the submerged moss Hygroamblystegium tenax (Figure 58)]. In New Zealand alpine streams, Suren (1992) found that harpacticoid copepods and ostracods were among the most abundant groups of non-chironomids. Suren found that there was a "strong positive relationship" between copepod density and high water velocity, with densities among the bryophytes there reaching twice that of macroinvertebrates. At first, this seems like a contradiction because meiofauna are intolerant of high water velocity (Winner 1975) and avoid it by burrowing into the hyporheic zone (sediment). Suren (1992) pointed out that the copepods Bryocamptus vejdovskyi and B. zschokkei (Figure 31) in Minnesota, USA, can only be found in the hyporheos in fast-flowing streams. He suggests that the bryophytes provide a "biotic hyporheic zone." The studies by Suren (1992) in New Zealand are in sharp contrast to those of Cox (1988) who found that in streams in Tennessee, USA, it was rotifers that dominated the bryophytic "hyporheic zone" in the mosses Fontinalis novae-angliae (Figure 59) and Platyhypnidium riparioides (Figure 60).

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Figure 60. Platyhypnidium riparioides, a moss that supports a dominant rotifer fauna rather than a copepod fauna in the hyporheic zone in Tennessee, USA. Photo by Michael Lüth, with permission.

Splash Zones Stream edges and waterfall splash zones provide a suitable habitat for some limnoterrestrial copepods (Lewis 1972a. In New Zealand one can find such taxa as Attheyella stillicidarum (see Figure 35) among the mosses and liverworts, preferring either permanently dripping mossy banks or areas in the splash zones of streams, apparently requiring moving (fresh, not stagnant) water. Attheyella humidarum and Attheyella fluviatalis likewise prefer dripping mossy banks and damp "bush" moss. In addition to these Attheyella species, Lewis (1972b) also described six new species in the genus Antarctobiotus (A. ignobilis, A. diversus, A. elongatus, A. australis, A. exiguus, A. triplex) from damp mosses in New Zealand. Cave Pool

Figure 58. Hygroamblystegium tenax, a submerged moss dominated by Chironomidae (midges - Diptera) rather than copepods in Germany. Photo by Barry Stewart, with permission.

Figure 59. Fontinalis novae-angliae at edge of stream, a moss that supports dominant rotifer fauna, not copepod fauna, in the hyporheic zone in Tennessee, USA. Photo by Janice Glime.

Galas et al. (1996) examined the decomposition of litter in a cave pool in Poland. These pools included copepods, among other fauna. Respiration released more energy by activity of microorganisms on mosses (Polytrichum, Figure 61) than on the litter of Sorbus and Alnus in the pool. This higher rate among the bryophytes suggests that they may have provided a better food source of fine particulates and microorganisms for small organisms such as copepods than that associated with the submersed leaf litter.

Figure 61. Polytrichum commune in a geothermal spring, Yellowstone, WY, USA. Photo by Janice Glime.

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Chapter 10-1: Arthropods: Crustacea – Copepoda and Cladocera

Summary Microcrustacea are primarily aquatic and marine, but some, including Copepoda and Cladocera, have developed characteristics that permit them to live on land in such habitats as wet bryophytes. Moisture, water chemistry, pH, and roughness of the moss habitat can be important determinants of microcrustacean diversity. Adaptations to land may include separate sexes, ability to change sex, and parthenogenesis. Cyclopoid copepods have short life cycles that permits them to increase recruitment. They can sometimes disperse with their bryophyte substrate. Truly terrestrial Cladocera are few, with Bryospilus being best represented among this group. Springs seem to be a transitional habitat between aquatic and terrestrial systems, with bryophytes serving as a refuge vertically and horizontally as moisture levels change. In streams, bryophytes can serve as a safety net to catch drifting organisms. The bottom of the moss clump provides a safe haven from the torrential waters above while being a collection site for food. Food is often fine detritus trapped by the bryophytes In these aquatic and wet habitats, the bryophytes can contribute significantly to increasing the faunal diversity. Peatlands/Sphagnum bogs increase diversity by offering multiple niches both in the mosses and among the tracheophyte vegetation. Alona and Alonella are among the most common there; Alona is also the most common drift cladoceran in streams. Cladoceran adaptations can include appendage reduction, shorter life cycle, eggs placed in dense masses of vegetation, and ability to swim in a thin film of water. Copepods on land use their antennae to swim in the larval stage. Dormancy permits them to survive dry periods, including resting eggs, arrested development, and encystment of both juveniles and adults. The ability of land-dwelling copepods to live among bryophytes is reflected in such names as Muscocyclops, Bryocyclops, and Epactophanes muscicola. Bryophytes can provide moist islands when copepods move from one location to another. Other species live among canopy epiphytes. Some even live among bryophytes in the Antarctic. Attheyella and Moraria are among the genera known from peat bogs, with genera such as Paracyclops found in pitcher plants there. Small copepods hide among the aquatic bryophytes. Harpacticoid copepods can dominate the moss fauna in springs, where temperature and pH are important factors in diversity. Canthocamptus and Attheyella are well represented in streams in New Zealand. Like the Cladocera, copepods often feed on periphyton or detritus among the bryophytes.

Acknowledgments I especially appreciate Dan Spitale for his contribution of the image of a copepod on Fontinalis and to Paul Davison for his anecdotal information, images, and

continued encouragement. Thank you to Larry Williams for numerous comments and suggestions that have improved the clarity of the manuscript. Thank you to all the photographers who have placed their images in Creative Commons on the internet.

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Glime, J. M. 2017. Arthropods: Crustacea – Ostracoda and Amphipoda. Chapt. 10-2. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. Ebook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 19 July 2020 and available at .

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CHAPTER 10-2 ARTHROPODS: CRUSTACEA – OSTRACODA AND AMPHPODA TABLE OF CONTENTS CLASS OSTRACODA ..................................................................................................................................... 10-2-2 Adaptations ................................................................................................................................................ 10-2-3 Swimming to Crawling ....................................................................................................................... 10-2-3 Reproduction....................................................................................................................................... 10-2-3 Habitats ...................................................................................................................................................... 10-2-3 Terrestrial ............................................................................................................................................ 10-2-3 Peat Bogs ............................................................................................................................................ 10-2-4 Aquatic ................................................................................................................................................ 10-2-8 Streams ........................................................................................................................................ 10-2-8 Springs ......................................................................................................................................... 10-2-8 CLASS MALACOSTRACA, ORDER AMPHIPODA .................................................................................... 10-2-8 Adaptations to Land – and Bryophytes ...................................................................................................... 10-2-9 Reproduction and Early Development .............................................................................................. 10-2-10 Food among the Bryophytes .................................................................................................................... 10-2-10 Habitats .................................................................................................................................................... 10-2-13 Terrestrial .......................................................................................................................................... 10-2-13 Aquatic .............................................................................................................................................. 10-2-14 Summary ......................................................................................................................................................... 10-2-15 Acknowledgments ........................................................................................................................................... 10-2-15 Literature Cited ............................................................................................................................................... 10-2-15

Chapter 10-2: Arthropods: Crustacea – Ostracoda and Amphipoda

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CHAPTER 10-2 ARTHROPODS: CRUSTACEA – OSTRACODA AND AMPHPODA

Figure 1. Terrestrial amphipod on leafy liverworts from New Zealand. Photo by Paddy Ryan, with permission.

The amphipods (Figure 1) and ostracods (Figure 2) might be considered as mimics that live in the bryophyte world. The amphipods look like miniature shrimp and the ostracods look like miniature mussel shells with a shrimp inside instead of a mussel.

become terrestrialized. They are not common among bryophytes, but they do sometimes occur there.

CLASS OSTRACODA Mark Papp (pers. comm. 19 November 2011) reported to me that he had a very sore neck and shoulders, but no ostracods to report. He had been looking at roof mosses where he had originally taken many ostracods at Chalfont St. Peter, UK. Their identity as ostracods was confirmed by a marine ecologist. He did find the remains of a copepod. The ostracods are evasive, making it that much more delightful when you find them. Those on the roof had apparently moved on. The name Ostracoda comes from the Greek óstrakon, meaning shell. Ostracods (sometimes known as seed shrimp) look like miniature clams (or seeds) with a tiny shrimp-like animal living inside the shell. They typically are marine and freshwater organisms, but some have

Figure 2. Ostracod, showing internal digestive system through the shell. Photo by Anna Syme through Wikipedia Commons.

Chapter 10-2: Arthropods: Crustacea – Ostracoda and Amphipoda

Adaptations Harding (1953) claimed the first find of a terrestrial ostracod (Mesocypris terrestris) as a new species occurring among mosses at the source of a small stream on Mt. Elgon in Kenya. Another occurred among mosses in a waterfall. But this ostracod is too large and globular for close alliance to the aquatic environment, so Harding (1953) reasoned that it must be more truly terrestrial. This ostracod is blind, presumably surviving loss of eyes because eyes are of little use among the mosses, and their swimming setae are very reduced as well. Instead, the second pair of antennae is especially powerful and Harding suggested that it might aid in movement in the water film among the mosses, a movement typically accomplished on mosses and liverworts by crawling (Powers & Bliss 1983). Excretion seems to be poorly understood, but some form of nitrogenous waste is excreted through glands on the maxillae, antennae, or both (Barnes 1982). Their food includes diatoms, bacteria, and detritus (Miracle 2014), items found not only in aquatic habitats, but also among terrestrial bryophytes. Swimming to Crawling A loss of ability to swim seems to be the result of an evolutionary loss of setae on antennae and reduction of setae on antennules (Harding 1953; De Deckker 1983; Martens et al. 2004). Instead, the terrestrial ostracods use their antennae to move along solid surfaces, much as benthic ostracods move along the bottom surface (Harding 1953; De Deckker 1983). On a moss, the ostracod is surrounded by a film of water at the bottom of the carapace (shell). This water is trapped by numerous hairs, especially ventrally and laterally, to about mid-height. This mechanism seems to work only on moist substrates. When Austromesocypris australiensis (=Mesocypris australiensis) was placed on a dry Petri plate, it was unable to retain all of the water when it moved (De Deckker 1983). Whereas most ostracods lie on their sides when at rest, this moss-dweller remains upright. As members of this species dry, they migrate to wetter conditions, but when it is too dry they close their shells (compare Figure 3 to Figure 9) to curtail water loss.

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Reproduction About half the non-marine ostracod species belong to the family Cyprididae (Wikipedia 2014). Many of these occur in temporary water bodies, requiring a degree of terrestrialization, and have drought-resistant eggs, mixed sexual and parthenogenetic reproduction, preadapting them to terrestrialization, and to living among bryophytes (Powers & Bliss 1983). There seems to be a prevalence of asexual reproduction among terrestrial ostracods compared to their aquatic counterparts (Pinto et al. 2005a). Nevertheless, terrestrialization of some may include retention of the fertilized eggs, protecting them from desiccation. Observations by Chapman (1961) suggest that the developing embryos of the moss-dweller Scottia audax (=Mesocypris audax) may be retained within the shell of the mother until they become free-living juveniles.

Habitats Terrestrial Although most ostracods are marine or aquatic, some, such as Mesocypris spp., live in wet terrestrial habitats, including mosses (Introduction to the Ostracoda 2002). This genus seems to be widespread among bryophytes in the Eastern Hemisphere from the Russian Far East (I'm unable to confirm this record) to Australia (Martens et al. 2004). Terrestrial species also occur in South America (Pinto et al. 2005a, b). Although Harding (1953) claimed the first record of terrestrial ostracods in Africa with his finding of Mesocypris terrestris, this one was still in the wet habitats of a waterfall and source waters of a stream among mosses. De Deckker (1983) collected Austromesocypris australiensis from Cammoo Caves in Queensland, Australia, from wet moss. De Deckker points out that although most ostracods are aquatic or marine, several species are able to live among leaf litter and mosses that are able to provide a moist environment. Among these, the type specimen of Austromesocypris australiensis was found among mosses, and others were living among Sphagnum (Figure 4) on the side of a road near a small creek in New South Wales, Australia. In fact, these individuals were unable to swim freely even in free water.

Figure 4. Sphagnum cristatum from a soil bank in New Zealand. Photo by Janice Glime.

Figure 3. Dead ostracod with its shell open, revealing the exoskeleton. When taken out of water, this shell immediately closes. Photo by Paul Davison, with permission.

In Queensland, the terrestrial ostracod Scottia audax (also known from mosses in New Zealand; Chapman 1961) occurred along with Austromesocypris australiensis in mosses (De Deckker 1983). Scottia birigida (Figure 5)

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Chapter 10-2: Arthropods: Crustacea – Ostracoda and Amphipoda

occurs among mosses in Japan (Robin James Smith, pers. comm. 31 March 2014). In Tasmania, Mesocypris tasmaniensis likewise occurs among mosses as well as litter (De Deckker 1983). Røen (1956) named Bryocypris grandipes from Africa (GBIF 2013), but I have only its name to suggest it dwells among bryophytes. De Deckker stated that terrestrial ostracods are known only from Gondwanaland: Africa, Madagascar, Australia, and New Zealand, but they have since been found in Europe (Pieri et al. 2009; Mark Papp, pers. comm. 19 November 2011) and South America, where Caaporacandona iguassuensis occurs among moist Brazilian forest mosses (Pinto et al. 2005a). Although members of the Cyprididae occur in

ophthalmica is known as a widespread species from the karst region of Italy (Wagenleitner 1990). All three species occur at the margins of lakes in the reed belt among the vegetation and on the sediment surface (Kiss 2007). The mosses were only examined from one site. One should note that these three species are also among the three most common taxa in the study (Figure 10), which included all the likely habitats for ostracods in the study area.

North America, thus far terrestrial representatives seem to be undocumented. Nevertheless, Paul Davison (pers. comm. 31 May 2014) reports them from dripping cliffs (Figure 6) among algae and suspects they could inhabit bryophytes under similar conditions. Bryologists should watch for them!

Figure 7. Cypria ophthalmica, a moss-dweller in Italy. Photo from Bold Systems through Creative Commons.

Figure 8. Cyclocypris laevis, a moss-dweller in Italy. Photo from Bold Systems through Creative Commons. Figure 5. Scottia birigida, a moss dweller in Japan. Photo by Robin James Smith, with permission.

Figure 9. Cyclocypris ovum, a moss-dweller in Italy, with its shell closed. Photo by Bold Systems Creative Commons.

Peat Bogs

Figure 6. Ostracod from wet wall, a potential bryophyte dweller. Photo by Paul Davison, with permission.

Pieri et al. (2009), reporting on ostracods from Friuli Venezia Giulia, Italy, found three species distributed on mosses: Cypria ophthalmica (Figure 7), Cyclocypris laevis (Figure 8), Cyclocypris ovum (Figure 9). It is not clear what the habitat was for these mosses. Cypria

Peat bogs seem to be a rich site for ostracod species. Harding (1953, 1955) states that ostracods tend to occur in Sphagnum (Figure 11) as well as in forest litter. Bryophytes influence the species composition by creating a diversity of niches, from pools to dry hummock tops, and many microniches among the stems and leaves. Likewise, a gradation of pH can sometimes be found vertically and horizontally, providing more niche choices. Temperature differs between the surface and deeper portions of peat. Figure 10 shows the relationships of four environmental parameters with the five most common ostracod species in 200 sites in the sampling of surface, interstitial, and ground waters of Friuli Venezia Giulia, Italy (Pieri et al. 2009).

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Figure 10. Comparison of environmental parameters for the five most common species in Friuli Venezia Giulia, Italy that also occur in peat bogs. The boxes show 25-75% quartiles. The horizontal line is the median, and vertical bars (whiskers) show the maximum and minimum values. The numbers of analyzed samples appear in parentheses below the species names. Redrawn from Pieri et al. 2009.

At Friuli Venezia Giulia, Italy, the five most widespread and common species of ostracods also occurred in peatlands (Pieri et al. 2009). Pieri and coworkers reported 24 species in 16 genera from peat bogs (Table 1).

Table 1. Ostracod species among those at Friuli Venezia Giulia, Italy, that occurred in peat bogs. From Pieri et al. 2009.

Figure 11. Sphagnum capillifolium representing a genus that houses several species of terrestrial ostracods. Photo by Blanka Shaw, with permission.

Darwinula stevensoni Figure 12 Penthesilenula brasiliensis Microdarwinula zimmeri Figure 13 Pseudocandona lobipes Pseudocandona compressa Figure 14 Pseudocandona pratensis Figure 15 Pseudocandona cf. sucki Cryptocandona vavrai Candonopsis scourfieldi see Figure 16 Cypria ophthalmica Figure 7 Cyclocypris globosa Figure 17 Cyclocypris laevis Figure 18

Cyclocypris ovum Figure 9 Ilyocypris bradyi Figure 29 Ilyocypris inermis Figure 19 Notodromas persica Figure 20 Eucypris pigra Figure 21 Herpetocypris sp. Figure 22 Herpetocypris reptans Figure 22 Scottia pseudobrowniana Cypridopsis elongata Figure 23 Cypridopsis vidua Figure 24 Cavernocypris subterranea Metacypris cordata Figure 25

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Figure 16. Candonopsis kingsleii, a peat bog species in Italy. Photo from Bold Systems through Creative Commons.

Figure 12. Darwinula stevensoni, an ostracod from mosses in peatlands in Italy. William Dembrowski through Creative Commons.

Figure 13. Microdarwinula zimmeri, a peat moss ostracod. Photo by Robin J. Smith, with permission.

Figure 17. Cyclocypris globosa, a peat bog species in Italy. Those white ovals near its surface are attached protozoa. Photo from Bold Systems through Creative Commons.

Figure 14. Pseudocandona compressa, a peat bog species in Italy. Photo from Bold Systems through Creative Commons.

Figure 18. Cyclocypris laevis, a peat bog species in Italy. Photo from Bold Systems through Creative Commons.

Figure 15. Pseudocandona pratensis, a peat bog species in Italy. Photo from Bold Systems through Creative Commons.

Figure 19. Ilyocypris inermis, a peat bog species in Italy. Photo from Bold Systems through Creative Commons.

Chapter 10-2: Arthropods: Crustacea – Ostracoda and Amphipoda

Figure 20. Notodromus sp., a peat bog species in Italy. Photo from Bold Systems through Creative Commons.

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Figure 24. Cypridopsis vidua, a peat bog species in Italy. Photo from Bold Systems through Creative Commons.

Figure 21. Eucypris pigra, a peat bog species in Italy. Photo from Bold Systems through Creative Commons.

Figure 25. Metacypris cordata, a peat bog species in Italy. Photo from Bold Systems through Creative Commons.

Figure 22. Herpetocypris reptans, a genus with members living in peat bogs in Italy. Photo from Bold Systems through Creative Commons.

Figure 23. Cypridopsis elongata, a peat bog species in Italy. Photo from Bold Systems through Creative Commons.

It is interesting that some of these Italian bog-dwelling species are so widespread. For example, Penthesilenula brasiliensis is known on all the continents except Antarctica and North America (Pieri et al. 2009). Its wide range of habitats (rivers, streams, interstitial water, bromeliad basins, rain forest leaf litter, and bog mosses) may permit this widespread geographic distribution. Furthermore, three of the most common species in this part of Italy have a wide altitudinal distribution (Figure 26). Surely they occur among bryophytes in other European countries as well. Some species seem to be restricted to bogs, making them tyrphobionts. In their study of Friuli Venezia Giulia, Italy, Cavernocypris subterranea and Cryptocandona vavrai were apparently restricted to peat bogs at high altitudes (Pieri et al. 2009). Barclay (1968) reported the new species Penthesilenula sphagna (=Darwinula sphagna) from New Zealand, living above the water among Sphagnum (Figure 4). Similar relationships of ostracods to Sphagnum are known from eastern Africa (Menzel 1916). The importance of mosses in bogs can be indirect. In Sphagnum (Figure 11) peatlands, mosses are a necessary habitat element to support the growth of pitcher plants (Sarracenia purpurea; Figure 27). The leaves of these plants form pitchers of water that provide a suitable habitat

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for some ostracods in North America (Hamilton et al. 2000), including those in Florida, USA (Harvey & Miller 1996).

pH range can reach into basic values, creating conditions that favor different communities. Bottazzi et al. (2011) compared ostracods collected in traps with those collected from mosses in rheocrene springs (where aquifer water reaches the surface) of the Northern Apennines. Ostracods were among the most abundant taxa, particularly among the permanent meiofauna. Such common inhabitants of springs can be called crenophiles (literally, spring-lovers). Only Psychrodromus bertharrami was collected in both traps and mosses, with similar numbers (20 individuals per sample in traps, 17 for mosses). Ilyocypris bradyi (Figure 29) was only recovered from mosses. All other taxa (except one of questionable identity) were collected in traps. Fryer (1955) described Potamocypris thienemanni (see Figure 28) as new to Britain, inhabiting bryophytes, including Sphagnum (Figure 11), in a spring. This species was also known from three springs in Germany.

Figure 26. Comparison of altitudinal ranges of the five most common ostracods in Friuli Venezia Giulia, Italy, all five of which also occur in peat bogs. The boxes show the 25-75% quartiles. The horizontal line is the median, and the vertical bars (whiskers) show the maximum and minimum values. The numbers of analyzed samples appear in parentheses below the species names. Redrawn from Pieri et al. (2009).

Figure 28. Potamocypris pallida, moss-dweller on sandy and rocky bottoms of Macedonian mountain springs and streams.. Photo by Elissa Dey, Zooplankton Project. Accessed 13 May 2014 at .

Figure 27. Sarracenia purpurea in a Sphagnum bog. Photo from Wikimedia Creative Commons.

Aquatic Streams Potamocypris pallida (Figure 28) in Macedonia occurs in moss cushions on the sandy and rocky bottoms of mountain springs and brooks (Petrovski & Meisch 1995). In my own stream bryophyte collections in Appalachian Mountain, USA, streams, I rarely encountered ostracods and considered them to be accidental or temporary residents since they more commonly occur in quiet water. Springs Spring habitats have a number of features in common with peat bogs. They typically have a dominant bryophyte flora, and they can be dry during part of the year. But their

Figure 29. Ilyocypris bradyi, an ostracod that in the northern Apennine springs seems to be limited to living among mosses. Note the hairy carapace that is typical of terrestrial ostracods. Photo from Bold Systems through Creative Commons.

CLASS MALACOSTRACA, ORDER AMPHIPODA I have occasionally found amphipods in my collections of stream mosses, but they are more typically in quiet water

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of larger streams. Nevertheless, it appears that in some cases they can be an important part of the aquatic moss faunal community (Minckley 1963; Minckley & Cole 1963; Matonickin & Pavletic 1964; Willoughby & Sutcliffe 1976). Badcock (1949) found Gammarus (Figure 30) to be most numerous in mosses and other protected niches, reporting an estimated fifty in a single tuft of moss (Cheney 1895). They are even known from terrestrial mosses (Merrifield & Ingham 1998).

Figure 32. Niphargus aquilex, a moss dweller. Photo by Grabow-Universität Koblenz-Landau, permission pending.

Adaptations to Land – and Bryophytes

Figure 30. Gammarus pulex showing the massive numbers in shallow pools. Photo through Creative Commons.

In some systems, amphipods can be quite abundant among the bryophytes. Wulfhorst (1994) found this to be true in two acid streams in the Harz Mountains, Germany, where they far exceeded those in the interstitial spaces (Figure 31).

Figure 31. Abundance (number of individuals per liter) of the amphipods Gammarus pulex and Niphargus aquilex (Figure 32) among mosses and the interstitial spaces at 10 and 30 cm depth at six stations in two Harz Mountain streams. Bars indicate 95% confidence interval; n = 14 for mosses and 28-36 for interstitial spaces. Redrawn from Wulfhorst 1994.

Stout (1963) summarized three evolutionary pathways for terrestrial plankton. Among these, Hurley (1959) proposed that amphipods moved from the supralittoral (splash zone) fauna directly to the forest floor. Another suggestion is that fauna such as amphipods may have originated in freshwater streams, extended to the wet mossy banks and Sphagnum (Figure 11) bogs to the forest floor and ultimately to mineral soil. Stout considers the latter route to be the most convincing. Hurley (1959, 1968) reported that all the terrestrial species of amphipods are in the family Talitridae, occurring in damp habitats. To survive in these terrestrial habitats required several morphological and behavioral changes, not to mention the physiological changes needed. They needed to become air breathers, jump instead of swim (accomplished by reduced pleopods, i.e. swimmerets, – to stumps in some species), adapt their life cycle to the changes in the seasons (Hurley 1959), and excrete uric acid instead of ammonia (Dresel & Moyle 1950). But they can have more than 50% ammonia excretion (Hurley 1959), perhaps releasing their ammonia as a gas like the isopods (O'Donnell & Wright 1995). It appears that they may have evolved different solutions to some of these problems from those of some of the other crustacean groups. We can understand the small number of terrestrial amphipod species by comparing them to the isopods, where both aquatic and terrestrial species likewise exist. Terrestrial amphipods are less adapted to their terrestrial life than the isopods, being restricted to more narrow niches (Hurley 1968). The amphipods lack the isopod advantages of evaporative cooling at high temperatures and have exoskeletons with greater permeability, leading to greater risk of desiccation (Hurley 1959). Terrestrial isopods have lost their antennae, whereas in amphipods they are merely simplified. Both groups have modified their behavior to stay where it is cool and moist. In wet leaf litter, the amphipods may move upward, a behavior we should look for among mosses (Hurley 1968). It is interesting that in the Fiordland of New Zealand the high level of rainfall and saturated ground has driven the amphipods to living among mosses or under bark of trees rather than their usual habitat of leaf litter. Avoidance of leaf litter there seems to be especially true for Arcitalitrus sylvaticus (=Talitrus sylvaticus; Figure 33). Its relative

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Arcitalitrus dorrieni (Figure 34) also occurs with mosses on bark in Australia.

Figure 33. Arcitalitrus sylvaticus, an amphipod that avoids leaf litter and lives among mosses in the New Zealand Fiordland. Photo by Arthur Scott Macmillan through Creative Commons.

she is more vulnerable to predation, and on land to desiccation. The eggs are deposited in the external brood pouch when she molts, followed by deposition of the sperm by the male. Following fertilization, embryos are carried by the female, but hatchlings, resembling miniature adults, are on their own. The terrestrial Talitrus saltator (a sandhopper; Figure 35) lays its eggs four days after molting, compared to laying them immediately after copulation (Figure 36) in the aquatic Gammarus (Hurley 1959). The latter species can hold the spermatozoa in a brood pouch for up to four days. The aquatic male Gammarus carries the female under him for several days (Figure 36), whereas the terrestrial male of Talitrus saltator does not carry the female, a behavior difference that seems backwards until you realize he is jumping around on the sand and the female would get in the way. The 1-10 terrestrial eggs are much larger than the small and numerous aquatic eggs. The eggs of the terrestrial species furthermore remain in the brood pouch longer, affording them greater protection from desiccation.

Figure 35. Talitrus saltator, a sand hopper that holds its eggs four days after molting. Photo by Arnold Paul through Wikimedia Commons.

Figure 34. Arcitalitrus dorrieni on bark among mosses. Photo by Dluogs through Creative Commons.

Obtaining water, no problem for aquatic species, requires special behavioral techniques for the land dwellers. It is interesting that the water-obtaining behavior is similar to that of the terrestrial oniscid isopods. The terrestrial amphipods both gain and eliminate water by dabbing the uropod tips (tails) onto wet or dry substrata, respectively (Moore & Richardson 1992). The water is exchanged rapidly in or out of the central channel through the capillary spaces between the body parts. Beating pleopods (abdominal appendages also known as swimmerets) transfer water from the abdomen to the thorax in most terrestrial taxa. Water that pools beneath the tail is taken in by anal drinking. Reproduction and Early Development Among amphipods, the male is typically larger than the female and mounts her dorsally when she is ready to molt (Sutcliffe 1992). This behavior of having the male carry the female beneath him, known as mate guarding, helps to protect her during the crucial mating molt while

Figure 36. Gammarus pulex copulating, with the larger male on top. Photo by J. C. Schou, with permission.

A further means to conserve both energy and water is neoteny. Orchestia (Figure 37) reaches sexual maturity at an earlier growth stage and smaller size (Powers & Bliss 1983). This results in fewer offspring. They have a female bias, somewhat compensating for the smaller number of offspring, and females are larger than males, which is atypical for amphipods. Stephensen (1935) reported Orchestia floresiana from moss in Java, where it grows in waterfalls, rivulets, and fountains.

Chapter 10-2: Arthropods: Crustacea – Ostracoda and Amphipoda

Figure 37. Orchestia cavimana at Colwick Park, Notts, UK This terrestrial genus has females larger than males. Photo by Roger S. Key, with permission.

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Figure 39. Comparison of mean percentage food types ±SD. of Gammarus fossarum as it relates to size. Modified from Felten et al. 2008.

Food among the Bryophytes Felten et al. (2008) found that some aquatic amphipods, or at least Gammarus fossarum (Figure 38), eat mosses, and that the relative proportion in the diet increases as they grow from 2 mm to 4 mm, increasing only slightly after that (Figure 39). Conversely, the proportion of fine amorphous detritus steadily decreases as the amphipods grow. Felten et al. suggest that the younger (smaller) amphipods do not have mouth parts that are developed well enough to eat the larger food items like bryophytes. The proportion of bryophytes in the diet also depends on where they are living, with those living among bryophytes eating a greater proportion of bryophytes (Figure 40). It is interesting that those in the detrital pools have a greater proportion of minerals, suggesting that they are unable to sort out the nutritious items from the nonnutritious items that surround them. It was unclear if the detrital pool populations were actually nibbling on the bryophytes or just eating fragments that had collected where they were.

Figure 38. Gammarus fossarum, an aquatic amphipod that eats mosses when its mouth parts are developed well enough to do so. Photo from BioLib.cz through public domain.

Figure 40. Comparison of mean proportion (±SD) of bryophytes vs other food items eaten by Gammarus fossarum in three habitat types. Modified from Felten et al. 2008.

Gladyshev et al. (2000) examined the gut contents of Gammarus lacustris (Figure 41) and found that they ingested mostly seston, obtaining omega 3 fatty acids from bottom sediment particles. They also consumed cells of the green alga Botryococcus. This alga not only survived the digestive tract, but its photosynthetic activity increased. They considered this activity to contribute to the dispersal of the alga, causing blooms in the littoral zone. Could this also be true of bryophytes they consume?

Figure 41. Gammarus lacustris, an amphipod that consumes mostly seston. Photo by Bold Systems Creative Commons.

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Food particle size can determine which species are able to survive in a habitat. Graca et al. (1994) found that Gammarus pulex (Figure 30-Figure 36) occupies different zones in rivers from those of the isopod Asellus aquaticus (Figure 42). The researchers found that the selection of substrate by G. pulex was based on size, with larger individuals choosing larger-sized substratum particles; juveniles were mostly associated with plants, including mosses. The substrate choices were most likely food choices. For the aquatic amphipods, it is likely that the detritus collected by the mosses serves as a food source. It would be interesting to determine the role of food sources in the choices of terrestrial amphipods for particular bryophytes.

One hypothesis is that increased density of bryophytes would increase available organic detritus and thereby increase invertebrate abundance. However, Suren and Winterbourn (1992) found that reducing stem density of mosses had little effect on periphyton biomass, but that the detrital biomass was reduced on low-density artificial mosses. In any case, stem density had little effect on invertebrate abundance. Nevertheless, detrital and periphyton availability seemed to be the determining factor for invertebrate density. Gladyshev et al. (2012) examined the gut contents of gammarids as part of a food chain study including Fontinalis antipyretica (Figure 44). Following Kalachova et al. (2011), they used acetylenic acids, considered as biomarkers for Fontinalis antipyretica, to trace the food through the mosses, periphyton, Trichoptera, gammarids, and Siberian grayling (Gladyshev et al. 2012). Gladyshev et al. (2012) found small amounts of consumption of the mosses among both the Trichoptera (caddisflies) and the gammarid Eulimnogammarus (Philolimnogammarus) viridis. The latter species had the highest concentrations of acetylenic acids in the winter and the lowest in summer (Kalachova et al. 2011), suggesting a shift to mosses in winter. It is likely that both the caddisflies and gammarids ate the moss to gain the periphyton and detritus accumulated there. The moss and associated periphyton and detritus are especially important in winter when other food sources are scarce (Gladyshev et al. 2012).

Figure 42. Asellus aquaticus, an aquatic isopod shown here on leaf litter. Photo by Malcolm Storey through Discover Life.

Acosta and Prat (2011) partially supported the idea of mosses as food collectors for the amphipod Hyalella sp. (Figure 43) in the headwaters of a High Andes river. Those living among layers of travertine had 69.5% fine particulate organic matter (FPOM), but even the bryophyte-dwellers had 56.8% FPOM. Those from leaf litter, on the other hand, had 68% of their gut contents from coarse particulate organic matter, suggesting a high level of flexibility in the diet.

Figure 44. Fontinalis antipyretica var gracilis, home for the amphipod Eulimnogammarus (Philolimnogammarus) viridis. Photo by Des Callaghan, with permission.

Figure 43. Hyalella azteca, a common bryophyte dweller in streams and rivers. Photo by Barbara Albrecht at , with permission.

But Parker et al. (2007) found that even when the moss Fontinalis novae-angliae (Figure 52) was cleaned of particulate matter, the amphipods still ate significant quantities of it. Earlier studies by Minckley and Cole (1963) likewise indicated that amphipods ate mosses. On the other hand, Mulholland et al. (2000) found that the amphipod Gammarus minus (Figure 45) depended on fine benthic (bottom) organic matter, despite the presence of bryophytes. One feeding possibility in nature that might not be evident in laboratory studies is the role of fungi. Barlocher and Porter (1986) demonstrated that Gammarus tigrinus (Figure 46) was able to digest plant polysaccharides and release sugars from maple leaves. They also had the right enzymes to break down glycosidic linkages in small molecules, much as that done in microbial decomposition.

Chapter 10-2: Arthropods: Crustacea – Ostracoda and Amphipoda

Furthermore, fungal carbohydrases ingested with the food of the Gammarus remained active in the gut. The implication seems to be that Gammarus could benefit from fungi associated with bryophytes in the field. Similarly, Sarah Lloyd (pers. comm.) has documented that terrestrial amphipods eat slime molds that live on mosses (Figure 47).

Figure 45. Gammarus minus, an amphipod that seems to prefer fine benthic organic matter over bryophytes. Photo through Creative Commons.

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It appears that at least some bryophytes are not suitable food for Gammarus (Figure 30-Figure 36). Willoughby and Sutcliffe (1976) conducted feeding experiments on Gammarus pulex (Figure 30) from the River Dutton. They found that those provided with only the liverwort Nardia sp. (Figure 48) were unable to grow or survive.

Figure 48. Nardia scalaris, a leafy liverwort genus in which a European species failed to sustain Gammarus pulex as a food source. Photo from Europe by Michael Lüth, with permission.

Habitats Terrestrial

Figure 46. Gammarus tigrinus, an amphipod that can digest fungi from leaves. Photo by Hugh MacIsaac, with permission.

Terrestrial amphipods are rare, but bryophytes can provide the kind of moist habitat needed for them to survive. Merrifield and Ingham (1998) found amphipods in their Oregon Coast Range, USA, study of the fauna of Eurhynchium oreganum (Figure 49). In most months they were not evident, but in the December collection their numbers rose to 1 per gram of moss in 10 5-cm samples. The second "peak" was in April, with 0.6 per gram. Sarah Lloyd (pers. comm.) found what appears to be Keratroides, possibly K. vulgaris, among mosses in a wet eucalypt forest in northern Tasmania.

Figure 47. Amphipod, probably Keratroides, possibly K. vulgaris, eating a slime mold (probably Diderma sp. ) on moss. Photo by Sarah Lloyd, with permission.

Figure 49. Eurhynchium oreganum, a moss that is known to house amphipods in North America. Photo by Adolf Ceska, with permission.

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bryophytes for larger stones. Juvenile densities in the daytime correlated positively with smaller particles and negatively to larger particles, also correlating with the weight of bryophytes. However, at night the densities were unrelated to particle sizes or bryophyte weight. Parker et al. (2007) found that the amphipod Crangonyx gracilis (see Figure 51) was a common inhabitant of the brook moss Fontinalis novae-angliae (Figure 52), where it used the moss shelter as a food source. Badcock (1949) found that Gammarus (Figure 30Figure 36) species were more numerous in sheltered sites such as mosses. Minckley (1963) found Gammarus among the moss Fissidens sp. (Figure 53) in a Kentucky, USA, stream. It not only lived there, but ate the moss that served as its home (Minckley & Cole 1963).

Figure 50. Terrestrial amphipod, probably Keratroides, possibly K. vulgaris, on mosses in wet sclerophyll (eucalypt) forest at Birralee in Northern Tasmania, Australia. Photo courtesy of Sarah Lloyd.

Friend (1987) described the new species Orchestiella neambulans from litter that accumulated between mosses in Tasmania. The Antarctic seems to be the most likely place to find limnoterrestrial Crustacea among mosses, but the amphipods are poorly represented. Pugh et al. (2002) found only one (Makawe insularis) in their study, a species with a broad niche of wood, leaf litter, lichens, tussock grass, under stones, penguin nests, and...among mosses.

Figure 51. Crangonyx pseudogracilis, relative of C. gracilis that lives among Fontinalis novae-angliae and also eats it. Photo from Discover Life - Creative Commons.

Aquatic Rocky streams are often dominated by mosses and liverworts in extensive mats over the rocks. These provide a foothold that protects their inhabitants from being swept away. Macan and Worthington (1951) found that amphipods such as Gammarus (Figure 30-Figure 36) were more likely on mosses that were not so thick, whereas thicker mosses were dominated by Chironomidae. They found that fish food organisms increased in number when the streams had rooted plants or mosses. One problem faced by the inhabitants of tracheophytes is that the plants begin die-off in late summer and the amphipods must find a new substrate with sufficient periphyton and detritus to provide food. Gammarus is among the slow colonizers (Fontaine & Nigh 1983), so it might benefit from the stable year-round habitat of bryophytes as a source of shelter and detrital and periphytic food. Elliott (2005) found that Gammarus pulex had significant day-night differences in its habitat distribution. These were explained by dry weights of bryophytes, leaf material, organic detritus, distance from bank, water depth, water velocity, and particle size class. The bryophyte weight correlated positively with larger particle sizes and negatively with smaller particle sizes, perhaps explaining some of the choices by G. pulex for bryophytes. But this correlation may have been due to the preference of

Figure 52. Fontinalis novae-angliae, shelter for Crangonyx gracilis. Photo by Janice Glime.

In an unlikely place, the depths of Yellowstone Lake, associated with active geothermal vents, Fontinalis abounds (Lovalvo et al. 2010). Associated with this unusual inhabitant are, among other invertebrates, the amphipods Hyalella (Figure 43) and Gammarus (Figure 30-Figure 36, Figure 41, Figure 45).

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interesting images and identifying the organisms. Robin James Smith not only gave me permission to use his images, but provided me with another moss record of an ostracod in Japan. Paul Davison has provided constant support with his interest, his images, and in this subchapter providing a critical review.

Literature Cited

Figure 53. Fissidens fontanus, both a home and food for some species of Gammarus. Photo by Michael Lüth, with permission.

Summary Both Ostracoda and Amphipoda are primarily marine, with fewer species in freshwater and much fewer in terrestrial habitats. Nevertheless, ostracods are known from roof mosses, and the genus Mesocypris is a common terrestrial moss dweller, especially in the Southern Hemisphere. Their adaptations to living among bryophytes (and other terrestrial locations) include swimming instead of crawling, small size, loss of eyes in some, reduced setae (used for swimming), excretion of nitrogenous waste from maxillae, antennae, or possibly through the carapace (perhaps as gaseous ammonia), droughtresistant eggs, and parthenogenesis. Food often consists of detritus, algae, and bacteria, but some amphipods eat bryophytes as well. Bogs offer habitats where ostracods can migrate vertically or horizontally to find suitable conditions as the temperature and moisture change. Some are even true tyrphobionts. But there appear to be few, if any, records for amphipods. Some ostracods live among the pitcher plants in the bogs. Few ostracods are known from among mosses in streams, but several amphipods can be found there. However springs seem to be suitable habitats for several species in both groups. The microcrustacea may have advanced onto land through wet mosses of springs and stream banks. They are represented by few families, the Cypridae among the ostracods and the Talitridae among the amphipods. Terrestrial amphipods are not well known, and thus far their presence among Sphagnum does not seem to be documented.

Acknowledgments Sarah Lloyd and her colleagues have been of invaluable help in this chapter by providing me with

Acosta, R. and Prat, N. 2011. Trophic ecology of Hyalella sp. (Crustacea: Amphipoda) in a High Andes headwater river with travertine deposits. Internat. Rev. Hydrobiol. 96: 274285. Badcock, R. M. 1949. Studies in stream life in tributaries of the Welsh Dee. J. Anim. Ecol. 18: 193-208. Barclay, M. H. 1968. Additions to the freshwater ostracod fauna of New Zealand. N. Z. J. Mar. Freshwat. Res. 2: 67-80. Barnes, R. D. 1982. Invertebrate Zoology. Holt-Saunders International, Philadelphia, pp. 680-683. Bottazzi, E., Bruno, M. C., Pieri, V., Sabatino, A. Di, Silveri, L., Carolli, M., and Rossetti, G. 2011. Spatial and seasonal distribution of invertebrates in Northern Apennine rheocrene springs. J. Limnol. 70(Suppl. 1): 77-92. Chapman, M. A. 1961. The terrestrial ostracod of New Zealand, Mesocypris audax sp. nov. Crustaceana 2: 255261. Cheney, A. N. 1895. Food for fishes. In: Davis, B. H., Lyman, H. H., Weed, W. R., Babcock, C. H., and Thompson, E. (eds.). First Annual Report of the Commissioners of Fisheries, Game and Forests, pp. 99-117. Deckker, P. De. 1983. Terrestrial ostracods in Australia. In: Lowrey, J. K. (ed.). Papers from the Conference on the Biology and Evolution of Crustacea. Australian Museum Memoir 18: 87-100. Dresel, E. I. B. and Moyle, V. 1950. Nitrogenous excretion of amphipods and isopods. J. Exper. Biol. 27: 210-225. Elliott, J. M. 2005. Day-night changes in the spatial distribution and habitat preferences of freshwater shrimps, Gammarus pulex, in a stony stream. Freshwat. Biol. 50: 552-566. Felten, V., Tixier, G., Guerold, F., Crespin De Billy, V. De, and Dangles, O. 2008. Quantification of diet variability in a stream amphipod: Implications for ecosystem functioning. Fund. Appl. Limnol. 170: 303-313. Fontaine, T. D. and Nigh, D. G. 1983. Characteristics of epiphyte communities on natural and artificial submersed lotic plants: Substrate effects. Arch. Hydrobiol. 96: 293301. Friend, J. A. 1987. The terrestrial amphipods (Amphipoda: Talitridae) of Tasmania: Systematics and zoogeography. Records of the Australian Museum, Supplement 7: 1-85. Fryer, G. 1955. XVI. – Potamocypris thienemanni Klie, a littleknown spring-inhabiting ostracod new to Britain. Ann. Mag. Nat. Hist. 8: 121-124. GBIF. 2013. The Global Biodiversity Information Facility: GBIF Backbone Taxonomy. Bryocypris Røen, 1956. Last updated 1 July 2013. Accessed 14 May 2014 at . Gladyshev, M. I., Emelianova, A. Y., Kalachova, G. S., Zotina, T. A., Gaevsky, N. A., andf Zhilenkov, M. D. 2000. Gut content analysis of Gammarus lacustris from a Siberian lake using biochemical and biophysical methods. Hydrobiologia 431: 155-163. Gladyshev, M. I., Sushchik, N. N., Kalachova, G. S., and Makhutova, O. N. 2012. Stable isotope composition of fatty

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acids in organisms of different trophic levels in the Yenisei River. PLoS one 7(3), e34059. Graca, M. A. S., Maltby, L., and Calow, P. 1994. Comparative ecology of Gammarus pulex (L.) and Asellus aquaticus (L.) I: Population dynamics and microdistribution. Hydrobiologia 281: 155-162. Hamilton, R. IV, Reid, J. W., and Duffield, R. M. 2000. Rare copepod, Paracyclops canadensis (Willey), common in leaves of Sarracenia purpurea L. Northeast. Nat. 7: 17-24. Harding, J. P. 1953. The first known example of a terrestrial ostracod, Mesocypris terrestris sp. nov. Ann. Natal Mus. 12: 359-365. Harding, J. P. 1955. The evolution of terrestrial habits in an ostracod. Bull. VII. Symposium on Organic Evolution, National Institute of Sciences of India, New Delhi, pp. 104106. Harvey, E. and Miller, T. E. 1996. Variance in composition of inquiline communities in leaves of Sarracenia purpurea L. on multiple spatial scales. Oecologia 108: 562-566. Hurley, D. E. 1959. Notes on the ecology and environmental adaptations of the terrestrial Amphipoda. Pacific Sci. 13: 107-109. Hurley, D. E. 1968. Transition from water to land in amphipod crustaceans. Amer. Zool. 8: 327-353. Introduction to the Ostracoda. 2002. University of California Museum of Paleontology. Accessed 6 November 2011 at . Kalachova, G. S., Gladyshev, M. I., Sushchik, N. N., and Makhutova, O. N. 2011. Water moss as a food item of the zoobenthos in the Yenisei River. Central Eur. J. Biol. 6: 236-245. Kiss, A. 2007. Factors affecting spatial and temporal distribution of Ostracoda assemblages in different macrophyte habitats of a shallow lake (Lake Fehér, Hungary). Hydrobiologia 585: 89-98. Lovalvo, D., Clingenpeel, S. R., McGinnis, S., Macur, R. E., Varley, J. D., Inskeep, W. P., Glime, J., Nealson, K., and McDermott, T. R. 2010. A geothermal-linked biological oasis in Yellowstone Lake, Yellowstone National Park, Wyoming. Geobiology 8: 327-336. Macan, T. T. and Worthington, E. B. 1951. Life in Lakes and Rivers. Collins, London, 272 pp. Martens, K., Deckker, P. De, and Rossetti, G. 2004. On a new terrestrial genus and species of Scottiinae (Crustacea, ostracods) from Australia, with a discussion on the phylogeny and the zoogeography of the subfamily. Zool. Anz. 243: 21-36. Matonickin, I. and Pavletic, Z. 1964. Postanak i razvoj najmladin sedrenih tvorevina u rijeci Uni s bioloskog stanovista. Jugoslav. Akad. Znan. Umjetn. Krs. Jugoslavje 4: 103-112. Menzel, R. 1916. Moosbewohnende Harpacticiden und Ostracoden aus Ost-Afrika. Arch. Hydrobiol. 11: 486-489. Merrifield, K. and Ingham, R. E. 1998. Nematodes and other aquatic invertebrates in Eurhynchium oreganum (Sull.) Jaeg., from Mary's Peak, Oregon Coast Range. Bryologist 101: 505-511. Minckley, W. L. 1963. The ecology of a spring stream Doe Run, Meade Co., Kentucky. Wildlf. Monogr. 11: 1-126. Minckley, W. L. and Cole, G. A. 1963. Ecological and morphological studies on gammarid amphipods (Gammarus spp.) in spring-fed streams of northern Kentucky. Occ. Papers C. C. Adams Center Ecol. Studies., W. Mich. Univ., Kalamazoo, 35 pp.

Miracle. 2014. Ostracods. Accessed 31 March 2014 at . Moore, M. L. and Richardson, A. M. M. 1992. Water uptake and loss via the urosome in terrestrial talitrid amphipods (Crustacea: Amphipoda). J. Nat. Hist. 26: 67-77. Mulholland, P. J., Tank, J. L., Sanzone, D. M., Wollheim, W. M., Peterson, B. J., Webster, J. R., and Meyer, J. L. 2000. Food resources of stream macroinvertebrates determined by natural-abundance stable C and N isotopes and a 15N tracer addition. J. N. Amer. Benthol. Soc. 19: 145-157. O’Donnell, M. J. and Wright, J. C. 1995. Nitrogen excretion in terrestrial crustaceans. In: Walsh, P. J. and Wright, P. (eds.). Nitrogen Metabolism and Excretion. CRC Press, Boca Raton, FL, pp. 105-118. Parker, J. D., Burkeile, D. E., Collins, D. O., Kubanek, J., and Hay, M. E. 2007. Stream mosses as chemically-defended refugia for freshwater macroinvertebrates. Oikos 116: 302312. Petrovski, T. K. and Meisch, C. 1995. Interesting freshwater Ostracoda (Crustacea) from Macedonia. Bull. Soc. Nat. Luxemb. 96: 167-183. Pieri, V., Martens, K., Stoch, F., and Rossetti, G. 2009. Distribution and ecology of non-marine ostracods (Crustacea, Ostracoda) from Friuli Venezia Giulia (NE Italy). J. Limnol. 68: 1-15. Pinto, R. L., Rocha, C. E. F., and Martens, K. 2005a. On new terrestrial ostracods (Crustacea, ostracods) from Brazil, Primarily from São Paulo State. Zool. J. Linn. Soc. 145: 145-173. Pinto, R. L., Rocha, C. E. F., and Martens, K. 2005b. On the evolution of the genus Microdarwinula Danielopol, 1968 (Ostracoda, Darwinulidae) with the description of a new species from semi-terrestrial habitats in São Paulo State (Brazil). Crustaceana 78: 975-986. Powers, L. W. and Bliss, D. E. 1983. Terrestrial adaptations. In: Vernberg, F. J. and Vernberg, W. B. The Biology of Crustacea 8, Environmental Adaptations. Academic Press, London, pp. 271-333. Pugh, P. J. A., Dartnall, H. J. G., and McInnes, S. J. 2002. The non-marine Crustacea of Antarctica and the Islands of the Southern Ocean: Biodiversity and biogeography. J. Nat. Hist. 36: 1047-1103. Stephensen, K. 1935. Terrestrial Talitridae from the Marquesas. Bernice P. Bishop Museum Bull. 142: 19-34. Stout, J. D. 1963. The terrestrial plankton. Tuatara 11: 58-64. Suren, A. M. and Winterbourn, M. J. 1992. The influence of periphyton, detritus and shelter on invertebrate colonization of aquatic bryophytes. Freshwat. Biol. 17: 327-339. Sutcliffe, D. W. 1992. Reproduction in Gammarus (Crustacea, Amphipoda): Basic processes. Freshwat. Forum 2: 102-129. Wagenleitner, H. 1990. Morphology and evolution of Cypria cavernae n. sp. (Ostracoda, Crustacea). Bull. Soc. Nat. Luxemb. 90: 199-226. Wikipedia. 2014. Ostracod. Last updated 22 March 2014. Accessed 31 March 2014 at . Willoughby, L. G. and Sutcliffe, D. W. 1976. Experiments on feeding and growth of the amphipod Gammarus pulex (L.) related to its distribution in the River Duddon. Freshwat. Biol. 6: 577-586. Wulfhorst, J. 1994. Selected faunal elements of the hyporheos and in submerged moss clumps (bryorheal) along acidification gradient in two brooks in the Harz Mountains, West Germany. Verh. Internat. Verein. Limnol. 25: 15751584.

Glime, J. M. 2017. Arthropods: Crustacea – Isopoda, Mysida, and Decapoda. Chapt. 10-3. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. Ebook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 19 July 2020 and available at .

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CHAPTER 10-3 ARTHROPODS: CRUSTACEA – ISOPODA, MYSIDA, AND DECAPODA TABLE OF CONTENTS CLASS MALACOSTRACA, ORDER ISOPODA ........................................................................................... 10-3-2 External Anatomy ...................................................................................................................................... 10-3-3 Adaptations to Terrestrial Life and to Bryophytes ..................................................................................... 10-3-4 Water Relations................................................................................................................................... 10-3-4 Waste Elimination............................................................................................................................... 10-3-5 Osmotic Balance ................................................................................................................................. 10-3-5 Respiration .......................................................................................................................................... 10-3-5 Temperature Tolerance ....................................................................................................................... 10-3-5 Moisture and Temperature Interaction ................................................................................................ 10-3-6 Behavior .............................................................................................................................................. 10-3-6 Congregating Behavior ................................................................................................................ 10-3-6 Sheltering ..................................................................................................................................... 10-3-7 Reproduction....................................................................................................................................... 10-3-7 Predators ............................................................................................................................................. 10-3-9 Overwintering ..................................................................................................................................... 10-3-9 Bryophytes as Food.................................................................................................................................. 10-3-10 Digestion ........................................................................................................................................... 10-3-10 Terrestrial Consumers ....................................................................................................................... 10-3-11 Defenses and Apparency Theory ............................................................................................... 10-3-18 Aquatic Consumers ........................................................................................................................... 10-3-19 Apparency or UV Protection?.................................................................................................... 10-3-21 Habitat ...................................................................................................................................................... 10-3-21 Terrestrial .......................................................................................................................................... 10-3-21 Peatlands ........................................................................................................................................... 10-3-25 Springs .............................................................................................................................................. 10-3-26 Waterfalls .......................................................................................................................................... 10-3-26 Aquatic .............................................................................................................................................. 10-3-26 Pollution..................................................................................................................................... 10-3-27 CLASS MALACOSTRACA, ORDER MYSIDA .......................................................................................... 10-3-27 CLASS MALACOSTRACA, ORDER DECAPODA .................................................................................... 10-3-27 Summary ......................................................................................................................................................... 10-3-29 Acknowledgments ........................................................................................................................................... 10-3-29 Literature Cited ............................................................................................................................................... 10-3-29

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Chapter 10-3: Arthropods: Crustacea – Isopoda, Mysida, and Decapoda

CHAPTER 10-3 ARTHROPODS: CRUSTACEA – ISOPODA, MYSIDA, AND DECAPODA

Figure 1. Porcellio scaber eating Pleurozium schreberi at midnight in Houghton, Michigan, USA. Photo by John Hribljan, with permission.

CLASS MALACOSTRACA, ORDER ISOPODA Then there are the Isopoda (Figure 1), the well-known pillbugs, woodlice, roly polies, potato bugs, or sowbugs (but they aren't bugs!). These aren't insects at all, but are arthropods with legs on each segment, sometimes included among the multipedes, which is an unofficial classification referring to arthropods with many legs. And at least some of them seem to love mosses. As a teacher, these were my favorite creatures. They have wonderful behavior responses to all sorts of things, especially light, moisture, and contact. Hence, they were excellent experimental organisms for behavior experiments for beginning students. They were easy to collect (just put out potatoes, with holes drilled through them, in a deciduous forest and give them 2-3 days to colonize). And they responded quickly and predictably. But for research on herbivory on bryophytes, these organisms are unparalleled. Both aquatic and terrestrial species eat mosses, are abundant, and can be used to test

for preferences. Nevertheless, they should not be considered as models for the feeding preferences of other invertebrates, as you will see when we discuss digestion. I have a small moss garden, and it is occasionally the site of my experiments, planned or otherwise! I had inherited a mat of mosses that had made themselves unwelcome on an asphalt parking lot. Some of these I had draped over a large rock in hopes that they would find it similar to their past home. In an attempt to keep them in place, I had used a mix of raw egg to act as glue. All seemed well for 2-3 weeks. Then one day when I went to look at them the mat looked like Swiss cheese! This carpet of a half-meter diameter had numerous relatively large holes in it! I found the carpet was loose, so I lifted it from the rock. As I did that, woodlice (mostly Porcellio scaber, Figure 2) fell to the ground and scrambled for cover. There were at least 20 of them! And many still remained on or within the mat.

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Figure 4. Trachelipus rathkii, an isopod that lives among epiphytic mosses in Hungary. Photo by Dragiša Savić, with permission. Figure 2. Porcellio scaber, a common moss inhabitant. Photo from , with permission.

Compared to other arthropods, the isopods, at least on land, probably have the most interaction with the bryophyte community. Božanić (2008) sampled 66 mosses and extracted their inhabitants using heat with a Tullgren apparatus. She recorded multiple factors to determine the niche requirements of the faunal species. The mosses represented 15 species. The Isopoda were the most abundant taxa (439 individuals); others included Chilopoda (centipedes), Diplopoda (millipedes), Araneae (spiders), Pseudoscorpionida (pseudoscorpions), Opilionida (daddy-long-legs), Lumbricidae (earthworms), and Formicidae (ants). The diplopods (another multipede) were second in abundance (240 individuals). The most important environmental factors in determining the faunal higher taxa were type of substrate, height above ground, and moss/sample area. The species factors, like those of the higher taxa, were substrate type and height above the ground, but in addition to these the tree diameter was important, possibly indicating colonization time. Farkas (2007) likewise found tree diameter to be important for the isopods Porcellium collicola (Figure 3), P. conspersum, and Trachelipus rathkii (Figure 4), all rather common among epiphytic mosses in Hungary.

Figure 3. Porcellium collicola, an isopod that lives among epiphytic mosses. Photo by Dragiša Savić, with permission.

External Anatomy Isopods have two compound eyes (Figure 5) that permit them to detect motion easily. They have a very small head, long thorax, and short abdomen (Figure 6). There are two pairs of antennae, but the first is short and not always visible (Figure 5). That pair may have a chemosensory function to detect odors and tastes (Massey University 2014). The second pair of antennae is large and easily seen; the function is tactile (touch sensation).

Figure 5. Isopod head showing compound eyes. Note the multiple small sections in each eye. Photo from NOAA, through public domain.

Figure 6. Ligia, a genus that sometimes inhabits bryophytes, showing typical isopod external anatomy. Redrawn from Richard Fox.

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Adaptations to Terrestrial Life and to Bryophytes Isopods are predominately aquatic and marine. Life on land requires special adaptations. Even so, some isopod taxa have rather broad niches. Porcellio laevis (Figure 7), a cosmopolitan species and largest member of the genus, at least in the UK (Harding & Sutton 1985), is known for its plasticity in response to the environment (Lardies & Bozinovic 2008), and this plasticity may be the most important adaptation of all. There seem to be few morphological adaptations specific to land dwelling, although one could argue there are no really large species like some of the marine species.

(Figure 13), Porcellio (Figure 7), Cylisticus], and Armadillidium nasatum (Figure 9) to A. vulgare (Figure 8). This order also reflects the progression from most moist to least moist habitat preferences. Armadillidium species further conserve water by curling (Figure 9), a behavioral adaptation that earned it the name of roly poly.

Figure 8. Armadillidium vulgare, the common roly poly that rolls into a ball. Photo from , with permission.

Figure 7. Porcellio laevis, an isopod from which we have learned many terrestrial adaptations. Photo by Roger S. Key, with permission.

Bryophytes make good homes for isopods. These organisms hide from light and require a moist environment, conditions which can be provided by bryophytes. There are probably many species still to be discovered on land, especially among bryophytes, because of the sheltering behavior of isopods in daylight. Water Relations For any organisms evolving from water to land, maintenance of hydration is a critical adaptation. Dias et al. (2013) experimented with 22 species of terrestrial northwestern European isopods to determine the importance of three traits related to desiccation resistance. They found that 90% of the interspecific variation could be explained by water loss rate and fatal water loss. Body surface area affects desiccation resistance through modification of water loss rate. Soil moisture affects species distributions, and by extension, it is likely that bryophyte moisture does as well. Edney (1951a) examined the evaporation of water from woodland isopods and found that in Armadillidium (Figure 8-Figure 9) and Porcellio (Figure 7) it was the pleopods (abdominal appendages also known as swimmerets, Figure 6, Figure 10) that lost water most rapidly, ranging 10-20 times as fast per unit area as the dorsal or ventral surfaces. However, the most water was actually lost from the dorsal and ventral surfaces because of the much greater area. Water loss rates differed among the terrestrial genera tested, in the order from greatest loss to least as Ligia (Figure 11), Philoscia (Figure 12), [Oniscus

Figure 9. Armadillidium nasatum curled into a ball, permitting it to reduce water loss. Photo by Lynette Schimming, through Creative Commons.

Figure 10. Oniscus asellus lying on its back and exposing its pereopods (see Figure 6). The pleopods are on the white abdomen behind these 7 pairs of legs and cannot be discerned in this picture. Note that the head is to the right where you can see two of the antennae. Photo by Brian Eversham, with permission.

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could rehydrate by absorption of water vapor or liquid water, but dead ones could not (Edney 1951a).

Figure 11. Ligia oceanica, member of a coastal genus that readily loses water. Photo by Gilles San Martin, through Wikimedia Commons.

Figure 12. Philoscia muscorum in bark crevice in the forest. Photo by Brian Eversham, with permission.

Figure 13. Oniscus asellus, a frequent moss-dweller in western and northern Europe. Photo by Brian Eversham, with permission.

When terrestrial isopods become desiccated, they can restore their original weight by absorption of moisture through the mouth and anus by contact with free water surfaces, and by mouth from moist surfaces (Edney 1954). This suggests a possible role for the bryophytes as pillbugs traverse such dry habitats as tree bark, rocks, or even soil. They could run from clump to clump of moss, rehydrating when they visit the mosses (or liverworts). Edney (1954) suggests that they are most susceptible to mortality during their wandering rather than while in their selected permanent shelter habitat. Interestingly, living isopods

Waste Elimination Even the elimination of waste products must be modified to conserve water on land. Digestive and bodily processes accumulate nitrogenous wastes, and these are toxic, requiring a means of efficient elimination. In aquatic animals, these are usually eliminated as toxic ammonia that is diluted in water (Dresel & Moyle 1950). But terrestrial animals cannot afford the large quantity of water needed to dilute ammonia to safe levels. Nevertheless, like aquatic isopods, most terrestrial isopods still excrete ammonia, but with a twist. They lack any organ homologous to the kidney or liver to detoxify or facilitate excretion of ammonia (Hartenstein 1968). Hartenstein studied this ammonia elimination mystery in Oniscus asellus (Figure 13) and concluded that rather than excreting liquid ammonia like most aquatic animals, the terrestrial isopods eliminate their ammonia as a gas. In addition, some of the nitrogen waste is stored in the body wall as uric acid and is eliminated during molting. Wieser and Schweizer (1970) likewise found that the terrestrial isopods Oniscus asellus and Porcellio scaber (Figure 1-Figure 2) eliminate their ammonia as gas. Their data refute earlier ideas that nitrogen metabolism is suppressed; instead, they accounted for loss of all the excess nitrogen intake through body wall storage and mostly through the body wall as ammonia gas, thus eliminating the need for large water losses – or kidneys. Osmotic Balance The osmotic pressure of the blood of terrestrial species is somewhat lower than that of sea water and adaptation to land seems to be achieved by osmotic tolerance rather than regulation (Edney 1954). Nevertheless, Porcellio scaber does not change its body fluid concentration as rapidly as it loses weight during desiccation (Horowitz 1970), implying it could have a limited balancing mechanism. Lindqvist and Fitzgerald (1976) explored this further and determined that initially the blood osmotic concentration remains essentially unchanged until about a 10% loss of body weight. Meanwhile, the oral fluid increases its osmotic concentration rapidly during about 90 minutes of drying. When severe desiccation occurs, these two compartments progress to an osmotic equilibrium, presumably due mostly to withdrawal of water from the gut lumen into the blood. Molting has the potential to affect the osmotic balance. Calcium is an important element in the exoskeleton. Before the animal molts, the calcium is resorbed and stored in the body of terrestrial isopods and little is lost, whereas in aquatic taxa, little is resorbed and most of the exoskeleton calcium is lost (Greenaway 1985). When needed, additional calcium is gained from food and exuviae (shed exoskeleton). Despite this resorption of high amounts of calcium in terrestrial species, most of it is not stored in ionic form and thus has little effect on the osmotic balance. Respiration Terrestrial isopods have pseudotracheae, assisting them with respiration in dry air (Edney 1954), whereas the importance of integumental oxygen absorption decreases in terrestrial species compared to aquatic species. The inner

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branch of each pleopod (Figure 6, Figure 10) is modified into a gill-like structure (Figure 14) with a thin, permeable cuticle where gas exchange occurs (Schotte et al. 20082014). These even somewhat resemble lungs in the terrestrial isopods.

and this at least partly explains their need for moist environments. On the other hand, if the air is saturated, they are unable to use evaporative cooling, and higher temperatures become lethal. Both temperature and moisture needs explain the migration of the isopods to deeper moss layers or even into the soil in the daytime, returning to the surface for feeding at night. Behavior The best adaptations of this group seem to be behavioral (Edney 1954), and these behaviors are what make them so interesting to watch. Pick up a rock and pillbugs scramble in all directions, soon disappearing under leaves or into the soil. They run from light, which might be an indicator of drying conditions. They seem to lack a well-developed cuticle, although both endocuticular and epicuticular layers are known from some species (Edney 1968), and thus they are able to use evaporative cooling, but this only works for a short time, hence making a behavioral solution essential. Edney (1968) suggests that the nightly activity of Porcellio scaber (Figure 1-Figure 2) on trees may permit them to transpire excess water. Armadillidium vulgare (Figure 8) also has greater activity at night when the air is more moist. It appears that males of Porcellio scaber and Armadillidium vulgare use surface shelters, including bryophytes, between foraging events (Dangerfield & Hassall 1994), sometimes providing them with a location to gain or reduce water content.

Figure 14. Porcellio siculoccidentalis pleopods modified to function in gas exchange and resembling lungs. Photo by Giuseppi Montesanto, with permission.

Temperature Tolerance The temperature tolerance follows the same sequence of genera as for water loss rates above [least in Ligia Figure 11 < Philoscia - Figure 12 < (Oniscus - Figure 13, Porcellio - Figure 7, Cylisticus) < Armadillidium (Figure 8)], with Armadillidium having the highest temperature tolerance (Edney 1951b). There was no difference in body temperature between living and dead woodlice, and once the animals reached equilibrium their temperatures differed from that of the air by no more than 0.1°C in moist air. However, in dry air the isopod temperatures were depressed relative to air temperature, apparently due to evaporative cooling.

Congregating Behavior Aggregating or congregating (Figure 15) in large numbers in a suitable habitat, as is easily observed under a log, board, or small rock, is generally accepted as a means to reduce their water loss to the atmosphere (Broly et al. 2013). This behavior is mostly thigmotactic (a contact response), and possibly olfactic (an odor response) (Edney 1968). Olfaction seems to play a role in seeking shelter. But the role of aggregation in preventing water loss may be misleading. Broly and coworkers suggest other potential benefits, including reduction of oxygen consumption, increase in body growth, stimuli for reproduction, better access to mates, shared predator defense, promotion of coprophagy, sheltering, and acquisition of internal symbionts. They suggest that congregating behavior provides terrestrial isopods with a non-physiological alternative to coping with climate constraints.

Moisture and Temperature Interaction Temperature and moisture rarely act alone in ecosystems, and responses by isopods to one of these typically depends on the other. In experiments with the isopods discussed above, Edney (1951b) found that after 30 minutes in dry air at 20° and 37°C, mean temperature depressions were for Ligia (Figure 11), 2.6°C and 6.8°C; Oniscus (Figure 13), 1.5 and 2.7°C; Porcellio (Figure 7), 0.4 and 1.3°C; Armadillidium (Figure 8), 0.5 and 1.8°C, respectively. Ligia differed from the others, with its body temperature rising for at least 2 hours, whereas the others reached equilibrium at a temperature lower than ambient air temperature after 25 minutes. It is the ability to evaporate water rapidly that permits these isopods to maintain a safe temperature for short intervals,

Figure 15. Isopod congregation. Photo by William Leonard, with permission.

Chapter 10-3: Arthropods: Crustacea – Isopoda, Mysida, and Decapoda

But crowding does not seem to have the same benefit for all terrestrial isopods. Armadillidium nasatum (Figure 16) and A. vulgare (Figure 8), members of the most xeric genus, had reduced growth rate, survivorship, and size at first reproduction as density increased in laboratory experiments (Ganter 1984). Since limited food reduced both growth rate and mortality in these experiments, these same detrimental factors might not exist in nature where foraging might be unlimited.

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Sheltering Sheltering (staying in one place that is protected) is common among some isopods, but not others (Hassall & Tuck 2007). Porcellio scaber (Figure 1-Figure 2), a common moss dweller, sheltered significantly more than either Platyarthrus hoffmannseggi (Figure 18) or Armadillidium vulgare (Figure 8), and Philoscia muscorum (Figure 17) sheltered the least, despite some individuals spending at least winter under mosses. Sheltering declined in all four species after the breeding season, continuing through winter. Porcellio scaber sheltered more where the soil was more calcareous (occurring on chalk or limestone), Philoscia muscorum more under the shade of trees, and both P. muscorum and Armadillidium vulgare more in grazed than in ungrazed areas. For A. vulgare sheltering was positively correlated with both rainfall and temperature of the day before sampling, whereas for Philoscia muscorum it was negatively correlated with rainfall.

Figure 16. Armadillidium nasatum showing two color variants. Photo by Stan Gilliam, through Creative Commons.

To put this in perspective, Hassall et al. (2010) experimented with aggregation behavior in Philoscia muscorum (Figure 17), Oniscus asellus (Figure 13), Porcellio scaber (Figure 1-Figure 2), and Armadillidium vulgare (Figure 8) from Norwich, UK. The first three are isopods known from terrestrial bryophytes, whereas Armadillidium vulgare tends to occur in drier habitats. The first three species clump more at lower levels of relative humidity and at higher temperature, whereas changing the humidity has little effect on clumping in A. vulgare.

Figure 18. Platyarthrus hoffmannseggi, an isopod that shelters under mosses. Photo by Jan van Duinen , with permission.

Dias et al. (2012) examined the influence of microclimate on sheltering in three terrestrial isopods: Porcellio scaber (Figure 1-Figure 2), Oniscus asellus (Figure 13), and Armadillidium vulgare (Figure 8, Figure 23). The first two are common among mosses, whereas A. vulgare typically lives in drier habitats. All three species spent more time sheltering and less in activities when the environment was drier (50% relative humidity) compared to more moist conditions (90% relative humidity). Oniscus asellus is the least terrestrialized of these three and thus the most susceptible to desiccation. Sheltering can also reduce the quality of food consumed because less time is spent on foraging. Reproduction

Figure 17. Philoscia muscorum on moss. Photo by Dick Jones, with permission.

Reproduction among terrestrial invertebrates usually requires modifications from that of aquatic taxa. Terrestrial isopods carry their young in a marsupium (brood pouch, Figure 19). The marsupium is filled with fluid and the eggs and embryos are surrounded by mucous. Warburg (1987) considers this to be one of the most important innovations for successful living on land. The mucous may contribute to nourishment of the young, possibly explaining their ability to survive when the mother doesn't eat.

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Chapter 10-3: Arthropods: Crustacea – Isopoda, Mysida, and Decapoda

Figure 19. Armadillidium vulgare lying on its back, showing young (cream-colored) isopods in a brood pouch on the ventral side. Photo by Malcolm Storey, through Creative Commons.

In the isopods, gonadal development is stimulated by a long photoperiod and high temperatures (Edney 1968). Temperature seems to play a role in controlling reproductive output and consequent water loss. Females are dominant throughout most of the year in Porcellio scaber (Figure 1-Figure 2) (Nair 1998), and this is likely true in other species as well, sometimes indicating parthenogenesis (reproduction from an egg without fertilization). Some species that exhibit parthenogenesis do not necessarily do so in their populations everywhere (Christensen 1979; Fussey & Sutton 1981; Fussey 1984). For example, in the British Isles some populations of an isopod that often lives among bryophytes, Trichoniscus pusillus (Figure 25), are parthenogenetic and others are not. Christensen (1979) demonstrated that the parthenogenetic populations represented different genotypes in this species. Fussey (1984) was unable to find a relationship between this parthenogenetic expression and latitude, longitude, altitude, or seven climatic variables, but it did correspond with calcareous habitats. But genes are not the only causes of alteration in the reproductive type. The bacterium Wolbachia pipientis is able to infect the isopods Hyloniscus riparius (Figure 20), Trachelipus rathkii (Figure 26), and Trachelipus ratzeburgii (Figure 21) (Nyirő et al. 2002), all species known to inhabit mosses (Božanić 2011). The bacterium lives in the ovaries and can cause such changes as loss of maleness and shift to parthenogenesis in these isopods. The bacterium also infects the eggs and thus is transferred from mother to offspring. Could the antibiotic properties of bryophytes protect the isopods from this populationaltering bacterium?

Figure 20. Hyloniscus riparius, whose gender is altered by the bacterium Wolbachia pipientis. Photo by Dragiša Savić, with permission.

Figure 21. Trachelipus ratzeburgii, an isopod whose gender is altered by the bacterium Wolbachia pipientis. Photo by Dragiša Savić, with permission.

Food quality can have a strong effect on the success of both reproduction and survival of the offspring. For example, Kautz et al. (2000) were only able to maintain a stable population of Trichoniscus pusillus (Figure 25) on a diet of Alnus litter with high microbial activity. Such needs may explain changes in the diet of isopods throughout the year. It would be interesting to test the effect of a bryophyte diet on reproductive success. On the other hand, Lavy et al. (2001) found that in Porcellio scaber (Figure 1-Figure 2) and Oniscus asellus (Figure 13) diet had no effect on the number of juveniles or their weight. Rather, the weight of the offspring was correlated with the weight of the female. Nair (1998) found that for Porcellio scaber in Benghazi, Libya, the total number of eggs correlated with body length of the female. High temperatures can be lethal or detrimental to developing isopods. In the terrestrial Porcellio ficulneus, at 25°C, oocytes matured sooner, and many were resorbed (Hornung & Warburg 1993). The Mediterranean population compensated for these losses by breeding earlier. Females must balance the advantages of faster brood development in higher temperatures with the risk of excessive water loss (Dangerfield & Hassall 1994). Incubation periods for Porcellio scaber (Figure 1Figure 2) in Benghazi were 18 days in summer and autumn but extended to 32 in late winter and spring (Nair 1998). Spring embryo production was higher in spring compared to summer and autumn. In Armadillidium vulgare (Figure 8), if females are dehydrated, they reproduce instead of growing (Warburg 1987). Terrestrial isopods care for their young, an uncommon feature in the aquatic habitat (Lardies et al. 2004). Such care can be costly energetically, but it increases the survival of the young in the terrestrial environment, and it might even reduce water loss of the adult, much like the congregating behavior. But there is a downside. Lardies and coworkers found that in Porcellio laevis (Figure 7) not only was the carrying of developing eggs energetically costly, the females carrying them had a lower ingestion rate and lower ability to digest food than non-carrying females. The net result was that egg-carrying females stored only about 20% as much energy as females with no eggs.

Chapter 10-3: Arthropods: Crustacea – Isopoda, Mysida, and Decapoda

Carrying eggs and young creates other problems for these woodlice. Females carrying broods were slower and moved shorter distances to avoid light than non-brooding females of Porcellio laevis (Figure 7) (Kight & Nevo 2004). Physical stress causes a reduction in both distance travelled and velocity in brooding females. Many eggs and embryos die before reaching their swimming stage. These are typically eaten by their siblings in the marsupium and larger larvae often eat the smaller ones (Warburg 1987). Once the young leave the marsupium they begin a life free of their mother. Predators Bryophytes can serve as a refuge for hiding from large predators like birds, but they may not be so safe from insect predators. Ants such as Tetramorium caespitum (Figure 22) influence the behavior of the isopods Armadillidium vulgare (Figure 8) and Porcellio laevis (Figure 7) (Castillo & Kight 2005). Armadillidium vulgare females were hidden better than those of P. laevis whether ants were present or not. But some of their behavior was rather strange. Isopods that had no experience with ants remained further from them than those with previous exposure, with P. laevis keeping a significantly greater distance than that of A. vulgare. This difference in behavior of the two species may be explained by the ability of A. vulgare to roll into a ball (Figure 23), whereas P. laevis is endowed with the ability of rapid locomotion.

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such exposure had no effect on brood time for Porcellio laevis (Figure 7) (Castillo & Kight 2005). This is a greater advantage for A. vulgare because it is unable to roll into a sealed ball when it is carrying its brood. Nevertheless, P. laevis is slowed down when carrying a brood (Kight & Ozga 2001; Kight & Nevo 2004). Overwintering It is difficult to find information on the use of bryophytes for overwintering of crustaceans. Samouelle (1819) reported that one could find Philoscia muscorum (Figure 17) under mosses in January in Great Britain. Le Gay Brereton (1957) reported that the isopod Porcellio scaber (Figure 1-Figure 2, Figure 24) overwintered "in large numbers" in the moss layers at the bases of oaks (Quercus) and ash (Fraxinus). These same aggregations did not occur at eye level, suggesting that the larger moss clumps at the tree bases were more suitable than the small clumps or shallow mats of the bole. One would presume that the tree base had both warmer and less desiccating conditions than any position on the bole.

Figure 24. Porcellio scaber, a common moss dweller and consumer that eats its own feces to assimilate more nutrients. Photo by Eric Schneider, with permission.

Figure 22. Ant Tetramorium caespitum eating larva, most likely of an insect. This ant causes soil isopods to stay hidden. Photo from Antwiki, through Creative Commons.

Figure 23. Armadillidium vulgare in a ball, a protection against predators. Photo from , with permission.

The brooding period of Armadillidium vulgare (Figure 8) was shortened when it was exposed to ants, whereas

Terrestrial isopods are not well adapted to cold temperatures and must seek locations where they are insulated from the cold. Porcellio scaber (Figure 1-Figure 2, Figure 24), a common species that is known from bryophytes and under many other objects, is able to adjust somewhat by acclimation, but is nevertheless susceptible to both freezing and chilling (Tanaka & Udagawa 1993). The temperature causing 50% mortality was -1.37°C in August but dropped to -4.58°C in December. At -7°C, the animal was unable to avoid freezing of its tissues, a temperature limit that was the same throughout the year. The winterization in Porcellio scaber (Figure 1-Figure 2, Figure 24) corresponded to the presence of low molecular weight carbohydrates that may have protected it against chilling injury (Tanaka & Udagawa 1993). The supercooling temperature of -7°C seemed to be associated with the year-round gut content. We know that at least in the autumn this species can live among mosses and deciduous and conifer leaf litter where it prefers mosses as food (Hribljan 2009; Hribljan & Glime in prep). Could the mosses help to prepare it for winter by contributing arachidonic acids that have lower freezing points (see Prins 1982)? Hansen and Rossi (1991) showed that Rhytidiadelphus triquetrus (Figure 49), a food of Porcellio scaber in autumn (Figure 53; Hribljan 2009; Hribljan &

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Chapter 10-3: Arthropods: Crustacea – Isopoda, Mysida, and Decapoda

Glime in prep), contains 30% arachidonic acid at 20°C, which slightly decreases at lower temperatures. Tanaka and Udagawa (1993) also suggest that food derivatives could serve as ice nucleation centers that could reduce freezing of tissues.

Bryophytes as Food In the other crustaceans, we have seen that bryophytes serve mostly as trapping devices, collecting detritus and growing periphyton that can serve as food for the crustaceans. The crustaceans have carried their aquatic habit of eating detritus into the terrestrial environment. Isopods are also litter-dwelling organisms that eat litter, but they eat bryophytes too. Digestion Generally bryologists might not care much about the digestive process of a bryophyte dweller, but understanding isopod digestion helps us explain bryophyte herbivory and why isopods can be such good bryovores whereas other invertebrates generally are not. As organisms derived from aquatic ancestry, isopods required adaptations to digest terrestrial food. They are among the few organisms known to readily eat bryophytes. Food sources on land can differ somewhat for isopods, although leaf litter, a common food for them, is available in streams and lakes as well as on land. Hames and Hopkin (1989) observed the digestive tracts of two terrestrial isopods known from mosses, Oniscus asellus (Figure 13) and Porcellio scaber (Figure 1-Figure 2, Figure 24), and determined that their digestive tracts are divided into five regions: foregut, anterior chamber, papillate region, rectum, and hepatopancreas. The latter opens into the foregut. There is a powerful muscular sphincter between the papillate region and the rectum. As food passes from the foregut to the hindgut it is mixed with secretions from the hepatopancreas (Hames & Hopkin 1989). When the hindgut is full, muscles contract to force the liquids and fine food particles back to the foregut through special channels. This re-entry can occur several times, each time being subject to further degradation by the microbial gut flora. Material left in the hindgut passes to the rectum where the fecal pellets are compacted for expulsion. And like a rabbit, Armadillidium vulgare (Figure 8) and Porcellio scaber (Figure 1-Figure 2, Figure 24) eat their own fecal pellets, preferring fresh ones to decaying ones (Hassall & Rushton 1982). Hames and Hopkin (1989) suggest that the ability to recycle the food, each time extracting more liquid, may be one of the major changes making their life on land successful. Isopods are good at digesting their food but poor at assimilating it. This may also help explain their ingestion of feces, to gain more nutrients from it (Warburg 1987). Food quality is important for growth (Merriam 1971), and fresh leaves are better for growth than decayed leaves (Beck & Bretowsky 1980). But we are left with the question of circumventing the high phenolic content of bryophytes and some types of leaf litter. Phenolic compounds are known for their production as a stress response in bryophytes (Graham et al. 2004). There is ample evidence that they deter herbivory in both

terrestrial and aquatic bryophytes, as discussed below. But it appears that not all isopods are created equal in their tolerance of phenolic compounds. And not all bryophytes are equal in making them. Zimmer (1997) showed that the common moss dweller Porcellio scaber (Figure 1-Figure 2, Figure 24) has significant ability to reduce gut surface tension. Phenolic compounds, well known to prevent digestion in other invertebrates due to the ability of the phenolics to increase the surface tension, seem to have a less negative effect on this species. These surfactants may be the key to the ability of Porcellio scaber to eat mosses without suffering from the typical binding of proteins suffered by many other kinds of organisms that eat phenolics. In insects, the phenolics precipitate proteins in the diet, preventing the insects from assimilating these essential nutrients, but in the isopods the surfactants bind the phenolics, leaving the proteins free for assimilation by the isopods. The concentration of surfactants in Porcellio scaber was 80 times as high as the "critical micelle concentration" needed to permit binding of the phenolics. Further research on Porcellio scaber (Figure 1-Figure 2, Figure 24) indicated that endosymbiotic bacteria residing in the hepatopancreas were able to oxidize the phenolics, disabling their adverse properties (Zimmer 1999). When the gut flora of Porcellio scaber was reduced, Zimmer demonstrated that bacteria in the gut apparently had an important role in hydrolyzing gallotannins. When galloylglucose esters were ingested, they greatly reduced the microbial component of the hindgut. Ingestion of gallic acid reduced both palatable fungi and bacteria, but not as strongly, and increased the gut microflora. Zimmer's study suggests that the ingestion of hydrolyzable tannins, as found in some mosses, can inhibit the digestion of other foods in the diet of this species. The gut differences among the isopod species can account for their preferences among bryophytes, and possibly account for those taxa that don't eat bryophytes at all. Similarly, differences in hydrolyzable tannin concentrations among bryophyte species can account for preferences for some bryophytes over others. Zimmer and Brune (2005) examined the physiological properties of the gut of four species of terrestrial isopods [Oniscus asellus (Figure 13), Porcellio scaber (Figure 1Figure 2, Figure 24), Trichoniscus pusillus (Figure 25), and Trachelipus rathkii (Figure 26)]. These adaptations were manifest as a steep gradient of oxygen, high at the periphery and low at the center of the gut transection. This gradient provides suitable habitat for both aerobic and anaerobic symbionts that can contribute to digestion. The pH gradient ran from acidic in the anterior hindgut to neutral in the posterior hindgut of O. asellus, P. scaber, and T. rathkii. In Trichoniscus pusillus, the pH in the hindgut lumen was nearly constant. Zimmer and Brune (2005) suggested that the pH gradient differences may be adaptive in providing differences in the digestion of lignocellulose from their food sources. Bryophytes lack true lignin, so the expenditure of resources to create the conditions suitable for digesting lignin could be spared in those isopods that eat mosses. These differences in gut physiology could also account for some of the differences in food preferences and survival of isopods on bryophytes vs other foods.

Chapter 10-3: Arthropods: Crustacea – Isopoda, Mysida, and Decapoda

Figure 25. Trichoniscus pusillus, a terrestrial isopod. From , with permission.

Figure 26. Trachelipus rathkii, a terrestrial isopod. Photo by R. E. Jones, with permission.

Terrestrial Consumers Terrestrial isopods seem to prefer a varied diet and exhibit food preferences (Dudgeon et al. 1990). Dudgeon and coworkers found that four species of isopods from a Hong Kong forest ate more food when given a mixture of leaves than when only one type was available. Their preferences did not seem to relate to ash, calcium, copper, soluble tannin, or energy content. Dudgeon and coworkers suggested that the isopods satisfied their nutrient requirements by consuming certain foods, then switching to others to avoid excessive tannins or other allelochemicals. Rushton and Hassall (1983a, b) examined the feeding preferences and rates of Armadillidium vulgare (Figure 8) among dicotyledonous and monocotyledonous plants and bryophytes (Calliergonella cuspidata, Figure 27). This pillbug, known as a roly poly due to its ability to roll into a ball, can live in drier habitats than Porcellio and is much less likely to be associated with mosses. These isopods initially preferred the dicotyledonous plants to the other two choices. But after the monocotyledonous plants began to decay, these were preferred. Nevertheless, eating monocots increased mortality and drastically reduced growth rates and reproductive output, even when it was in a later decay state. Defenses in the food become more concentrated as the food decays and carbon sources are removed. Chemical defenses in mosses may play a role in the isopod choice of leaf litter over mosses in Armadillidium vulgare.

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Figure 27. Calliergonella cuspidata, a moss that seems to deter feeding by Armadillidium vulgare. Photo by J. C. Schou, through Creative Commons.

Rushton and Hassall (1983a) suggested that Armadillidium vulgare (Figure 8) compensates for low quality food by eating more, but that plant defenses can interfere with this compensation. Even though the moss is likely to provide a suitable moist habitat, and Armadillidium vulgare may be able to absorb at a high rate on low quality food by increasing its rate of consumption, it appears that plant defenses of Calliergonella cuspidata (Figure 27) might outweigh its habitat desirability (Rushton & Hassall 1983a). Dead mosses may be less desirable than dead tree leaves or even monocot leaves, particularly after the tracheophyte leaves begin to decay. It is likely that very little nutritional material is available relative to cell wall material in dead mosses (see Pakarinen & Vitt 1974 for lower N content), especially if nutrients are moved from dead portions to living portions, but that relationship requires further testing.

Figure 28. Hypopterygium didictyon from Chile, a moss in the same genus as one grazed in Costa Rica. Photo by Juan Larrain, with permission.

Nevertheless, at times isopods can be voracious consumers of bryophytes. Angela Newton (Bryonet, 20 November 2006) reported seeing extensive grazing on Hypopterygium sp. (Figure 28) in the montane rainforest of Costa Rica. The isopods sheared off the green lamina and left the branches and costa, much like the feeding behavior

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Chapter 10-3: Arthropods: Crustacea – Isopoda, Mysida, and Decapoda

of insects on tracheophyte leaves. However, grazing of isopods and silverfish on damp herbarium labels and plants in packets made her question whether they were simply feeding on the associated fungi and consumed the mosses in the process. Isopods can be downright pests on garden mosses. Henk Greven, in a communication to Bryonet (23 October 2003) writes: "Apart from mammals, birds and slugs, sowbugs (Oniscus asellus L.; Figure 13) are fervent bryophyte eaters. In my garden, I keep several Grimmia species on pieces of rock. When I put these on the ground, sow-bugs are hiding during the day under these rocks. In the evening they climb above and start eating my Grimmias. They have a special preference for Grimmias growing on limestone, basic sandstone, and basic basalt. In no time, they have eaten all my Grimmia plagiopoda (Figure 29), G. crinita (Figure 30), and G. orbicularis (Figure 31). Species on acidic rock, however, are not safe either. The only species they don't like is Ptychomitrium polyphyllum (Figure 32). I had this species nearly ten years on a piece of rock on my garden floor. I learnt my lesson and now I keep pieces of rock on a table where they are safe from sow-bugs."

Figure 29. Grimmia plagiopoda, a species that seems to be preferred food for Oniscus asellus on limestone rocks. Photo by Michael Lüth, with permission.

Figure 30. Grimmia crinita, a species that seems to be preferred food for Oniscus asellus. Photo by Michael Lüth, with permission.

Figure 31. Grimmia orbicularis with capsules, growing on rock. This seems to be a preferred food for Oniscus asellus in limestone habitats. Photo by Michael Lüth, with permission.

Figure 32. Ptychomitrium polyphyllum, a moss that is not eaten by Oniscus asellus. Photo by David T. Holyoak, with permission.

Likewise, I have already reported above on my own sad experience with Porcellio scaber (Figure 24) eating my carpet of mosses so that it looked like Swiss cheese. And Daniel Marsh (Bryonet, 18 November 2006) reported that wood lice (isopods) have usually consumed any liverwort he tried to cultivate in his garden or greenhouse. "The attraction seems to be immediate." In contrast, he reports that he has not noticed such consumption of liverworts by isopods in wild communities. We (Weston 1995; Liao & Glime unpubl) attempted to find out what sorts of things might deter pillbugs (Porcellio scaber (Figure 24). Using Polytrichum juniperinum (Figure 33) and P. commune (Figure 34) from Houghton, MI, USA, we compared consumption of stems and leaves. Polytrichum juniperinum leaves were consumed 3:1 over stems; P. commune leaves were consumed 5.5:1 over stems (Figure 35). It made no difference whether the leaves were still connected to the stems or not.

Chapter 10-3: Arthropods: Crustacea – Isopoda, Mysida, and Decapoda

Figure 33. Polytrichum juniperinum, a species in which Porcellio scaber prefers eating leaves over stems. Photo by Li Zhang, with permission.

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In both Polytrichum juniperinum (Figure 33) and P. commune (Figure 34), the leaves had roughly double the protein content per dry weight compared to the stems (Figure 36) (Weston 1995; Liao & Glime unpubl), suggesting that the isopod Porcellio scaber (Figure 24) could gain more protein nutrition from eating leaves. And structurally leaves would seem to be easier to chew than the tough stems endowed with thickened walls and coloration suggesting phenolic compounds. But it is surprising to find that the leaves of at least P. commune seem to have a higher concentration of phenolic compounds than do their stems (Figure 37), yet that species had the higher consumption ratio of leaves to stems. Perhaps the presence of folded-over leaf edges in Polytrichum juniperinum (Figure 38), absent in P. commune (Figure 39), makes it easier to obtain the nutritious photosynthetic lamellae in P. commune (Figure 39).

Figure 34. Polytrichum commune, a species in which Porcellio scaber prefers eating leaves over stems. Photo by David T. Holyoak, with permission.

Figure 36. Comparison of mean protein ± 95% CI in stems and leaves of two Polytrichum species. Based on Weston 1995; Liao & Glime unpublished data; Bradford's (1976) test, n = 3.

Figure 35. Comparison of mean isopod (Porcellio) consumption ± 95% CI of excised leaves and stems vs intact leaves and stems in two species of moss. Data based on unpublished laboratory data of Weston 1995; Liao & Glime unpublished data; n = 3.

Figure 37. Comparison of mean phenolic content ± 95% CI in stems and leaves of two Polytrichum species. Based on Weston 1995; Liao & Glime unpublished data; Folin-Denis test (Swain & Hillis 1959) and Prussian Blue test for tannin; n = 3.

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Chapter 10-3: Arthropods: Crustacea – Isopoda, Mysida, and Decapoda

Figure 38. Polytrichum juniperinum leaf cross section showing margin of leaf rolled over the lamellae, partially covering them. Photo by John Hribljan, with permission.

Figure 39. Polytrichum commune leaf cross section showing absence of rolled over leaf margin, thus giving exposure to all the lamellae. Photo by Kristian Peters, through Creative Commons.

Isopods can be a major inhabitant of bryophytes, becoming especially obvious at night when they migrate to the surface to feed (Hribljan & Glime in prep.). But even the isopods are fussy about which bryophytes they eat. Phenolic content seems to deter isopod consumption of various leaves (Warburg 1987). But deterrents may not be the only answer to these food preferences. Porcellio scaber (Figure 24) will eat Thuidium delicatulum (Figure 40-Figure 41) ravenously, but ignore Dicranum polysetum (Figure 42) and sometimes ignore Pleurozium schreberi (Figure 43) (Liao 1993; Glime 2006). When faced with Polytrichum (Figure 33), they eat the leaves, but little of the stems (Liao 1993, unpublished data; Hribljan 2009). This suggests that chemistry might be more important than structure, as Thuidium delicatulum is a crunchy moss with papillae (but small leaves, Figure 41) whereas P. schreberi and D. polysetum are softer and more flexible, lacking papillae (but with large leaves). But it appears that we may not have examined enough potential deterrents in Thuidium. And we need to beware of differences between populations and seasons. Fatoba et al. (2003) found that whereas Thuidium gratum from the Nigerian tropics lacked detectable phenolics, it had tannins, alkaloids, and cardiac glycosides. In a different location in tropical Nigeria (and a different date), Adebiyi et al. (2012) found that this same species had a high content of saponins (absent in the Fatoba et al. 2003 study) and flavonoids, but also had a very low content of phenolics. Perhaps isopods, like many humans, just prefer a crunchy snack.

Figure 40. Thuidium delicatulum, a moss readily eaten by Porcellio scaber. Photo by Bob Klips, with permission.

Figure 41. Thuidium delicatulum branch leaf showing small cells and papillae (note bumps on cells). Photo from Dale A. Zimmerman Herbarium, Western New Mexico University, with permission.

Figure 42. Dicranum polysetum, a moss that is ignored, not eaten, by Porcellio scaber. Photo by Bob Klips, with permission.

Figure 43. Pleurozium schreberi, a moss that is sometimes eaten and sometimes ignored by the wood louse Porcellio scaber. Photo by Janice Glime.

Chapter 10-3: Arthropods: Crustacea – Isopoda, Mysida, and Decapoda

Hribljan and Glime (in prep) explored the food preferences of populations of Porcellio scaber (Figure 24) in the Keweenaw Peninsula of Michigan, USA. In food preference experiments, these isopods preferred the moss Pleurozium schreberi (Figure 44) over leaf litter from Acer saccharum (Figure 45) and Pinus strobus (Figure 46) in each of three study months of September – November (Figure 47). They compared the carbohydrates, proteins, and phenolics in these three species for the three months of the feeding trials and found that Pleurozium schreberi had the lowest levels of phenolics and highest levels of carbohydrates of the three choices of food (Figure 48). This is interesting because some studies (e.g. Pakarinen & Vitt 1974) have suggested that mosses were unable to provide enough energy for herbivores, but it appears that compared to leaf litter the mosses may, at least at times, have more carbohydrates than litter and be preferred food for isopods. Furthermore, all five mosses tested [Pleurozium schreberi, Thuidium delicatulum (Figure 40Figure 41), Polytrichum juniperinum (Figure 33), Rhytidiadelphus triquetrus (Figure 49), and Dicranum polysetum (Figure 42)] had higher carbohydrate contents than the leaf litter of the trees tested (Figure 50-Figure 51). However, protein was higher in both types of tree leaf litter tested compared to that of Pleurozium schreberi (Figure 48).

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Figure 46. A common sight of needles of Pinus strobus (white pine) mixed with the moss Pleurozium schreberi. The needles are a food less preferred in September to November by Porcellio scaber than the moss Pleurozium schreberi. Photo by Janice Glime.

Figure 44. Pleurozium schreberi showing damage from Porcellio scaber that feeds on it at night. Note the less green plants on left that have suffered considerable damage. On the right you can see naked red stem tips where leaves and buds have been eaten. Photo by John Hribljan, with permission. Figure 47. Comparison of mean air-dried mass (±95% CI) consumed by isopods in 24 hours when given the choice of the moss Pleurozium schreberi and the tree leaves of Acer saccharum and Pinus strobus. The same letters signify means that are not significantly different from each other (α = 0.05 post two-way ANOVA & Tukey test, n = 10). Hribljan 2009; Hribljan & Glime in prep.

Figure 45. Freshly fallen Acer saccharum (sugar maple) leaves, a food source less preferred by Porcellio scaber than the moss Pleurozium schreberi in September to November. Photo by Janice Glime.

Based on these experiments, Hribljan and Glime (in prep) compared the preferences among five species of mosses that occurred within the foraging distance of the isopods. Porcellio scaber (Figure 24) significantly preferred the moss Pleurozium schreberi (Figure 44) to the mosses Rhytidiadelphus triquetrus (Figure 49), Thuidium delicatulum (Figure 40), Dicranum polysetum (Figure 42), and Polytrichum juniperinum (Figure 50), with Pleurozium schreberi and Rhytidiadelphus triquetrus having lower phenolic concentrations than Dicranum polysetum and Polytrichum juniperinum (Figure 51). The

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Chapter 10-3: Arthropods: Crustacea – Isopoda, Mysida, and Decapoda

Thuidium delicatulum, preferred in earlier experiments over Pleurozium schreberi (Liao 1993; Glime 2006), was not among the top preferences, perhaps due to its lower carbohydrate content at a time of year when the isopods were preparing for winter.

Figure 50. Comparison of moss consumed (mean ±95% CI) with mean percent by weight of phenolics, proteins, and carbohydrates in leaves of the mosses Pleurozium schreberi (PLE), Thuidium delicatulum (THU), Polytrichum juniperinum (POL), Rhytidiadelphus triquetrus (RHY), and Dicranum polysetum (DIC). n = 10. Hribljan 2009; Hribljan & Glime in prep.

Figure 48. Comparison of percent of carbohydrates, proteins, and phenolics in freeze-dried leaves of the moss Pleurozium schreberi, sugar maple tree Acer saccharum, and white pine Pinus strobus. Samples were taken once each month during to compare stages of decay in the tree leaves. Values are means of 10 samples. Redrawn from Hribljan & Glime (in prep).

Chemical analysis revealed that P. schreberi contains a high protein:phenolic ratio (Figure 55) (Hribljan & Glime in prep). Despite the high phenolic content and low protein content of Rhytidiadelphus triquetrus (Figure 49), these isopods would still consume it (Figure 52-Figure 53), perhaps for its high carbohydrate content, but it was not a preferred food (Figure 50-Figure 51). On the other hand, the feces indicated that this moss had not been well digested (Figure 54). As a terrestrial moss, it collects only minimal detritus, suggesting that it could have limited food value. Dicranum polysetum was least preferred despite a relatively high carbohydrate content (Figure 50-Figure 51).

Figure 49. Rhytidiadelphus triquetrus, a less preferred bryophyte as autumn food for Porcellio scaber, growing as it typically does amid leaf litter. Photo by Michael Lüth, with permission.

Figure 51. Comparison of means ±95% CI of phenolics, proteins, and carbohydrates in leaves of the mosses (arranged from most to least eaten) Pleurozium schreberi (PLE), Thuidium delicatulum (THU), Polytrichum juniperinum (POL), Rhytidiadelphus triquetrus (RHY), and Dicranum polysetum (DIC). n = 10. Bars with the same letters are not significantly different (α=0.05, n=10).

Figure 52. Branches of Rhytidiadelphus triquetrus that have been nibbled by Porcellio scaber. Photo by John Hribljan, with permission.

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mosses despite phenolics, whereas other arthropods like the cranefly Tipula montana, a moss-food-avoider, cannot (Smith et al. 2001).

Figure 55. The mean protein:phenolic ratio of leaves (± 95% CI), arranged in order from most to least consumption, of freezedried mosses Pleurozium schreberi (PLE), Thuidium delicatulum (THU), Polytrichum juniperinum (POL), Rhytidiadelphus triquetrus (RHY), and Dicranum polysetum (DIC). n = 10; bars with the same letter are not significantly different, post ANOVA Tukey test, α = 0.05).

Figure 53. Moss branches of Rhytidiadelphus triquetrus being eaten by Porcellio scaber. Photos by John Hribljan, with permission.

We cannot rule out the possible importance of carbohydrates, and Forman (1968) provides evidence that caloric content is highest in two of the mosses that seem to be preferred in our experiments (Hribljan & Glime in prep). Forman showed that Thuidium delicatulum (Figure 40) had the highest caloric value (4305 cal/gdw) among the ten mosses he tested; Pleurozium schreberi (Figure 43) had the second highest caloric content (4240 cal/gdw), fitting with our data on carbohydrates. On the other hand, the lowest content was that of Dicranella heteromalla (Figure 56) (3749 cal/gdw), a moss in the same family as Dicranum polysetum (Figure 42), the latter being least preferred in our experiments. Furthermore, Sveinbjörnsson and Oechel (1991) found that the carbohydrate concentration varied with season in Polytrichum commune (Figure 34), but not in Polytrichastrum alpinum (Figure 57). Could it be that some bryophytes become more desirable in autumn due to higher carbohydrate concentrations?

Figure 54. Moss leaf fragments extracted from feces of Porcellio scaber fed only Rhytidiadelphus triquetrus. Photo by John Hribljan, with permission.

Hribljan (2009) suggested that the protein:phenolic ratio might be more important in determining isopod herbivory than concentration of phenolic compounds alone. In this case, Pleurozium schreberi (Figure 44) had the highest ratio of proteins:phenolics (Figure 55), but it was not significantly different from that of Dicranum polysetum (Figure 42), which had the lowest mass eaten, suggesting that this ratio alone did not account for the preference (Hribljan & Glime in prep). With their unusual digestive tracts (see Digestion above), the terrestrial isopods may be able to gain sufficient nutrition from

Figure 56. Dicranella heteromalla in its typical soil bank habitat. This moss has a relatively low caloric content. Photo by Janice Glime.

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Chapter 10-3: Arthropods: Crustacea – Isopoda, Mysida, and Decapoda

Figure 57. Polytrichastrum alpinum with capsules. Photo by Michael Lüth, with permission.

Several other factors could account for the preferences. First, we know that other deterrents such as saponins, alkaloids, and steroids are present in some mosses and were not tested here (Adebiyi et al. 2012). Leaf structure could make it difficult to obtain energy from the leaves or they might be harder to chew and break off (toughness). We have no measures of such toughness differences for these species, so we must keep an open mind about that possibility. The structure of the cell wall might make it difficult to obtain the cell contents easily (Figure 58-Figure 59). As seen in Figure 58, Pleurozium schreberi has much thinner cell walls than the much less preferred Dicranum polysetum (Figure 42, Figure 59). But does this really translate to toughness? Or edibility? And the leaves might differ from the stems in their phenolic content, making measurements of whole plants meaningless if only leaves are eaten. However, Hribljan and Glime (in prep) used only leaves for their analyses of proteins, carbohydrates, and phenolics. They did compare the chemistry of stems and leaves in Pleurozium schreberi; for all three chemical groups (phenolics, protein, carbohydrates), leaves had the higher content (Figure 60).

Figure 59. Leaf cell structure of Dicranum polysetum showing thick cell wall and low ratio of cell contents to cell wall. This species was least consumed among the five moss species in the study by Hribljan and Glime (in prep.). Photo by Walter Obermayer, with permission.

Figure 60. Comparison of mean phenolic, protein, and carbohydrate content (± 95% CI) of Pleurozium schreberi between freeze-dried leaves and stems (paired t-test, an asterisk indicates a significant difference between the two bars, α = 0.05, n = 3).

These studies leave many questions unanswered, especially regarding season. Do the concentrations in the bryophytes change with season? Do the isopod needs change with season? Does the tree litter change in such a way that bryophytes are preferable at some times and not others without requiring any change in the bryophytes? And are the relationships the same if liverworts are presented instead of mosses? Finally, what evolutionary patterns can we observe and how do they relate to habitat and dominant herbivores?

Figure 58. Leaf cell structure of Pleurozium schreberi showing thin cell wall and high ratio of cell contents to cell wall. This species was most consumed among the five moss species in the study by Hribljan and Glime (in prep.). Photo from Wikimedia Commons.

Defenses and Apparency Theory Plant defenses can be grouped into physical and chemical defenses. Physical defenses include structural modifications into such deterrents as thorns and spines or tissue modifications that include hard cell walls (Cooper & Owen-Smith 1986). The small bryophyte structure does not permit the large thorns found in some tracheophytes, but hard cell walls and hard papillae as extensions of the cell wall do fall into this category.

Chapter 10-3: Arthropods: Crustacea – Isopoda, Mysida, and Decapoda

Chemical defenses can be divided into quantitative and qualitative defenses (Feeney 1975, 1976; Rhoades & Cates 1976; Yamamura & Tsuji 1995). Qualitative defenses include toxic substances like the milky juices of milkweed plants. Few bryophytes have been tested for such substances as those found in the milkweed, but as mentioned above, similar compounds do exist in the few that were tested (Fatoba et al. 2003; Adebiyi et al. 2012). Quantitative defenses, on the other hand, are quite common in bryophytes and typically interfere with digestion (Yamamura & Tsuji 1995), creating malnutrition in the herbivore. Phenolics typically fall in this category. The apparency theory (Feeney 1976) was developed to explain the production of secondary compounds such as phenolics among some plants and not others (Coley et al. 1985). Coley and coworkers contended that resource availability in the environment was a primary determinant of both the amount and type of plant defense. Under resource limitation, slow-growing plants are favored by the environment over fast-growing plants because the former use lower levels of resources. At the same time, slow growth rates favor larger investments in antiherbivore defenses because growth is not fast enough to replace effects of herbivory. Since bryophytes are slow-growing, they are often able to inhabit locations with low levels of resources, including sunlight, where few other plants grow robustly, making the bryophytes one of the obvious, or apparent, plants in the area. Hence, bryophytes could benefit in these situations by the production of antiherbivore compounds. In fact, development of such compounds may have been essential to their success on land as the arthropods likewise became terrestrialized (Graham et al. 2004). Phenolic compounds, occurring in varying concentrations from the bryophytes tested, are useful as antiherbivore compounds. And it appears that bryophytes are not eaten by many kinds of organisms. Isopods are a notable exception to that avoidance. But even they have preferences. We have seen above that for the isopod Porcellio scaber (Figure 1-Figure 2), Dicranum polysetum (Figure 42, Figure 59) is a less-preferred moss compared to Thuidium delicatulum (Figure 50) (Hribljan & Glime in prep). The former is an apparent moss (one with high visibility in its habitat) with high concentrations of secondary compounds (phenolic compounds), whereas Thuidium delicatulum is unapparent (grows with other potential food plants) and is low in secondary compounds (Liao 1993). Furthermore, Thuidium delicatulum tends to grow where there is more sun and often more nutrients, thus supporting the concept that production of phenolic compounds may be related to resource limitation (see Coley et al. 1985). But it is not so simple. Pleurozium schreberi (Figure 43) is a very apparent moss, sometimes covering hectares with 100% cover, yet had the highest consumption. The study by Liao (unpublished) and the discussion here related to the study by Hribljan and Glime (in prep) seem to be the only studies that have tested the apparency theory in bryophytes. This should be an interesting topic for study.

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al. (2007) traced food and fatty acids in macroinvertebrates and determined that the isopods in a stream food web fed on terrestrial food sources and on algae. Asellus species consume a variety of aquatic vegetation. Marcus et al. (1978) experimented with a sometimes moss dweller, Asellus aquaticus (Figure 61), and demonstrated that it ate both Elodea canadensis and periphyton (adhering algae), being able to survive on either. They found fragments of Elodea leaves and pieces of oak (Quercus), as well as the alga Oedogonium in the guts of some individuals of this species from Lake Windermere, England.

Figure 61. Asellus aquaticus, an aquatic isopod that dines on Fontinalis novae-angliae. Photo from Wikimedia Commons.

Parker et al. (2007) found that Asellus aquaticus (Figure 61) consumed large quantities of the brook moss Fontinalis novae-angliae (Figure 62) but rejected the riverweed Podostemum ceratophyllum (Figure 63), despite having similar protein content in both. The isopods continued to eat the F. novae-angliae even when the organic matter was removed from the plants, demonstrating that the moss itself was most likely a food source. They suggested that the mosses served as a refuge against larger predators that could eat the A. aquaticus, largely because such predators as crayfish (Procambarus spiculifer, Figure 64; Figure 95) and Canada geese (Branta canadensis; Figure 65) avoided the mosses despite its comprising 89% of the plant cover in the stream. It seems that the chemical deterrents to the geese and crayfish served to protect the many macroinvertebrates living there. And to the advantage of the A. aquaticus, these isopods rejected the riverweed. On the other hand, this species was not deterred by the chemical defenses of the mosses.

Aquatic Consumers Among the aquatic isopods, some consume bryophytes, but others apparently do not. Torres-Ruiz et

Figure 62. Fontinalis novae-angliae, a habitat and a food source for species of Asellus. Photo by John Parker, with permission.

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Chapter 10-3: Arthropods: Crustacea – Isopoda, Mysida, and Decapoda

F. antipyretica were found in the feces of freshly collected A. cf. militaris (Figure 67), and when A. cf. militaris was cultured in the lab with the moss as a substrate it produced fecal pellets containing the moss. Gut analysis revealed diatoms and detrital matter along with small fragments of Fontinalis (Figure 68). LaCroix found that even terrestrial isopods would eat F. antipyretica.

Figure 63. Podostemum ceratophyllum (riverweed) in upper left, appearing as fine red threads here. This plant has been heavily grazed, whereas the Fontinalis novae-angliae on the right has not. Photo by John Parker, with permission.

Figure 66. Asellus cf. militaris on a branch of Fontinalis antipyretica, where it lives in slow-moving streams and uses the moss as a food source. Photo by Jacob LaCroix, with permission.

Figure 64. Procambarus spiculifer, a crayfish that avoids mosses, thus making the mosses a protected site for the isopods dwelling there. Photo by Chris Lukhaup, with permission.

Figure 67. Asellus cf. militaris feces containing Fontinalis antipyretica and detrital matter. Photo by Jacob LaCroix, with permission.

Figure 65. Branta canadensis (Canada Goose), a large bird that avoids mosses, thus permitting the mosses to protect wouldbe food items that hide there. This one is feeding on riverweed (Podostemum ceratophyllum). Photo by John Parker, with permission.

Asellus cf. militaris (Figure 66) eats Fontinalis antipyretica (Figure 66) in lab experiments and in the field (LaCroix 1996a). Likewise, A. cf. militaris feeds on Fontinalis novae-angliae (Figure 62) in its native aquatic habitat (LaCroix 1996a; Parker et al. 2007). Fragments of

Figure 68. Fontinalis antipyretica and diatoms in gut of Asellus cf. militaris. Photo by Jacob LaCroix, with permission.

Stern and Stern (1969) determined the greatest abundance in February and the lowest in July in a cold springbrook in Putnam County, Tennessee, USA. Asellus

Chapter 10-3: Arthropods: Crustacea – Isopoda, Mysida, and Decapoda

militaris occurs on Fontinalis antipyretica for the first few instars, then moves to the leaf litter. Observations by LaCroix and Glime (unpublished) suggest that this species can live among the mosses for a much greater part of the life cycle in northern Michigan, USA. Like terrestrial isopods, Asellus cf. militaris (Figure 66) avoids the light. Hence, more of these isopods were on the mosses in the shade in the stream than in the sun (LaCroix 1996a; Glime 2006). When both sun and shade mosses were brought to the lab and placed under the same light conditions, the isopods preferred those that had grown in the sun. Furthermore, the isopods chose to go to the mosses collected from the sunny location under both light and dark conditions (LaCroix 1996a). Surprisingly, the shade populations had higher concentrations of phenolic compounds (LaCroix 1996a), a phenomenon contrary to the use of phenolic compounds as light protectants in tracheophytes (Swain & Hillis 1959; Martin & Martin 1982; Mole et al. 1988; Vergeer et al. 1995), but consistent with the preference for those grown in the sun when light was no longer a factor. Bryophytes often take advantage of phenolic compounds as protection against UV radiation (Jorgensen 1994; Clarke & Robinson 2008; Wolf et al. 2010), suggesting that herbivory was a stronger factor in this case than light. This combination of circumstances raises several questions. First, how can we explain isopod preference for high phenolic shade bryophytes in the field but preference for lower phenolic sun bryophytes in the lab (Figure 69)? Parker et al. (2007) showed Asellus aquaticus (Figure 61) was not deterred by extracts from Fontinalis novaeangliae (Figure 62). Parker et al. suggested these isopods have some means to render the deterrent compounds ineffective, as suggested above in the discussion of the digestive system. LaCroix (1996a, b) concluded that food quality of the moss determined what isopods ate, but that shade was a more important determining factor controlling their location (and hence available food) in the field. This combination can structure communities in which small invertebrates live among unpalatable hosts that provide enemy-free space, and isopods have the benefit of avoiding their own predators while being able to eat the substrate.

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Apparency or UV Protection? Having suggested an explanation for the behavior of the isopods, we are left with the question of the higher production of phenolic compounds by the bryophytes in the shade compared to those in the sun. As suggested above for terrestrial bryophytes, it is possible that the production of phenolic compounds by mosses in the shade is an evolutionary response to apparency. In shady locations of streams, mosses are likely to be the dominant macrophyte vegetation, with aquatic tracheophytes preferring sun LaCroix 1996a). As the dominant (most apparent) organism, probability would make the bryophytes the most likely to be eaten. Furthermore, the Fontinalis had phenolic compounds in both locations, so it is likely that they had sufficient levels in the sun to provide the needed protection against UV radiation. Could it be that the Fontinalis produces phenolic compounds in response to herbivory? If so, are they able to signal (chemically) to the nearby mosses to do likewise? Or might this moss have evolved to produce more phenolic compounds in the shade under the selective pressure of one of its primary herbivores, aquatic isopods, that spends most of its time in the shade?

Habitat You know where isopods hang out. Look under anything with a tiny bit of space to give access and you will find them. They go scurrying away in seek of shelter when you lift their cover. But look out at night. They come out in force to eat your vegetables – and your mosses. Bryophytes seem to play multiple roles in the niches of isopods. For terrestrial species, bryophytes provide refuge against some predators, but even for litter-dwelling species they may represent islands for rehydration amid a dry food area. But the bryophytes can also serve as food, especially at night when desiccation is less of a problem. Aquatic bryophytes likewise serve as a refuge against predators and can also serve as food or a food substrate for periphyton and detritus. Zimmer and Topp (1997) found that Porcellio scaber (Figure 1-Figure 2) populations decreased in response to acidification, and that microorganisms, often reduced by acid conditions, were important in the maintenance of juveniles. It seems logical that the first consideration for a habitat for isopods is a moist place with good aeration that provides shelter and darkness, but that also has a food source. In the water, detritus and periphyton can serve as the food source, but on land periphyton is too minor and detritus is more likely to be in the soil. Hence, bryophytes that provide these physical characteristics and are also palatable and chewable become a food source and provide a suitable habitat. Terrestrial

Figure 69. Comparison of moss Fontinalis antipyretica mean phenolic content (± 95% CI) and number of moss-dwelling isopods Asellus cf. militaris (± 95% CI) choosing to inhabit it. Most of the isopods in the lab chose to go to the sun-grown Fontinalis antipyretica that had a lower phenolic content than that in the shade plants. Based on LaCroix 1996b.

Terrestrial habitats require special adaptations for these groups, as discussed above. Edney (1954) found that terrestrialization increased in the order of Ligiidae, Trichoniscidae, Oniscidae, Porcellionidae, to Armadillidiidae. This order can be interpreted as their order for tolerating drought. And each of these families has members known from bryophytes.

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Chapter 10-3: Arthropods: Crustacea – Isopoda, Mysida, and Decapoda

Božanić (2011) sampled the moss invertebrate fauna in a forest in the Vrapač National Nature Reserve, Czech Republic. The most abundant groups were Acarina (mites – 2946 individuals), Collembola (springtails – 1341 individuals), and Isopoda (320 individuals). Within moss colonies on the forest floor and tree trunks they found the isopods Androniscus roseus, Hyloniscus riparius (Figure 86), Hyloniscus spp., Lepidoniscus minutus, Ligidium hypnorum, Porcellium collicola (Figure 3), Porcellium conspersum, Trachelipus rathkii (Figure 26), Trachelipus ratzeburgii (Figure 85), Trachelipus spp., and Trichoniscus pusillus (Figure 25). In the adjoining forest floor, the isopods were not among the most numerous groups sampled. Sample size was important in determining abundance, with more isopods occurring in larger sample sizes of ~400 sq cm. Trichoniscus pusillus and Hyloniscus riparius in particular preferred thicker mosses, especially in Plagiomnium undulatum (Figure 70) with a 50-mm thickness. These two species are known to be hygrophilous (water-loving) (Tajovský 2000), perhaps explaining their preference for thicker mats that could retain moisture longer. This preference could create danger as this thicker moss was also in the range of preference of a predator ant, Myrmica ruginodis (Figure 71), that occurred primarily in mosses having 40-50 mm thickness (Božanić 2011). In poplar forests of Hungary, Hyloniscus riparius (Figure 86) occurs primarily in wet, decaying trees that are covered with mosses (Farkas 1998).

woodlice in the UK (Stenhouse 2007). Its ability to feed on bryophytes is discussed above. Diver (1938) examined the common woodlouse (Porcellio scaber, Figure 24) in five coastal animal successional zones in the British Isles where the plant carpet played a major role in characterizing the habitat. In the Calluna-Psamma zone, there was a well-developed lichen-moss carpet that replaced the grass turf. Nevertheless, only one species of isopod occurred there, whereas two more were added in the Calluna zone where the ground cover was nearly 100% Calluna. In a separate study that compiled many records, Harding and Sutton (1985) reported Trichoniscus pusillus (Figure 25) from all five dune zones, but primarily in dune slacks, where it was associated with mossy areas as well as damp hollows, large pieces of concrete, or decaying wood.

Figure 71. Myrmica ruginodis, an ant that lives among the same mosses as the isopods Trichoniscus pusillus and Hyloniscus riparius, and is a known arthropod predator. Photo by Boris Ginestet and Nicolas Calmejane, through Creative Commons. Figure 70. Plagiomnium undulatum, a moss that forms 50 mm deep mats where the isopods Trichoniscus pusillus (Figure 25) and Hyloniscus riparius (Figure 86) take shelter. Photo by Ralf Wagner , with permission.

Philoscia muscorum (Figure 17), an isopod with a mossy name, is common and widespread in the UK among mosses and other substrata (Stenhouse 2007). Porcellio is perhaps the most common genus in the Northern Hemisphere, occurring with mosses in Europe and North America. Porcellio scaber (Figure 1-Figure 2) is often found among mosses and is one of the commonest of the

Božanić and coworkers (Božanić 2008; Božanić et al. 2013) used heat to extract invertebrates from 61 terrestrial bryophyte samples from forests of the Czech Republic. They found 45 invertebrate species (13 higher taxonomic groups) from among 15 bryophyte species. The moss Brachythecium oedipodium (Figure 72) seems to be a preferred habitat, exhibiting the highest invertebrate diversity on decaying wood, where Isopoda were the most abundant (439 specimens), but diversity was also high in B. salebrosum (Figure 73) (mean 4 spp. per sample) and B. rutabulum (Figure 74) (mean 5.5 spp. per sample).

Chapter 10-3: Arthropods: Crustacea – Isopoda, Mysida, and Decapoda

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Atrichum undulatum (Figure 75), B. rutabulum, and Hypnum cupressiforme (Figure 76) were the most frequent mosses and presented a high number of invertebrate taxa. This abundance is despite the content of hydroxycinnamic and phenolic acids present in B. rutabulum (Davidson et al. 1989).

Figure 75. Atrichum undulatum, mosses where clump size is important in determining isopods (smaller clumps) vs annelids (larger clumps). Photo by Michael Lüth, with permission. Figure 72. Brachythecium oedipodium, a preferred habitat for invertebrates, including Isopoda. Photo by Michael Lüth, with permission.

Figure 73. Brachythecium salebrosum, a bryophyte with a high diversity of invertebrates. Isopods were most abundant in small cushions. Photo by Michael Lüth, with permission.

Figure 74. Brachythecium rutabulum capsules, a moss with high invertebrate diversity, including isopods. Photo by Martin Cooper, through Creative Commons.

Figure 76. Hypnum cupressiforme, a bryophyte with a high diversity of invertebrates. Isopods were most abundant in small cushions. Photo by Michael Lüth, with permission.

Type of substrate, size of cushion, and height above the ground were important determinants of the invertebrate species in these Czech forests (Božanić 2008; Božanić et al. 2013). Isopoda were numerous in small cushions, in contrast to the Enchytraeidae (Annelida) that were abundant in larger moss carpets. The woodlice (isopods) were most abundant among the moss Plagiomnium (Figure 77) on the ground. Tree size also played a role, with isopods Trichoniscus pusillus (Figure 78) and Porcellium collicola (Figure 3) living among mosses on smaller trees, whereas the isopod Trachelipus rathkii (Figure 26) occurred among mosses growing on larger trees. It is possible that correlation with tree diameter resulted from colonization rates and succession of the community. Nevertheless, T. pusillus also occurred among mosses on volcanic rock in the Azores (Vandel 1968). Because the bryophyte habitat was one of the earliest ones available to invasion of land, Božanić and coworkers (2013) suggest that the bryophytes may serve as refugia in expected future climate change.

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Figure 77. Plagiomnium drummondii on rocks in forest, a moss where isopods are abundant. Photo by Janice Glime.

Figure 80. Armadillidium pictum, an isopod that lives under stones and among mosses in the UK. Photo by Jan van Duinen , with permission.

The genus Ligia (Figure 81) is one of the less terrestrialized isopods, requiring more moisture than other terrestrial genera that have been studied, often living in tidal zone cliffs and rocky beaches. But on the Hawaiian Island of Kauai, L. perkinsi commonly occurs among wet mosses of indigenous trees in the montane rainforests above 600 m, whereas on Oahu it is known instead from a windward wet rocky cliff at only 300 m (Taiti et al. 2003).

Figure 78. Trichoniscus pusillus, an isopod that lives among mosses on small trees and among mosses on exposed lava rocks. Photo by Graham Montgomery, with permission.

While pillbugs require moisture, a boggy habitat can be too moist. Although Armadillidium is among the best adapted of isopods to terrestrial life, surviving in relatively dry habitats, some species do use mosses as a habitat. Dale and Dale (1986) report Armadillidium pulchellum (Figure 79) in moss mats of the coastal cliff slopes in the UK. They were surprised to find this species also inland in abundance under mosses on a wall. Harding and Sutton (1985) likewise report them under mats of mosses as well as under stones and mats of the flowering plants Thymus spp. and Sedum anglicum in the UK. In the daytime, one can also find Armadillidium pictum (Figure 80) under stones and among mosses in the UK (Harding & Sutton 1985).

Figure 79. Armadillidium pulchellum, a coastal isopod found among coastal mosses in the UK. Photo by Jan van Duinen , with permission.

Figure 81. Ligia sp., related to the moss dweller Ligia perkinsi that occurs among wet mosses on trees in Hawaiian rainforests. Photo by Steve Nanz, through Creative Commons.

Isopods even live in the exposed higher parts of trees. In the neotropical montane forests of Costa Rica, isopods dwell in both the ground litter and canopy litter, which includes bryophytes (Nadkarni & Longino 1990). But in the montane forests, the isopods had higher densities on the ground. In the Polynesian islands, Philoscia truncata occurs both under stones and among mosses at 500 m on the Society Islands (Jackson 1938). On the Mangareva Islands Spherillo marquesarum occurs under mosses and rocks. In the Tasmanian temperate rainforests, isopods and other invertebrates often occur among mosses in places where they are not common on other substrates (Greenslade 2008). The higher moisture content of the mosses most likely accounts for the higher species richness, with 28 species of isopods among the mosses there. Styloniscus nichollsi is common in Tasmania and can occur among Sphagnum (Figure 83) at 1600 m at Point Lookout (Green

Chapter 10-3: Arthropods: Crustacea – Isopoda, Mysida, and Decapoda

1974). In the Antarctic, several species of Styloniscus occur among mosses: S. otakensis (Figure 88), S. pallidus, S. thompsoni, S. verrucosus (Pugh et al. 2002). Working in the Azores and Madeira, Vandel (1968) found a number of bryophyte-dwelling species not mentioned in other locations cited here, including Trichoniscus pygmaeus among mosses, Miktoniscus chavesi among mosses in a lava field and the bottom of a crater, but also among liverworts in Erica bush, Chaetophiloscia guernei among mosses in the Erica forest and other indigenous vegetation, and Eluma purpurascens among mosses at snowline, under mosses at the roadside of an old lava field, and among mosses in the Erica forest and heath. Androniscus dentiger (Figure 82) occurred on exposed lava rocks covered with mosses and lichens

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Antonović et al. (2012) used pitfall traps to study the isopods living in the Dubravica peat bog and surrounding forest in Croatia. They found eight species of isopods, comprising 389 individuals, during their two-year study, with little difference in species richness between the bog and forest. They considered the small size of the bog peatland, progressive succession of plant life, and interactions among species to account for the high species richness there. Where the grass Molinia spread into the bog, the Sphagnum (Figure 83) was less humid and provided habitats for forest isopod species. The edge (ecotone) had the highest diversity, probably due to multiple factors: greater variety of niches, seasonal immigration, and less predator abundance relative to the open bog. Within the bog, cohabiting lycosid spiders (see Chapter 7-4 on Peatland Spiders) and Myrmica ants (Figure 84) were a threat to the isopods. In the bog Trachelipus rathkii (Figure 26), a known bryophyte dweller, was the most common isopod, whereas in the forest it was Protracheoniscus politus (Figure 85). Bogspecific species were absent. Instead the isopod fauna was dominated by widespread species with wide niche requirements, which Antonović et al. attributed to the degradation process on the bog. Antonović and coworkers considered one bog inhabitant here, Hyloniscus adonis (see Figure 86), to be tyrphoxenous, i.e., a vagrant not reproducing in the bog.

Figure 82. Androniscus dentiger, an inhabitant of mosses and lichens on lava rock in the Azores. Photo by Gilles San Martin, through Creative Commons.

Peatlands Sphagnum (Figure 83) in peatlands often has its own unique fauna, in part due to the unique assemblage of plants. The pH can influence some species. The surface can get quite hot, thus being inhospitable to isopods. But within the peat mats, the gradient of temperature and moisture often provides suitable habitat with the possibility for vertical migration as conditions fluctuate.

Figure 83. Sphagnum cristatum, a moss from boggy habitats where the isopod Trachelipus rathkii (Figure 26) lives in New Zealand. Photo by Jan-Peter Frahm, with permission.

Figure 84. Myrmica sp, an ant predator genus to isopods in bogs. Photo by Alex Wild , with permission.

Figure 85. Protracheoniscus politus (top) and Trachelipus ratzeburgii (bottom), the upper being the most common moss dweller in a forest surrounding a bog in Croatia. Photo by Walter Pfliegler, with permission.

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Chapter 10-3: Arthropods: Crustacea – Isopoda, Mysida, and Decapoda

Figure 86. Hyloniscus riparius, relative of the vagrant isopod Hyloniscus adonis in bogs. Photo by Tom Murray, through Creative Commons.

Springs I expected to find a number of records of isopods among mosses in springs and was surprised to find relatively few. In some of these, although mosses were abundant, the isopods were in the open water and bottom sediments, but not among the mosses (Gooch & Glazier 1991; Erman 2002). Erman (2002) could find no relationship between moss mats and invertebrate diversity, including that of isopods. The only relationship he found was that the mosses indicated that the spring had constancy and persistence. In his study of isopods in habitats of the Azores and Madeira, Vandel (1968) found Trichoniscus pusillus (Figure 25) among mosses in a spring on the mountain slope and among Sphagnum at another spring. But the other spring species were less familiar among moss dwellers, including Miktoniscus chavesi, Chaetophiloscia guernei, and Eluma purpurascens among mosses. Oniscus asellus occurred among mosses in sheltered ravines and under wet moss in the ravine.

Figure 87. Nitellopsis obtusa, an alga that provides habitat for isopods like Asellus but that can disappear in some habitats for part of the year, causing the isopods to seek other shelter. Photo through Public Domain.

Although Asellus aquaticus (Figure 61) is well known from bryophytes, it is the juveniles that are most abundant in algal and bryophyte mats, whereas the larger adults are typically associated with large-sized substratum particles (Graca et al. 1994). As already noted in discussing bryophytes as food, Asellus cf. militaris (Figure 66) occurs in mats of Fontinalis spp. in streams where it feeds on both the mosses and associated detritus and periphyton. On Macquarie Island in the sub-Antarctic, Styloniscus otakensis (Figure 88) lives among mosses on margins of streams, among other places (Greenslade 2008). Cowie and Winterbourn (1979) found that the isopod Styloniscus otakensis was the only common invertebrate on the moss Cratoneuropsis relaxa (Figure 89) in the outer spray zone of a spring brook in the Southern Alps of New Zealand. They attributed differences in fauna among the moss species to differences in flow rates, availability of detritus, and differences in water saturation.

Waterfalls Waterfalls provide a variety of niches from very aquatic to damp terrestrial. These microhabitat niches change as water levels recede and may be quite dry in summer when the waterfall recedes or disappears altogether. Stephensen (1935) found terrestrial Talitridae in such habitats in Java in the Marquesas where Orchestia floresiana occurred among mosses of rivulets, fountains, and waterfalls. Aquatic Aquatic isopods can also be moss inhabitants. Fontaine and Nigh (1983) suggest that aquatic isopods like Asellus (Figure 61) may be limited by their slow colonization rate. When such host plants as Nitellopsis (Figure 87) die off, the isopods need an alternative substrate with sufficient food available (Hargeby 1990). In habitats where bryophytes occur, these bryophytes could provide the permanence needed by the slow isopod colonizers.

Figure 88. Styloniscus otakensis, an aquatic species in a genus with a number of terrestrial moss-dwelling members in forests and bogs of Tasmania, New Zealand, and nearby islands. Photo by Mark Stevens. PERMISSION PENDING.

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Figure 90. Mysis relicta, a species that feeds on diatoms and detritus among mosses in some habitats. Photo by Perhols, through Creative Commons.

CLASS MALACOSTRACA, ORDER DECAPODA

Figure 89. Cratoneuropsis relaxa, genus of mosses that occur in springbrooks in the Southern Alps of New Zealand and home to Styloniscus otakensis. Photo by Tom Thekathyil, with permission.

South Africa may have species unfamiliar to most of us in the Northern Hemisphere. Enckell (1970) found Protojanira prenticei among mosses in the upper part of a streamlet there. Pollution Pollution in the form of heavy metals can quickly move up the food chain in streams. Detrital feeders like Asellus species can concentrate the metals from the detritus on the streambed or among mosses, then get eaten by larger invertebrates or fish, further concentrating the pollutants (Eimers et al. 2001). However, Eimers and coworkers found that when the sediment organic content was increased (20% peatmoss), the cadmium concentration in Asellus racovitzai decreased compared to that of mineral sediment treatments, indicating that bryophytes, especially Sphagnum (Figure 83), might be able to protect the isopods and organisms higher up the food chain by sequestering the heavy metals and keeping them out of the water column. Other mosses, for example Fontinalis antipyretica (Figure 66), occurring in the same waters with Asellus aquaticus (Figure 61), also accumulate heavy metals. Lithner et al. (1995) found that when the pH decreased, the bioconcentration factors decreased in the bryophytes while several of the metals simultaneously increased in fish. Hence, using aquatic bryophytes as bioaccumulators to protect the organisms is complicated, but they could be a useful tool to predict imminent fish dieoff.

Decapods include such animals as crayfish, lobsters, crabs, and hermit crabs. For such large invertebrates to succeed on land they have developed morphological, physiological, biochemical, and behavioral adaptations (Bliss & Mantel 1968). Adult land crabs maintain water balance through the coordinated action of gills, pericardial sacs, and the gut, taking up, storing, and redistributing both salts and water to maintain an osmotic and water balance. In larvae, on the other hand, this suite of responses is not practiced. As is known for the isopods, there is evidence that at least some decapods excrete some of their ammonia as a gas (Weihrauch et al. 2004). Adult land crabs use both gills and the highly vascularized lining of the branchial chambers for gas exchange (Bliss & Mantel 1968). They generally cannot survive low temperatures, but their cytochrome C seems to help in their survival of high temperatures. Finding a mate is typically accomplished by both visual and acoustic signals, coupled with ritualistic behavior. Decapods generally are too large to live among most bryophytes, but they are not without interesting bryological interactions. The decapod Thalassina anomala (Figure 91-Figure 93), a mud lobster, forms soil mounds (Figure 92-Figure 93) when it builds its nest (Yamaguchi et al. 1987). It is on these soil mounds in the mangrove forests of Japan that Fissidens microcladus dwells. By living on the soil mounds, the moss is never submerged at high tide and most likely benefits from the moist air.

CLASS MALACOSTRACA, ORDER MYSIDA The Mysida are known as oppossum shrimps because of the brood pouch where females carry their larvae. Mysids are not common on bryophytes, but they can use them as a restaurant in aquatic habitats. Mysis relicta (Figure 90) in Char Lake, Northwest Territories, Canada, feeds primarily on diatoms and inorganic particles on moss substrata (Lasenby & Langford 1973). It is known as an opportunistic feeder, permitting it to survive on a variety of resources (Grossnickle 1982).

Figure 91. Thalassina anomala, a mud lobster that makes mounds in mangrove forests – mounds that have somewhat unique flora including Fissidens microcladus. Photo by Ariff Aziz, through Creative Commons.

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Chapter 10-3: Arthropods: Crustacea – Isopoda, Mysida, and Decapoda

Figure 92. Mound of the mud lobster, Thalassina anomala, in a mangrove forest. Photo by Ariff Aziz, through Creative Commons.

The relationship of the Parenephrops species with stream mosses contrasts with the avoidance of mosses by the crayfish Procambarus spiculifer (Figure 64, Figure 95; see also discussion under Isopoda – Aquatic Consumers) reported by Parker et al. (2007). The latter crayfish is selective in its plant habitat, choosing the flowering plant Podostemum ceratophyllum (riverweed; Figure 96) over Fontinalis novae-angliae (Figure 62; Figure 96), despite the greater abundance of the moss (89% of total biomass) (Parker et al. 2007). Furthermore, the mosses supported twice as many macroinvertebrates as did the riverweed. This revelation suggests that the mosses might provide a safe refuge for macroinvertebrates, allowing them to escape from larger predators, perhaps due to their chemical defenses. This hypothesis is supported by the presence in the moss of C18 acetylenic acid, octadeca-9,12-dien-6-ynoic acid, a defense compound that inhibits crayfish feeding. A similar avoidance was absent in the amphipods and isopods in the stream, permitting them to find safe refuge there. This discriminatory behavior of the antifeedant against crayfish but not microcrustacea permits these small arthropods to live where they can avoid the predation of larger arthropods.

Figure 93. Close view of a mound of the mud lobster, Thalassina anomala, showing greenish patches that could be protonemata of the moss Fissidens microcladus. Photo by Ariff Aziz, through Creative Commons.

Coffey and Clayton (1988) have suggested that deep water bryophytes in New Zealand lakes do not occur in the presence of freshwater crayfish. It appears that in the presence of the crayfish Paranephrops spp. (Figure 94), the bryophytes suffer both mechanical damage and browsing. In Lake Wanaka, there is a deep water (down to 50 m) community of bryophytes (Coffey & Clayton 1988). But in other New Zealand lakes the mosses were absent. This absence correlated with the presence of large crayfish (Paranephrops spp.) populations. Coffey and Clayton suggest that the mosses are absent not due to different habitat needs from the crayfish, but from the browsing and mechanical damage caused by the crayfish.

Figure 94. Paranephrops planifrons, member of a genus of crayfish that inflicts mechanical damage on bryophytes. Photo by David Wilson, through Creative Commons.

Figure 95. Procambarus spiculifer eating Egeria densa. This crayfish avoids eating the moss Fontinalis novae-angliae, thus protecting its invertebrates as well. Photo by John Parker, with permission.

Figure 96. Podostemum ceratophyllum (left) and Fontinalis novae-angliae (right) showing effects of grazing by the crayfish Procambarus spiculifer on the P. ceratophyllum. The moss remains untouched. Photo by John Parker, with permission.

Chapter 10-3: Arthropods: Crustacea – Isopoda, Mysida, and Decapoda

Summary Isopods include a number of terrestrial genera, many of which include bryophyte dwellers, including the families Ligiidae, Trichoniscidae, Oniscidae, Porcellionidae, and Armadillidiidae. Asellus seems to be the most common genus in streams. Springs seem to have few isopods inhabiting mosses. Other taxa benefit from the moisture of bogs, migrating vertically to achieve optimum moisture and temperature. As descendents of aquatic and marine organisms, isopods benefit from the moisture and protection of bryophytes, finding food among them as detritus, periphyton, and the bryophytes themselves. Their digestive system is modified by reducing gut surface tension and culturing gut flora to render the phenolic compounds safe in their diet. They are known to eat a wide range of bryophytes, but they do have preferences, and some taxa are ignored. In addition to sheltering, the isopods use the bryophytes as a place to remove excess water or gain needed water. They conserve water by releasing their nitrogenous waste as ammonia gas. Isopods are sensitive to temperature, and bryophytes can provide shade and evaporative cooling. Isopods often go into the soil in the daytime, emerging and climbing to the tips of the bryophytes to dine at night. They congregate under bryophytes, as well as rocks, logs, and boards, reducing water loss and oxygen consumption, stimulating reproduction, increasing predator defense, promoting coprophagy, and acquiring internal symbionts. Reproduction is typically sexual, but parthenogenesis is possible in some taxa. The eggs and young are carried by the mother. Some isopods overwinter under bryophytes or in the soil under bryophytes. They generally cannot survive temperatures below -7°C. At least some bryophytes exemplify the apparency theory. The bryophytes are small and slow-growing. They contain a wide range of antiherbivore compounds that deter most herbivores. Isopods, on the other hand, circumvent the antiherbivore compounds through their digestive system, permitting them to gain a food source (bryophytes) where they are protected from a number of would-be predators. However, ants are a predatory threat even among the bryophytes. Members of the order Mysida are rarely reported from bryophytes, but in Char Lake they feed on diatoms and inorganic particles among mosses. The Decapoda (crayfish) generally do not live among mosses, in some cases actually avoiding them, apparently due to the presence of C18 acetylenic acid, octadeca-9,12-dien-6-ynoic acid in the mosses (and possibly other compounds). Others damage the bryophytes by moving their heavy bodies across them. Invertebrates are able to avoid predation by crayfish by living among the mosses.

Acknowledgments Thank you to all the people who have contributed images or posted them through Creative Commons. Much

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of the content of this chapter presents results from undergraduate and graduate research by my students, who were a constant source of inspiration to me. Eileen Dumire reviewed the chapter from the perspective of a novice and Gipo Montesanto provided a scientific review. John Parker provided the images to complete the story of predatory Canada Geese and crayfish that avoid the mosses. John Hribljan provided many discussions, did much of the research on Porcellio scaber, and commented on the chapter.

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Fatoba, P. O. Omojasola, P. F., Awe, S., and Ahmed, F. G. 2003. Phytochemical screening of some selected tropical African mosses. Nigerian Soc. Exper. Biol. J. 3(2): 49-52. Feeny, P. 1975. Biochemical coevolution between plants and their insect herbivores. In: Gilbert, L. E. and Raven, P. H. (eds.). Coevolution of Animals and Plants. University of Texas Press, Austin and London, pp. 3-19. Feeny, P. 1976. Plant apparency and chemical defense. In: Wallace, J. and Mansell, R. A. (eds.). Biochemical Interaction between Plants and Insects. Recent Advances in Phytochemistry, Vol. 10. Plenum Press, New York, pp. 140. Fontaine, T. D. and Nigh, D. G. 1983. Characteristics of epiphyte communities on natural and artificial submersed lotic plants: Substrate effects. Arch. Hydrobiol. 96: 293301. Forman, R. T. T. 1968. Caloric values of bryophytes. Bryologist. 71: 344-347. Fussey, G. D. 1984. The distribution of the two forms of the woodlouse Trichoniscus pusillus Brandt (Isopoda: Oniscoidea) in the British Isles: A reassessment of geographic parthenogenesis. Biol. J. Linn. Soc. 22: 309-321. Fussey, G. D. and Sutton, S. L. 1981. The identification and distribution of the bisexual and parthenogenetic forms of Trichoniscus pusillus (Isopoda: Oniscoidea) in Ireland. Irish Nat. J. 20: 196-199. Ganter, P. F. 1984. The effects of crowding on terrestrial isopods. Ecology 65: 438-445. Gay Brereton, J. Le. 1957. The distribution of woodland isopods. Oikos 8: 85-106. Glime, J. M. 2006. Bryophytes and herbivory. Cryptog. Bryol. 27: 191-203. Gooch, J. L., and Glazier, D. S. 1991. Temporal and spatial patterns in mid-Appalachian springs. Mem. Entomol. Soc. Canada 123: 29-49. Graca, M. A. S., Maltby, L., and Calow, P. 1994. Comparative ecology of Gammarus pulex (L.) and Asellus aquaticus (L.) I: Population dynamics and microdistribution. Hydrobiologia 281: 155-162. Graham, L. E., Kodner, R. B., Fisher, M. M., Graham, J. M., Wilcox, L. W., Hackney, J. M., Obst, J., Bilkey, P. C., Hanson, D. T., and Cook, M. E. 2004. Early land plant adaptations to terrestrial stress: A focus on phenolics. The Evolution of Plant Physiology. Elsevier Academic Press, Boston, pp. 155-169. Green, M. A. J. 1974. Oniscoidea (terrestrial Isopoda). In: Biogeography and Ecology in Tasmania, Monographiae Biologicae Vol. 25. Springer, Netherlands, pp. 229-249. Greenaway, P. 1985. Calcium balance and moulting in the Crustacea. Biol. Rev. 60: 425-454. Greenslade, P. 2008. Distribution patterns and diversity of invertebrates of temperate rainforests in Tasmania with a focus on Pauropoda. Mem. Museum Victoria 65: 153-164. Grossnickle, N. E. 1982. Feeding habits of Mysis relicta – an overview. Hydrobiologia 93: 101-107. Hames, C. A. C. and Hopkin, S. P. 1989. The structure and function of the digestive system of terrestrial isopods. J. Zool. 217: 599-627. Hansen, C. E. and Rossi, P. 1991. Effects of culture conditions on accumulation of arachidonic and eicosapentaenoic acids in cultured cells of Rhytidiadelphus squarrosus and Eurhynchium striatum. Phytochemistry 30: 1837-1841. Harding, P. T. and Sutton, S. L. 1985. Woodlice in Britain and Ireland: Distribution and habitat. Institute of Terrestrial Ecology. Lavenham Press, Huntingdon, UK, 151 pp.

Chapter 10-3: Arthropods: Crustacea – Isopoda, Mysida, and Decapoda

Hargeby, A. 1990. Macrophyte associated invertebrates and the effect of habitat permanence. Oikos 57: 338-346. Hartenstein, R. 1968. Nitrogen metabolism in the terrestrial isopod, Oniscus asellus. Amer. Zool. 8: 507-519. Hassall, M. and Rushton, S. P. 1982. The role of coprophagy in the feeding strategies of terrestrial isopods. Oecologia 53: 374-381. Hassall, M. and Tuck, J. M. 2007. Sheltering behavior of terrestrial isopods in grasslands. Invert. Biol. 126: 46-56. Hassall, M., Edwards, D. P., Carmenta, R., Derhé, M. A., and Moss, A. 2010. Predicting the effect of climate change on aggregation behaviour in four species of terrestrial isopods. Behaviour 147: 151-164. Hornung, E. and Warburg, M. R. 1993. Breeding patterns in the oniscid isopod, Porcellio ficulneus Verh., at high temperature and under different photophases. Invert. Repro. Devel. 23: 151-158. Horowitz, M. 1970. The water balance of the terrestrial isopod Porcellio scaber. Entomol. Exper. Appl 13: 173-178. Hribljan, J. A. 2009. The Influence of Moss and Litter Chemical Traits on Bryophagy in a Northern Temperate Forest Invertebrate, Porcellio scaber Latr. M.S. Thesis, Michigan Technological University, Houghton, MI, USA, 73 pp. Jackson, H. G. 1938. Terrestrial isopods of southeastern Polynesia. Occ. Papers Bernice P. Bishop Museum 14: 167192. Jorgensen, R. 1994. The genetic origins of biosynthesis and light-responsive control of the chemical UV screen of land plants. In: Ellis, B. E., Kuroki, G. W., and Stafford, H. A. (eds.). Genetic Engineering of Plant Secondary Metabolism. Plenum Press, N.Y., pp. 179-192. Kautz, G., Zimmer, M., and Topp, W. 2000. Responses of the parthenogenetic isopod, Trichoniscus pusillus (Isopoda: Oniscidea), to changes in food quality. Pedobiologia 44: 7585. Kight, S. L. and Nevo, M. 2004. Female terrestrial isopods, Porcellio laevis Latreille (Isopoda: Oniscidea) reduce brooding duration and fecundity in response to physical stress. J. Kans. Entomol. Soc. 77: 285-287. Kight, S. L. and Ozga, M. 2001. Costs of reproduction in the terrestrial isopod Porcellio laevis Latreille (Isopoda: Oniscidea): Brood-bearing and locomotion. J. Kans. Entomol. Soc. 74: 166-171. LaCroix, J. 1996a. Food and light preferences of Asellus. Bull. N. Amer. Benthol. Soc. (abstr.) 13(1): 121. LaCroix, J. J. 1996b. Phenolics from Fontinalis antipyretica Hedw. and light as causes of differential distribution of Asellus militaris Hay in Gooseneck Creek. Unpublished M.S. Thesis, Mich. Tech. Univ., Houghton, MI, 47 pp. Lardies, M. A. and Bozinovic, F. 2008. Genetic variation for plasticity in physiological and life-history traits among populations of an invasive species, the terrestrial isopod Porcellio laevis. Evol. Ecol. Res. 10: 747-762. Lardies, M. A., Cotoras, I. S., and Bozinovic, F. 2004. The energetics of reproduction and parental care in the terrestrial isopod Porcellio laevis. J. Insect Physiol. 50: 1127-1135. Lasenby, D. C. and Langford, R. R. 1973. Feeding and assimilation of Mysis relicta. Limnol. Oceanogr. 18: 280285. Lavy, D., Rijn, M. J. Van, Zoomer, H. R., and Verhoef, H. A. 2001. Dietary effects on growth, reproduction, body composition and stress resistance in the terrestrial isopods Oniscus asellus and Porcellio scaber. Physiol. Entomol. 26: 18-25.

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Liao, C.-L. 1993. Chemical defence in bryophytes with high apparency. In: Glime, J. M. Ecology Column, The Bryological Times 75: 1-4. Lindqvist, O. V and Fitzgerald, G. 1976. Osmotic interrelationship between blood and gut fluid in the isopod Porcellio scaber Latr. (Crustacea). Compar. Biochem. Physiol. A Physiol. 53: 57-59. Lithner, G., Holm, K., and Borg, H. 1995. Bioconcentration factors for metals in humic waters at different pH in the Roennskaer area (N. Sweden). In: Grennfelt, P., Rodhe, H., Thoerneloef, E., and Wisniewski, J. (eds.). Acid Reign '95? Proceedings from the 5th International Conference on Acidic Deposition: Science and Policy, held in Goteborg, Sweden, 26-30 June 1995. Water Air Soil Pollut. 85: 785-790. Marcus, J. H., Sutcliffe, D. W., and Willoughby, L. G. 1978. Feeding and growth of Asellus aquaticus (Isopoda) on food items from the littoral of Windermere, including green leaves of Elodea canadensis. Freshwat. Biol. 8: 505-519. Massey University. 2014. Guide to New Zealand Soil Invertebrates. Isopoda. Accessed 6 April 2014 at . Merriam, H. G. 1971. Sensitivity of terrestrial isopod populations (Armadillidium) to food quality differences. Can. J. Zool. 49: 667-674. Mole, S., Ross, J. A., and Waterman, P. G. 1988. Light-induced variation in phenolic levels in foliage of rain-forest plants. J. Chem. Ecol. 14: 1-21. Nadkarni, N. M. and Longino, J. T. 1990. Invertebrates in canopy and ground organic matter in a Neotropical montane forest, Costa Rica. Biotropica 22: 286-289. Nair, G. A. 1998. Reproductive and population biology of Porcellio scaber (Isopoda, Oniscidea) in Benghazi, Libya. Israel J. Zool. 44: 399-412. Nyirő, G., Oravecz, O., and Márialigeti, K. 2002. Detection of Wolbachia pipientis infection in arthropods in Hungary. Eur. J. Soil Biol. 38: 63-66. Pakarinen, P. and Vitt, D. H. 1974. The major organic components and caloric contents of high Arctic bryophytes. Can. J. Bot. 52: 1151-1161. Parker, J. D., Burkeile, D. E., Collins, D. O., Kubanek, J., and Hay, M. E. 2007. Stream mosses as chemically-defended refugia for freshwater macroinvertebrates. Oikos 116: 302312. Prins, H. H. 1982. Why are mosses eaten in cold environments only? Oikos 38: 374-380. Pugh, P. J. A., Dartnall, H. J. G., and McInnes, S. J. 2002. The non-marine Crustacea of Antarctica and the Islands of the Southern Ocean: Biodiversity and biogeography. J. Nat. Hist. 36: 1047-1103. Rhoades, D. F. and Cates, R. G. 1976. Toward a general theory of plant antiherbivore chemistry. In: Wallace, J. W. and Nansel, R. L. (eds.). Biological Interactions Between Plants and Insects. Recent Advances in Phytochemistry 10. Plenum Press, New York, pp. 169-213. Rushton, S. P. and Hassall, M. 1983a. Food and feeding rates of the terrestrial isopod Armadillidium vulgare (Latreille). Oecologia 57: 415-419. Rushton, S. P. and Hassall, M. 1983b. The effects of food quality on the life history parameters of the terrestrial isopod (Armadillidium vulgare (Latreille)). Oecologia 57: 257-261. Samouelle, G. 1819. The Entomologist's Calendar, exhibiting the time of appearance and habitation of near three thousand species of British insects. In: The Entomologist's Useful Compendium; An Introduction to the Knowledge of British Insects. R. and A. Taylor, Shoe-lane, 496 pp.

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Schotte, M., Boyko, C. B, Bruce, N. L., Poore, G. C. B., Taiti, S., Wilson, G. D. F. (eds.). 2008-2014. World List of Marine, Freshwater and Terrestrial Isopod Crustaceans. Accessed 28 May 2014 at . Smith, R. M., Young, M. R., and Marquiss, M. 2001. Bryophyte use by an insect herbivore: Does the crane-fly Tipula montana select food to maximise growth? Ecol. Entomol. 26: 83-90. Stenhouse, D. 2007. Appendix G5, Entomologial Survey Report. Accessed 18 March 2012 at . Stephensen, K. 1935. Terrestrial Talitridae from the Marquesas. Bernice P. Bishop Museum Bull. 142: 19-34. Stern, M. S. and Stern, D. H. 1969. A limnological study of a Tennessee cold springbrook. Amer. Midl. Nat. 82: 62-82. Sveinbjörnsson, B. and Oechel, W. C. 1991. Carbohydrate and lipid levels in two Polytrichum moss species growing on the Alaskan tundra. Holarctic Ecol. 14: 272-277. Swain, T. and Hillis, W. E. 1959. The phenolic constituents of Prunus domesticus. I. The quantitative analysis of phenolic constituents. J. Sci. Food Agric. 10: 63-68. Taiti, S., Arnedo, M. A., Lew, S. E., and Roderick, G. K. 2003. Evolution of terrestriality in Hawaiian species of the genus Ligia (Isopoda, Oniscidea). Koninklijke Brill NV, Leiden, pp. 85-102. Tajovský, K. 2000. Mnohonožky (Diplopoda), stonožky (Chilopoda) a suchozemští stejnonožci (Oniscidae) vybraných aluviálních ekosystémů střední a severní Moravy (Litovelské Pomoraví a Poodří). In: Kovařík, P. and Machar, I. (eds.). Mokřady 2000. Sborník z konference při příležitosti 10. výročí vzniku CHKO Litovelské Pomoraví. Správa CHKO ČR a Český Ramsarský výbor, Praha, pp. 230-232. Tanaka, K. and Udagawa, T. 1993. Cold adaptation of the terrestrial isopod, Porcellio scaber, to subnivean environments. J. Compar. Physiol. B 163: 439-444. Torres-Ruiz, M., Wehr, J. D., and Perrone, A. A. 2007. Trophic relations in a stream food web: Importance of fatty acids for macroinvertebrate consumers. J. N. Amer. Benthol. Soc. 26: 509-522. Vandel, A. 1968. The terrestrial Isopoda of the Azores. Report No. 52. Lund University Expedition in 1957 to the Azores and Madeira.

Vergeer, L. H. T., Aarts, T. L., and Groot, J. D. De. 1995. The 'wasting disease' and the effect of abiotic factors (light intensity, temperature, salinity) and infection with Labyrinthula zosterae on the phenolic content of Zostera marina shoots. Aquat. Bot. 52: 35-44. Warburg, M. R. 1987. Isopods and their Terrestrial Environment. Academic Press, New York. Weihrauch, D., Morris, S., and Towle, D. W. 2004. Ammonia excretion in aquatic and terrestrial crabs. J. Exper. Biol. 207: 4491-4504. Weston, M. 1995. The effects of phenolic and protein contents in Polytrichum commune and P. juniperinum on isopod feeding behavior. Unpublished report, Department of Biological Sciences, Michigan Technological University, Houghton, MI. Wieser, W. and Schweizer, G. 1970. A re-examination of the excretion of nitrogen by terrestrial isopods. J. Exper. Biol. 52: 267-274. Wolf, L., Rizzini, L., Stracke, R., Ulm, R., and Rensing, S. A. 2010. The molecular and physiological responses of Physcomitrella patens to ultraviolet-B radiation. Plant Physiol. 153: 1123-1134. Yamaguchi, T., Nakagoshi, N., and Nehira, K. 1987. Terrestrial bryophytes in mangrove forests in Japan. Proc. Bryol. Soc. Japan 4: 137-140. Yamamura, N. and Tsuji, N. 1995. Optimal strategy of plant antiherbivore defense: Implications for apparency and resource-availability theories. Ecol. Res. 10: 19-30. Zimmer, M. 1997. Surfactants in the gut fluids of Porcellio scaber (Isopoda: Oniscidea), and their interactions with phenolics. J. Insect Physiol. 43: 1009-1014. Zimmer, M. 1999. The fate and effects of ingested hydrolyzable tannins in Porcellio scaber. J. Chem. Ecol. 25: 611-628. Zimmer, M. and Brune, A. 2005. Physiological properties of the gut lumen of terrestrial isopods (Isopoda: Oniscidea): Adaptive to digesting lignocellulose? J. Comp. Physiol. B 175: 275-283. Zimmer, M. and Topp, W. 1997. Does leaf litter quality influence population parameters of the common woodlouse, Porcellio scaber (Crustacea: Isopoda)? Biol. Fert. Soils 24: 435-441.

Glime, J. M. 2017. Aquatic insects: Biology. Chapt. 11-1. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. Ebook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 19 July 2020 and available at .

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CHAPTER 11-1 AQUATIC INSECTS: BIOLOGY TABLE OF CONTENTS Aquatic Insects .................................................................................................................................................. 11-1-2 Life Cycle Stages .............................................................................................................................................. 11-1-3 Collembola ................................................................................................................................................. 11-1-3 Hemimetabolous Insects ............................................................................................................................ 11-1-4 Nymphs ............................................................................................................................................... 11-1-4 Naiads ................................................................................................................................................. 11-1-4 Holometabolous Insects ............................................................................................................................. 11-1-4 Adaptations to Aquatic Bryophyte Life ............................................................................................................ 11-1-5 Life Cycle Strategies .................................................................................................................................. 11-1-5 Life Cycle Cues .................................................................................................................................. 11-1-8 Temperature Relations ...................................................................................................................... 11-1-10 Overwintering ................................................................................................................................... 11-1-10 Structural .................................................................................................................................................. 11-1-12 Attachment ............................................................................................................................................... 11-1-14 Behavioral ................................................................................................................................................ 11-1-17 Oxygen Conditions .................................................................................................................................. 11-1-18 Obtaining Food ........................................................................................................................................ 11-1-21 Who Lives There? ........................................................................................................................................... 11-1-22 Specificity ................................................................................................................................................ 11-1-27 Seasons..................................................................................................................................................... 11-1-30 Sampling ......................................................................................................................................................... 11-1-33 Preservative .............................................................................................................................................. 11-1-34 Extraction ................................................................................................................................................. 11-1-35 Flotation ................................................................................................................................................... 11-1-35 Artificial Mosses ............................................................................................................................................. 11-1-36 Summary ......................................................................................................................................................... 11-1-36 Acknowledgments ........................................................................................................................................... 11-1-37 Literature Cited ............................................................................................................................................... 11-1-37

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CHAPTER 11-1 AQUATIC INSECTS: BIOLOGY

Figure 1. Rhyacophila carolina larva, a free-living caddisfly that occurs commonly on bryophytes. Photo by Bob Henricks, with permission.

Aquatic Insects Cascading waterfalls, silt-laden torrents, lurking predators, limited oxygen, unpredictable water levels, icy winters – all these dangers face the insects (Figure 1) that call lakes, and especially streams, their homes. So why do the insects choose to live there, and how do bryophytes help to make life in such unfriendly conditions possible? The relationship between aquatic insects and bryophytes is a topic dear to my heart. When I was working on my M.S. project on the bryophytes, my roommate was working on aquatic insects. Never passing up an opportunity for a field trip, I accompanied her on all her collecting trips. We both soon realized that in her rocky mountain streams of northern West Virginia, USA, there were typically more insects among the bryophytes than in any other microhabitat in these streams. It was this discovery that led me to my Ph. D. research topic on the insects associated with Appalachian stream bryophytes and the many studies I have done on ecology of aquatic mosses since then. These wonderful bryophyte-insect communities are not a new discovery. Stream ecologists in particular have observed the importance of mosses as cover for aquatic insects and other aquatic invertebrates and even fish (Thienemann 1912; Carpenter 1927; Percival & Whitehead 1929, 1930; Humphries & Frost 1937; Jones 1941, 1948, 1951; Frost 1942; Badcock 1949; Illies 1952; Hynes 1961; Minckley 1963; Egglishaw 1969; Arnold & Macan 1969; Lindgaard et al. 1975; Hawkins 1984; McKenzie-Smith 1987; Suren & Winterbourn 1992a, b; Gislason et al. 2001; Linhart et al. 2002; Paavola 2003).

In Idaho, USA, Maurer and Brusven (1983) found that Fontinalis neomexicana (Figure 2) housed 5-30x the densities of insects found associated with the mineral substrates; biomass, however, was only 2x as great. The moss did not alter insect densities in the underlying hyporheic zone (saturated zone beneath the bed of a river or stream that can support invertebrate fauna). The diversity of functional groups was greater among mosses, but the species richness was similar to that of the mineral substrate.

Figure 2. Fontinalis neomexicana, a moss that greatly increases the density of stream insects. Photo by Belinda Lo, through Creative Commons.

Chapter 11-1: Aquatic Insects: Biology

The numbers of insects among bryophytes can be extensive (Figure 3). Minckley (1963) found that mosses had the highest densities of insects compared to sand, stones, and tracheophytes in a Kentucky, USA, stream. Lillehammer (1966) found that moss-covered stones had 606 individuals m-2 compared to 471 m-2 on stones with no mosses.

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When more sophisticated statistical methods became available, bryophyte biomass emerged as one of the factors accounting for the variation in insect fauna among streams, and as we might expect, it has a positive influence on the insect fauna (Gislason et al. 2001). Furthermore, bryophytes can occupy deeper waters, forming a zone that is lower than that of tracheophytes, and this zone is able to support fauna that could not otherwise live at those depths (Blackstock et al. 1993). Minshall (1984) considered bryophytes to be a major factor in increasing insect numbers because of the increased surface area offered by them. Egglishaw (1969) found that most species of invertebrates, including insects, were less aggregated in clumps among the mosses than they were under stones. One might interpret that this is due to the complex nature of the mosses and the large space in which they can be distributed. On the other hand, it would seem that the stone habitat would be more homogeneous and thus one might expect less clumping. Another mystery.

Figure 3. This branch of Palustriella commutata demonstrates the variety and density of aquatic insects that can occur on aquatic mosses. Photo by Dan Spitale.

Table 1. Orders of insects and their abundances among bryophytes in various locations around the world. NR refers to not recorded, which may mean the researcher(s) didn't look at the group. Collembola Odonata Diptera Coleoptera sample size Ephemeroptera Plecoptera Trichoptera

Straffan, River Liffey, Ireland 200 g Ballysmuttan, River Liffey, Ireland 200 g Cold Springbrook, TN, USA 0.1 m2 Bystřice, Czech Republic 10 g dry Mlýnský náhon, Czech Republic 10 g dry Welsh Dee Tributary, Wales ~300 cm2 Mouse Stream, Alpine, NZ 1 m2 Tim's Creek, Alpine, NZ 1 m2 West Riding, Yorkshire, UK – loose moss % West Riding, Yorkshire, UK – thick moss % alpine unshaded stream, NZ % alpine shaded stream, NZ % River Sawdde, Wales

NR NR NR NR NR NR NR NR NR NR NR NR rare

533 16 7.1 1103 176 9.7 NR NR 13.42 8.03 NR NR NR

NR NR NR NR NR NR NR NR NR NR NR NR NR

Life Cycle Stages Life cycle stages play a major role in the occupancy of water habitats by insects. Most of these orders of insects have poor ability to survive freezing, so escape into water can maintain their temperatures above freezing. The flowing part of water generally remains at ~1°C throughout the winter, and lakes and ponds that don't freeze to the bottom have water just above 0 up to 4°C. Because of the importance of water in the life cycle of the major groups of aquatic insects, we must understand the types of life cycles among them before we can begin a discussion of the biology and ecology of these groups. There are two major groups of classification among the insects, based on life cycles and their developmental stages.

22 310 8 18 0 513 540 270 154 0.65 2.1 2.5 very rare

11446 10482 215 44762 11035 82.8 61270 24580 65.3 42 58l.8 69.9 NR

492 148 24.6 359 13 0.4 730 260 3.1 8 NR NR NR

262 1095 0.4 184 5 7.4 0 90 6.7 4.4 NR NR very rare

Reference

Frost 1942 Frost 1942 Stern & Stern 1969 Vlčková et al. 2001-2002 Vlčková et al. 2001-2002 Hynes 1961 Suren 1991a Suren 1991a Percival & Whitehead 1929 Percival & Whitehead 1929 Suren 1991b Suren 1991b Jones 1949

Collembola The Collembola (Figure 4), or springtails, long considered to be insects, have been kicked out of the Insecta by cladistics, due to linkages shown by their DNA and supported by their morphology. Because they have much of their ecology in common with insects, and their earlier inclusion among Insecta, they will be discussed among these aquatic insect subchapters. The Collembola have the simplest life cycle, one in which the hatchling is a miniature of the adult. The immature stage is known as a nymph. Their life cycle consists of egg/embryo, nymph, and adult. The Collembola hatch from their egg casing and look like the adults, perhaps in somewhat different proportions; they

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continue to increase in size without changing their basic form as they become adults.

Figure 4. Collembola Arthropleona oruarangi, a group of "pre-insects" that are born looking like little adults. Photo by Stephen Moore, Landcare Research, NZ, with permission.

creature, with or without legs, or in some cases with prolegs that are of soft tissues. The aquatic larvae have gills in many taxa, but not in others. Some have fleshy legs with hooks at the posterior end.

Figure 5. Plecoptera exuvia. Photo by Jason Neuswanger at , with permission.

Hemimetabolous Insects (Hemimetabola) Nymphs Among the aquatic insects, this group includes the true bugs (order Hemiptera), a group that lacks gills in all stages. The Hemiptera are hemimetabolous insects and thus lack the pupal stage (familiar to most people as the chrysalis of butterflies). Instead, they have only the egg/embryo (Figure 8), nymph (including naiads in the other hemimetabolous orders), and adult. [The holometabolous insects, on the other hand, have an egg/embryo, larva, pupa, and adult (imago).] Naiads Those orders with obligate aquatic immature stages that do not resemble the adults, but that do not pass through a second stage as a pupa before becoming an adult, have an aquatic stage known as a naiad. The naiad is a specialized nymph stage known only among aquatic insects and occurs in the orders Plecoptera (Figure 5, Figure 73, Figure 74, Figure 77), Ephemeroptera (Figure 6), and Odonata (Figure 7). The naiad usually differs from the adult in having some form of gills to aid in gaining oxygen in the aquatic environment. When it is time for the adult to emerge, these insects climb to the surface or out of the water, often on an emergent plant, and often hang vertically while they climb out of their naiad exoskeleton (Figure 5). The shed exoskeleton is the exuvia (pl. exuviae; Figure 5). In the Ephemeroptera, the emergent stage is a subadult known as a subimago (Figure 6). This subimago goes through one additional moult to become the adult (imago). Holometabolous Insects (Holometabola) The remaining orders of aquatic insects are holometabolous and have what is known as complete metamorphosis. These insects have four life cycle stages: egg/embryo (Figure 8), larva (Figure 1), pupa (Figure 9), adult (imago; Figure 10). The larva stage looks nothing like the adult. It is familiar to most people in the moths and butterflies as the caterpillar. The larva is a worm-like

Figure 6. Baetis male subimago emerging to adult. Photo by Jason Neuswanger at , with permission.

Figure 7. Enallagma damselfly naiad. Murray, through Creative Commons.

Photo by Tom

The pupa is usually a stationary phase (known as a chrysalis in butterflies). As the pupa develops, the larva develops a chitinous outer covering that has the imprint of parts like wings and antennae. The insect is likely to be dormant or in diapause (in insects, period of suspended development, especially during unfavorable environmental

Chapter 11-1: Aquatic Insects: Biology

conditions) during its pupal stage, providing it reprieve from winter's cold or tropical drought. But during this time the insect goes through a number of changes in both form and physiology. When the insect has matured into an adult and conditions are right for its emergence, it breaks out of the pupa. In most cases, those that spend their larval lives in the water emerge into the atmosphere, spending their adult lives as terrestrial organisms (except in most of the beetles).

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The holometabolous insect orders that live among bryophytes include Coleoptera, Trichoptera, Megaloptera, Neuroptera, and Diptera.

Adaptations to Aquatic Bryophyte Life Bryophyte dwellers might benefit from several behavioral and structural adaptations to make life among the bryophytes easier. They need to be able to gain sufficient oxygen (Hynes 1970), to move about freely, to avoid being pulled out if a predator catches a tail or leg, to avoid being swept away by the current, and to eat the available food. In streams where the water level varies a lot or dries up, they need to have a means to avoid desiccation. Life Cycle Strategies

Figure 8. Emerald dragonfly with eggs. Armstrong, with permission.

Photo by Bob

Figure 9. Chironomidae (midge) pupa. Henricks, with permission.

Photo by Bob

Figure 10. Chironomidae adult male. Photo by Roger S. Key, with permission.

Although I would normally discuss structural adaptations first, the life cycle adaptations appear to be the most important ones among the insects. Differing requirements among life cycle stages permit insects to survive from year to year in changing environmental conditions. Blackstock et al. (1993) found the insects in a clear sequence of bryophyte to herbaceous swamp to woody plant community occupying different depth zones in the basin of Pant-y-llyn, Wales. These changes, on a large scale, require a degree of mobility on the part of the insect inhabitants as the habitat changes from aquatic to terrestrial seasonally. But even more permanent aquatic habitats have their down times. Success for an aquatic insect means having a strategy to survive during stages when the habitat is dry (Blackstock et al. 1993), too cold, or too hot. To understand the role of bryophytes in the life of their insect inhabitants, one must understand these life cycles. Only twelve orders of insects plus the Collembola (Figure 4) are generally considered to have aquatic members, but even these aquatic members typically live out of the water during part of their lives (Thorp & Covich 1991; Ward 1992). Since most of the aquatic insects live in the water in immature stages, an understanding of these stages is necessary to understand fully how bryophytes are so important for them. Danks (1991) points out that we can understand insect life cycle adaptations best by understanding the options. These include the choices (evolutionarily) to develop or to enter diapause (period of suspended development) and to grow rapidly or grow slowly. These developmental options respond to photoperiod and temperature, among other things (Danks 1991; Zwick 1996). Because of dependency on these cues, eggs of some stoneflies are able to remain in the sediments for years, providing a "seed bank" (Zwick 1996). The choices that have been programmed into the life cycle impact the life span of the insect. Eggs (Figure 8) are an important stage for insects with a terrestrial adult stage and aquatic immature stage(s). The term egg is used somewhat loosely, referring to both the unfertilized egg and the embryonic stage that remains within the egg "shell," indicated herein as egg/embryo. Most of these insects lay their eggs in the water, so a substrate that anchors and protects them from both flowing water and predation is important. Even such freeswimming insects as the dragonfly Sympetrum (Figure 11) in the Odonata sometimes lay their eggs in plates on moss

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growths, securing the eggs and hatchlings (Figure 12) (Wesenberg-Lund 1943).

development, long or repeated dormancy, or adults that live a long time (Danks 1991, 1992). Others, in particular the mayflies (Ephemeroptera), may live for only one day as adults (Figure 13), just long enough to mate and lay eggs, but can spend about one year in the naiad stage in the water. The long life cycles are usually coupled with several factors, including cold, unpredictable temperatures, unreliable or low quality food supplies, natural enemies, and large adult size. Life cycle traits relate strongly to the predictability of the environment where the insect lives and the environmental signals that are provided (Danks 2006). But Danks cautions that much more detail is needed to understand these life cycle patterns in insects.

Figure 13. Callibaetis ferrugineus subimago. Jason Neuswanger, with permission. Figure 11. Sympetrum sanguineum mating. Photo by Qartl through Creative Commons.

Photo by

Radford and Hartland-Rowe (1971) examined the life cycles of stream insects from Alberta, Canada. Several of these represent genera [Nemoura/Zapada/Prostoia (Figure 14), Ephemerella/Drunella (Figure 15)] that are common among bryophytes. Of these, Prostoia (=Nemoura) besametsa (see Figure 16) and Drunella (=Ephemerella) coloradensis (Figure 17) are characterized as fast seasonal types. But in the same family, Zapada (=Nemoura) cinctipes (Figure 18), Z. columbiana (Figure 19), Z. oregonensis (Figure 20-Figure 21), and Drunella doddsii (Figure 22) are slow seasonal types. None of these species has more than one brood per year except Zapada cinctipes, which has two. Temperature is important in determining growth rate in these species.

Figure 12. Sympetrum striolatum egg-laying among grasses and mosses. Photo by Hugh Venables through Creative Commons.

Some of the aquatic insects live in immature stages in the water for more than one year (Danks 1992; Ulfstrand 1968b). These extended lives may result from slow

Figure 14. Nemoura sp. naiad, a genus with both fast and slow development. Photo by Bob Henricks, with permission.

Chapter 11-1: Aquatic Insects: Biology

Figure 18. Zapada cinctipes naiad. Armstrong, with permission.

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Photo by Bob

Figure 15. Ephemerella invaria naiad, a genus with both fast and slow development. Photo by Bob Henricks, with permission.

Figure 19. Zapada columbiana adult on snow, emerging in winter. Photo by Jason Neuswanger, with permission.

Figure 16. Prostoia naiad, a common bryophyte dweller. Photo by Jason Neuswanger, with permission.

Figure 20. Zapada oregonensis naiad showing gills. Photo by Jim Moore, through Creative Commons.

Figure 17. Drunella coloradensis naiad, having a fast seasonal type of development. Photo by Bob Henricks, with permission.

Figure 21. Zapada oregonensis adult. Photo by Jim Moore, through Creative Commons.

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Chapter 11-1: Aquatic Insects: Biology

Figure 23. Diamesa (Diptera) pupal exuvium, a genus that may produce 8-10 generations in a single year. Photo by Will Bouchard, with permission.

Figure 22. Drunella doddsii naiad, having a slow seasonal type of development. Photo by Bob Henricks, with permission.

Since insects have little tolerance for low temperatures (Dunman et al. 1991; Moore & Lee 1991), they must spend winter in a way that avoids the dangers of freezing (Ramløv 2000), as will be discussed in more detail below. It is this need to avoid freezing that forces some insects to spend part of their lives in the water. Bryophytes provide a habitat that helps them to cope with this watery habitat. As Danks (1991) points out, the life cycle strategies provide options that facilitate survival: develop or enter diapause; grow rapidly or grow slowly. These are typically under the control of such environmental parameters as temperature and photoperiod. Life Cycle Cues As already stressed, changes in life cycle phases are often necessary to survive changing weather conditions as the seasons change. Danks (1999) pointed out that life cycles are influenced by climate severity, seasonality, unpredictability, and variability. Some insects solve the unpredictability and variability problems by having flexible life cycles. These modifications can be determined by factors such as food availability and temperature. Danks (1991) points out that various stages in the life cycle are used in combination to adapt the insects to the changes of the seasons in nature. In cold environments, some of the Chironomidae (Diamesa incallida; Figure 23) may produce 8-10 generations in a single year, with egg-laying occurring throughout the year (Nolte & Hoffmann 1992). Diamesa incallida is a hot-spring-dwelling midge that lives in water at 76-80°C, a community where we are not likely to find bryophytes, but it demonstrates the role of temperature and the wide range of capabilities in a family that is common among bryophytes. Some Arctic Chironomidae solve the problem of finding a sexually mature mate by negating the need for mating and being parthenogenetic (producing offspring without fertilization) (Langton 1998).

Shama and Robinson (2009) demonstrated that an alpine caddisfly (Allogamus uncatus, a bryophyte dweller) in Switzerland responded to late season photoperiod cues by accelerating development, but the species showed adaptive plasticity in response to season length, making responses different among populations with only small geographic differences. Furthermore, the responses of the two sexes can differ (Shama & Robinson 2006). On the other hand, the bryophyte-dwelling caddisfly Limnephilus externus (Figure 24-Figure 26) did not make developmental adjustments in response to diet supplementation, although it did grow to a larger size (Jannot et al. 2008). Furthermore, this caddisfly was unable to adjust to pond drying, responding by reduced growth rates and delayed development. This indicates the danger of an unpredictable environment for the aquatic insects.

Figure 24. Limnephilus externus larva in case. Photo by Wendy Brown , with permission.

Figure 25. Limnephilus externus adult, a caddisfly that does not adjust its development in response to food supplements. Photo by Jason Neuswanger, with permission.

Chapter 11-1: Aquatic Insects: Biology

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temperature was more important than number of days for development, with 34 instars being produced in the laboratory at 20°C. That number is most likely plastic in response to environmental conditions.

Figure 26. Limnephilus externus larva showing abdominal gills. Photo by Wendy Brown , with permission.

In unpredictable or unstable environments, flexibility in the life cycle is important (Brittain & Saltveit 1989). Knispel et al. (2006) found that the bryophyte-dwelling mayfly Baetis alpinus (Figure 27) in the Swiss floodplains has synchronous egg development with high hatching success. By developing faster in warmer habitats it is able to hatch when conditions are favorable in the autumn. Long development time and delayed hatching permit success in unpredictable habitats in the cold glacial conditions. The mayfly Rhithrogena nivata (see Figure 28) has a long incubation period; the timing of hatching and glacial discharge conditions determine the success of development. This plasticity permits it to live in the very unstable, cold habitats that are limiting to other species.

Figure 28. Rhithrogena impersonata naiad, a genus in which some species have life cycle plasticity that depends on local weather. Photo by Donald S. Chandler, with permission.

Figure 29. Leptophlebia cupida naiad, a species with only one reproductive cycle per year. Photo by Jason Neuswanger, with permission.

Figure 27. Baetis alpinus naiad, a mayfly with synchronous egg development that promotes high hatching success. Photo by Andrea Mogliotti, with permission.

Many insects have developmental cues similar to those of plants. These include degree-days (calculated by taking the average of the daily maximum and minimum temperatures compared to a base temperature necessary for growth by the species). As in many plants, degree days may be important in determining the rate of development. For example, the mayfly Leptophlebia cupida (Figure 29) in the Bigoray River, Alberta, Canada, has only one reproductive period each year (Clifford et al. 1979). Clifford et al. (1979) found that degree days of water

For aquatic insects, the temperatures are much more tempered than in the terrestrial environment. In a study of 95 aquatic species, Pritchard et al. (1996) found that only 4 of 92 possible comparisons among congenerics (members of same genus) demonstrated significant differences in degree of cold adaptation. All Odonata (damselflies and dragonflies), 71% of Diptera (true flies), and 81% of Ephemeroptera (mayflies) had significant slopes indicating that they were warm adapted. They suggested that the Plecoptera are cold-adapted species that may use the egg stage to survive when the temperatures are too high. In the stonefly family Leuctridae, commonly represented among bryophytes, the length of the naiad stage depends on the temperature. In Leuctra ferruginea (Figure 30) those individuals living in the coolest streams required two years for their life cycle, whereas those in the warmest waters were able to complete the life cycle in one

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Chapter 11-1: Aquatic Insects: Biology

year (Harper 1973). The Leuctridae (Figure 30) and Nemouridae (Figure 14) are both common at the cooler upstream stations in Southern Ontario. Six species of the stonefly Isogenoides (Figure 31) from Colorado, USA, a genus also known from mosses, varied in hatching time both among the species and within some species (Sandberg & Stewart 2004). In one species the eggs hatched over an extended period of time, stopped hatching for the winter, then resumed hatching in May-June the following year. Some eggs even survived and hatched two years later. In one species, a summer diapause was needed before the eggs would hatch. Members of the genus required three months to four years before hatching.

Figure 30. Leuctra ferruginea naiad, a stonefly that has modified its life cycle to suit the climatic conditions. Photo by Tom Murray at BugGuide.

Figure 31. Isogenoides hansoni naiad, in a genus with moss-dwelling members in which life cycles vary both between and within species. Photo by Bob Henricks, with permission.

Temperature Relations As already noted, temperature plays an important role in determining when life cycle stages occur. Freezing, desiccation, and anoxia are all lethal among aquatic insects, from egg to adult (Lencioni 2004). When in the aquatic habitat, these three factors are related, with ice preventing the renewal of oxygen, and ice crystals drawing water from the cells, causing desiccation. Some of the aquatic insects enter diapause during winter. This usually requires storage of food as glycogen and lipids, hormonal control, and depression or suppression of oxidative metabolism with mitochondrial degradation. But the mosses themselves seem to present a relatively constant temperature. Thorup (1963) considered the

temperature among mosses in springs to be so constant that it would not provide the developmental temperature point needed to trigger changes in stages. Correlated with the moss habitats in springs was an insect life cycle with only one generation per year. Overwintering Duman et al. (1991) defined two physiological mechanisms by which insects survive winter: freeze tolerance and freeze avoidance or freeze resistance (see also Ramløv 2000). Aquatic insects have only limited ability to survive at temperatures below freezing (Moore & Lee 1991). They can supercool to only -3 to -7°C and only some members in the order Diptera are known to be freeze tolerant. The adults seem to be somewhat more cold tolerant. Thus this is a group of insects for which aquatic habitats that do not freeze provide them with an escape to suitable temperatures for the winter. What is fascinating is the plasticity of their responses. Duman et al. (1991) found that not only do different populations of the same species exhibit different overwintering mechanisms, but that even the same population may change its overwintering mechanism from year to year. Because of their need for warmer temperatures in immature stages than that needed by terrestrial insects, most of the aquatic insects spend their egg and immature stages in the water. In fact, warm-water insects avoid the freezing dangers of winter by surviving as eggs. This is particularly true for the blackflies (Simuliidae; Figure 51Figure 53) (Hynes 1970). Insects rarely spend their entire lives in the water, but some spend larval stages there, pupal stages on land, then return to the water as adults, as in many Coleoptera (beetles). Others, particularly some of the Trichoptera (caddisflies) overwinter as adults. In fact, some even emerge mid-winter in cold climates. And the adult stonefly Zapada cinctipes (Nemouridae; Figure 18) re-enters the water when air temperatures drop below freezing (Tozer 1979). However, the stream chironomid Diamesa mendotae (Diptera; Figure 32-Figure 33) does things quite differently – its freeze tolerance is actually greater in the larval (stream) stage (Figure 33). Although it has a larval super-cooling-point (SCP) temperature of -7.4°C and pupal SCP of -9.1°C, compared to -19.7°C for the adults (Bouchard et al. 2006), the larvae of D. mendotae are freeze tolerant, with a lower lethal temperature (99% dead) of -25.4°C, ~10°C lower than their minimum super cooling point (-15.6°C). They change from freeze tolerant as larvae to freeze intolerant as adults! Nevertheless, the adults are able to tolerate cold temperatures sufficiently to mate on the snow (Ferrington et al. 2010). Furthermore, they can survive under the snow for extended periods of time (Anderson et al. 2013). The often moss-dwelling Serratella ignita (Figure 60) overwinters from late summer until late the next spring as an egg (Arnold & Macan 1969). On the other hand, the mayfly Ameletus inopinatus (Figure 34) and stonefly Leuctra hippopus (Figure 35), a stony bottom dweller, do the most developing in the naiad stage while their stream is iced over, at least in northern Sweden (Ulfstrand 1968b). The low temperatures slow, but usually do not stop, development and growth.

Chapter 11-1: Aquatic Insects: Biology

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Figure 32. Diamesa mendotae adult on snow. Permission to reproduce given by Leonard Ferrington on behalf of the Chironomidae Research Group at the University of Minnesota. Figure 35. Leuctra hippopus, a stonefly that develops in Sweden while the stream is iced over. Photo by Niels Sloth, with permission.

Figure 33. Diamesa mendotae larvae alive in Petri dish after freezing. Permission to reproduce given by Leonard Ferrington on behalf of the Chironomidae Research Group at the University of Minnesota.

But ice is also a good insulator, so those insects living on the bottom of lakes and ponds are usually able to avoid lethal low temperatures there. Such insects as the Chironomidae (Figure 90) typically live in sediments where oxygen content is low. Cold water holds more oxygen, and since these organisms are adapted to low oxygen conditions, there is sufficient oxygen in the cold water. Some Chironomidae and Trichoptera (Figure 83) actually occur in ice and frozen sediment, as noted in a north Swedish river (Olsson 1981). Olsson found that 80100% of these frozen insects survived thawing. Chironomidae survived exposure to -4°C for five months. Danks and Oliver (1972a) found that in the Arctic Chironomidae that overwinter are mature larvae and are ready to emerge as soon as the winter season is over. They take advantage of the warm sun by emerging in the middle of the day when the water temperature is highest (Danks & Oliver 1972b). It is interesting that Plecoptera, Ephemeroptera, Trichoptera, Diptera, and Coleoptera have all been recovered alive from anchor ice (submerged ice anchored to the bottom; Figure 36). Anchor ice can encase bryophytes as well, and when it breaks loose, it can take the entire patch of bryophytes with it. Hence, it would likewise take all the insect inhabitants as well, moving them downstream to a new location.

Figure 34. Ameletus ludens naiad, member of a genus where some species develop under the ice in streams. Photo by André Wagner, with permission.

It is interesting that in alpine streams that have snow cover for 6-9 months of the year, taxa richness and abundance of the insects seems to have no seasonal pattern. Nevertheless, the species composition differs significantly from summer to winter. Schütz et al. (2001) found two strategies for larval survival. The insects either had to be adapted to the extreme conditions of summer or avoid these by developing during the winter (typical of Ephemeroptera and Plecoptera).

Figure 36. Anchor ice, Alberta, Canada, visible here as cloud-like mounds of ice attached to the rocks under water. Courtesy of Pacific Northwest National Laboratory.

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Chapter 11-1: Aquatic Insects: Biology

Structural Hynes (1970) summarized the adaptations of stream insects to include flattening, streamlining, friction discs, close application to the surface of stones, and in some the presence of hydraulic suckers. But many of these adaptations pertain to a life on rocks or other relatively smooth substrate. Such characters as flattening, friction discs, close application to the surface, and hydraulic suckers are of little value among the chambers of a bryophyte mat. This leaves us with only one adaptation from his list, that of streamlining (Figure 37), present in the stoneflies [Plecoptera: Leuctridae (Figure 37), Capniidae (Figure 38), Chloroperlidae (Figure 39), and some Gripopterygidae (Figure 40)], and mayflies [Ephemeroptera: Leptophlebiidae (Figure 41) and Baetidae (Figure 45)] – all known from bryophytes. Others have retained the dorsi-ventral flattening, but it is better described as compressing (Figure 42) since these insects do not quite fit the definition of flat. And compression is useful among bryophytes. Other bryophyte adaptations include small size, attachment hooks, and gill covers or gills absent (Glime 1968).

Figure 39. Chloroperlidae naiad. Photo by Bob Henricks, with permission.

Figure 37. Leuctra laura naiad showing streamlining. Photo by Tom Murray at BugGuide, through Creative Commons. Figure 40. Zelandobius illiesi, a stonefly naiad with streamlining. Photo by Stephen Moore, Landcare Research NZ, with permission.

Figure 38. Allocapnia sp. naiad showing streamlining. Photo by Bob Henricks, with permission.

Figure 41. Paraleptophlebia mollis naiad, a mayfly illustrating streamlining. Photo by Tom Murray through Creative Commons.

Chapter 11-1: Aquatic Insects: Biology

Figure 42. Ephemerella naiad showing dorsi-ventral compression. Photo by Bob Henricks, with permission.

Bryophyte-dwelling insects therefore do not necessarily have the same adaptations as stream insects in general. Streamlining helps, but does not need to be as severe. Steinmann (1907, in Muttkowski 1929) found that about 30% of the bryophyte-dwelling taxa were streamlined. But in the streams of the Appalachian Mountains, streamlining was not common (Glime 1994). For example, the common bryophyte-dwelling mayfly Ephemerella (Figure 42) is neither flattened nor streamlined (Arnold & Macan 1969), but has a shape more like a terrestrial insect – it is dorsiventrally compressed. Small size is also an advantage and seems to be the most important characteristic of bryophyte dwellers. Bryophytes provide small spaces where invertebrates can hide, but these same small spaces limit the sizes of the organisms that can occur there. This explains why bryophytes tend to harbor small species and hatchling insects (Figure 43).

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Figure 44. Leuctra inermis adult, a species whose early naiad instars live among mosses in riffles. Photo by James K. Lindsey, with permission.

Frost (1942) remarked that because of the very young and thus small specimens, identification was both difficult and questionable, forcing identification to genus or subfamily only. Glime (1994) found that Baetis sp. was present among mosses (10 per gram) in summer, but were absent in later stages when the larger naiads were present among rocks in the stream bed. Others that moved out of the bryophytes when they got larger were the cranefly Limonia (Figure 47), stonefly Taeniopteryx (Figure 48), and caddisflies Lepidostoma (Figure 49) and Neophylax (Figure 50). Similar migration of older stages occurs in Europe (Thienemann 1912; Carpenter 1927; Egglishaw 1969).

Figure 45. Baetis rhodani, a mayfly that starts its life among bryophytes, but moves out as it grows larger. Photo by J. C. Schou through Creative Commons.

Figure 43. Taeniopteryx naiad on the edge of a Syracuse watch glass, demonstrating the small size of this bryophyte dweller. Photo by Bob Henricks, with permission.

Dudley (1988) suggested that while the complex structure of bryophytes might interfere with attachment by larger larvae, it reduces frequency of encounter between such predators and the small insect inhabitants. In the Appalachian, USA, streams 70% of the bryophyte dwellers were less than 6 mm long (Glime 1994). Egglishaw (1969) found that a higher proportion of smaller animals occurred on mosses than on stones of riffles. In Leuctra inermis (see Figure 37, Figure 44), Baetis rhodani (Figure 45), and Isoperla grammatica (Figure 46) it was the young (small) stages that occurred among the bryophytes.

Figure 46. Isoperla grammatica naiad showing dorsiventral compression. Photo by Dragiša Savić, with permission.

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Chapter 11-1: Aquatic Insects: Biology

Figure 47. Limonia sp., an insect that lives among bryophytes until it gets too large; then it moves out. Photo by Stephen Moore, Landcare Research, NZ, with permission. Figure 50. Neophylax atlanta larva and case, a caddisfly that moves from bryophytes to other substrates as it grows. Photo by Bob Henricks, with permission.

Figure 48. Taeniopteryx sp. naiad, a moss-dwelling stonefly that moves to substrates with more space when it gets larger. Photo by Bob Henricks, with permission.

Figure 49. Lepidostoma larva and case, a caddisfly that moves out of the bryophytes as it grows. Photo by Bob Henricks, with permission.

Attachment While torrents bring much-needed oxygen, they also are treacherous, dislodging the insects and sweeping them downstream. Black flies (Simuliidae; Figure 51-Figure 53) are among the best adapted of the aquatic insects for surviving this torrential onslaught, living on the upper surface of the bryophyte mats (Niesiolowski 1979). On both rocks and mosses, they are able to anchor themselves with a circle of hooks on the rear of the abdomen (Figure 51) (Arnold & Macan 1969). Furthermore, they manufacture a silken thread that they lay down on their substrate surface as an anchor. When they do become dislodged by chance or choice, they have a tether that prevents them from travelling too far and helps them to gain a "foothold" on their new downstream substrate. Those hooks, on both the abdomen and the single proleg foot (Figure 52), enable blackfly larvae to grab onto the silken mat (Figure 53) they have made. They are able to use these same two sets of hooks to move along their silken mat like inch worms.

Figure 51. Simuliidae larva showing anal hooks. Photo by Bob Henricks, with permission.

Chapter 11-1: Aquatic Insects: Biology

Figure 52. Prosimulium mixtum larva showing single proleg. Photo by Tom Murray at BugGuide, through Creative Commons.

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Figure 55. Rhyacophila fuscula larva showing anal hooks that serve as anchors. Photo by Jason Neuswanger, with permission.

Figure 53. Simuliidae larvae on leaf where silken threads form a mat, aiding in attachment. Photo by Bob Henricks, with permission.

The net-spinning caddisflies (Hydropsychidae) accomplish anchorage by a pair of hooks on the posterior end (Figure 54), a modification of many caddisflies for pulling themselves into their cases. But among the freeliving caddisflies like the Hydropsychidae and Rhyacophilidae [e.g. Rhyacophila dorsalis (Badcock 1949)], these hooks (Figure 55) serve as anchors among the bryophytes. Other insects have hooked claws that help them to clamber among the bryophytes, including the beetles (e.g. Elmidae, Figure 56) and some mayflies (e.g. Ephemerellidae, Figure 60) and stoneflies [e.g. Nemoura (Figure 57) and Acroneuria (Figure 58)]. Others, like the Chironomidae, achieve anchorage by nestling at the leaf bases (Figure 59) where little flow occurs.

Figure 54. Hydropsyche larva showing posterior prolegs with hooks that provide anchorage. Photo by Bob Henricks, with permission.

Figure 56. Elmidae adult showing clawed feet that help it climb among mosses. Photo by Stephen Moore, Landcare Research, NZ, with permission.

Figure 57. Nemoura sp. naiad showing hooked claws. Photo by Bob Henricks, with permission.

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Figure 58. Acroneuria abnormis naiad showing hooked claws. Photo by Tom Murray, through Creative Commons.

Figure 61. Zelandobius illiesi (Gripopterygidae) showing backward-pointing dorsal spines. Photo by Stephen Moore at Landcare Research, NZ, with permission.

Figure 59. Rheotanytarsus exiguus (Chironomidae) group nestled in leaf bases. This species makes a tube where it lives. Photo by D. N. Bennett, with permission.

Hora (1930) and Ward (1992) suggested that backward pointing dorsal spines (Figure 60-Figure 64) of some moss dwellers, e.g. the Gripopterygidae (Figure 61), are adaptations to reduce chances of being swept downstream. Illies (1961) reported large dorsal spines on a mossdwelling stonefly from Chile. Similar (but smaller) spines are known on the common moss-dwelling mayfly Ephemerella ignita (Figure 60; Hynes 1970). Even Diptera larvae [e.g. Psychodidae (Figure 62), Tipulidae (Figure 63-Figure 64)] can have backward-directed spines. But the tipulid larvae of Phalacrocera (Figure 63) and Triogma (Figure 64-Figure 65) have such projections and live mostly among semiaquatic mosses where there is no flow to dislodge them. This suggests the spines may serve either as camouflage or as trapping devices to prevent would-be predators from pulling them out of the moss mat.

Figure 60. Serratella ignita naiad showing spinelike structures on the dorsal side of the abdomen. Photo by J. C. Schou through Creative Commons.

Figure 62. Clogmia albipunctata (Psychodidae) larva with backward pointing spines. Photo by Ashley Bradford through Creative Commons.

Figure 63. Phalacrocera replicata larva showing green color and projections that help to camouflage it among mosses. Photo from Wikimedia Commons.

Chapter 11-1: Aquatic Insects: Biology

Figure 64. Triogma larva showing backward pointing spines. This larva also has cryptic coloration that makes it difficult to detect among the bryophytes. Photo by Janice Glime.

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Figure 67. Serratella gills showing gill covers and fibrillate gills on successive abdominal segments. Photo by Bob Henricks, with permission.

Figure 65. Triogma trisulcata larva among Sphagnum showing appendages that mimic moss leaves. Photo by Walter Pfliegler, with permission.

Gill covers help to keep silt from accumulating among the gills, since the mosses often reside where they collect large amounts of silt. The gill covers can also be used to fan the fills, hence moving the water and facilitating oxygen exchange. Gill covers are common among the Ephemeroptera, especially in the Ephemerellidae (Figure 66-Figure 67) and Caenidae (Figure 68).

Figure 66. Drunella grandis naiad showing raised gill covers and fimbrillate gills. Photo by Bob Newell, with permission.

Figure 68. Caenis latipennis naiad showing large gill covers over the dorsal abdomen. Photo by Donald S. Chandler, with permission.

Behavioral Behavior often permits organisms to change their locations, providing the best location available to them as the season changes and required resources are in new locations. Behavioral adaptations can help them capture prey, avoid being prey themselves, gain sufficient oxygen, avoid being swept away by the current, and escape cool or freezing temperatures. Bryophytes provide a series of zones (Figure 69) that permit insects to live in the flow regime they require. As will be seen, oxygen can be a limiting factor, requiring some insects to live near the surface of the bryophyte where torrential waters trap oxygen from the air. Hence, these insects require a means of anchorage lest they themselves become part of the torrent. Others are well adapted to the low oxygen levels and live at the base where detritus accumulates and predators seldom venture. But it is advantageous that they can move about and seek the zone within the stream or lake and within the bryophyte community that best meets their needs.

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Chapter 11-1: Aquatic Insects: Biology

Figure 69. Fontinalis zonation of insects. Redrawn from Niesiolowski 1979.

Aquatic insects tend to avoid light, exhibiting negative phototaxis (Moon 1940; Shelford 1945). Mayflies, in particular, demonstrate a negative phototaxis, preferring darker locations (Wodsedalek 1911; Gros 1923; Percival & Whitehead 1926). This may account for the presence of some taxa among the darker spaces of mosses, particularly in rapid water where rock surfaces may be highly exposed to light. Others may avoid light to be less conspicuous to their prey. On the other hand, Baetis harrisoni (Figure 70) chose illuminated stones 112 times compared to 14 for shaded stones, exhibiting strong positive phototaxis (Hughs 1966).

Figure 70. Baetis harrisoni naiad, a mayfly that prefers illuminated stones. Photo by Helen James through Creative Commons.

Insects often escape adverse conditions in their environments by modifying the environments themselves. Such modifications may include making shelters (Figure 71), excavating, aggregating (Figure 53), forming colonies, and parental actions (Danks 2002). Although all of these actions may be found among aquatic insects, not all of these occur among those living among bryophytes. The bryophyte itself sometimes makes such actions as excavating and making shelters unnecessary. For example, several families of caseless caddisflies live among bryophytes. But the very tiny Hydroptilidae may take advantage of the bryophytes for case-building materials.

Figure 71. Helicopsyche case, made by the caddisfly as a shelter. Photo by Mike Quinn, through Creative Commons.

Oxygen Conditions Ponds can become quite anoxic in winter when the surface is frozen (Nagell & Brittain 1977). Streams are less likely to become anoxic, but within the bryophyte mat water can be quite quiet and oxygen can be used up quickly by decaying organisms. However, insects have a wide array of adaptations to help them through places and times of anoxia (Hoback & Stanley 2001). For example, 10 Arctic species of Collembola (springtails) are known to survive anoxia at 5°C for up to 36 days (Hodkinson & Bird 2004). The mayfly Cloeon dipterum (Figure 72) is able to survive 3-4 months in anoxic ponds, and naiads survived up to 155 days at 0°C in the lab (Nagell 1977).

Figure 72. Cloeon dipterum, a mayfly that can survive 3-4 months in anoxic pond water. Photo by Malcolm Storey, through Creative Commons.

As is obvious from previous studies, oxygen relations in the insects are dependent on temperature (Jacob & Walther 1981). More oxygen can dissolve at low temperatures. In fact, oxygen limitations due to temperature are so important that they set the thermal limits in at least some species of aquatic insects (Verberk & Bilton 2011). Furthermore, since smaller insects use less oxygen, large insects may have been an adaptation to excess oxygen in the Carboniferous Era (Verberk & Bilton 2011). Oxygen limitations may explain in part the presence of small insects among the bryophytes, whereas

Chapter 11-1: Aquatic Insects: Biology

the larger stages move to rock faces where flow is uninterrupted and able to replenish the oxygen more easily. Knight and Gaufin (1966) measured oxygen consumption as a function of temperature in two stonefly naiads that associate with bryophytes: Hesperoperla pacifica (Figure 73) and Pteronarcys californica (Figure 74). These insects followed the general trend of consuming more oxygen at higher temperatures. This relationship is problematic because gasses are lost from the water at higher temperatures, thus limiting the most available oxygen to winter.

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Insects living in low oxygen conditions may be adapted by developing enlarged respiratory organs (Figure 75) (Dodds & Hisaw 1924), including enlargement of tracheal gills (Figure 76) (Golubkov et al. 1992). Behavior can play an important role, with most species moving away from the anoxic sediments when oxygen becomes limiting (Kolar & Rahel 1993). But moving is not always a good choice because it can result in being swept into the current and usually means becoming more visible, hence being more obvious to predators. The movement itself attracts attention through the excellent vision in the well developed eyes of other arthropods and fish.

Figure 73. Hesperoperla pacifica with its pompom-like gills peeking out from the ventral thorax. Photo by Arlen Thomason, with permission.

Figure 75. Relationship of gill size in seven species of Ephemeroptera to oxygen availability in aquatic systems. The outlier species on the right is the genus Iron, a genus for which the gills form a suction cup, preventing one side of the gills from functioning in oxygen uptake. Its position when only half the area is used is shown by the square at the base of the dotted line on the right. Redrawn from Dodds & Hisaw 1924.

Figure 74. Pteronarcys californica, probably the largest insect inhabitant of bryophytes. Photo by Bob Henricks, with permission.

Among the common bryophyte dwellers, the mayflies (Ephemeroptera) are the least tolerant of low oxygen (Gaufin et al. 1974), making them good indicator organisms. These are followed by stoneflies (Plecoptera), then caddisflies (Trichoptera), flies (Diptera), and damselflies (Odonata) in that order. Of course there are exceptions within the orders.

Gills are a common adaptation to low oxygen, especially in Ephemeroptera (Figure 76), Plecoptera (Figure 77-Figure 79), and Trichoptera (Figure 80). These are placed in almost every position (e.g. Figure 78), depending on the genus or family, and are useful taxonomic characters in some groups. But they also tend to be protected, between legs or under gill covers. Others have cutaneous breathing – providing the expanse of the insect's surface and avoiding the danger of collecting sediments.

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Chapter 11-1: Aquatic Insects: Biology

Figure 79. Acroneuria carolinensis naiad showing gills on the ventral thorax. Photo by Tom Murray, through Creative Commons.

Figure 76. Leptophlebia nebulosa showing abdominal (tracheal) gills. Photo by Don S. Chandler, with permission.

Figure 80. Hydropsyche sp. larva showing gills on ventral side. Photo by Bob Henricks, with permission.

Figure 77. Nemoura sp. naiad showing clusters of white thoracic gills at the "neck." Photo by Bob Henricks, with permission.

Figure 78. Coxal gills on a winter stonefly. Photo by Bob Henricks, with permission.

As early as 1907, Babak and Foustka concluded that as the oxygen concentration in the water decreased, movement of the gills of mayflies increased. Dodds and Hisaw (1924) showed a relationship between gill area and oxygen concentration in mayflies. But in the mayfly Baetis (Figure 45, Figure 70) used for testing, the gills never beat and it seems that they do not use their gills for oxygen consumption in the range of 5.0 to 8.0 cc L-1 (Wingfield 1939). Rather, these mayflies live in rapid streams where oxygen concentrations are usually above 4 cc L-1 and rapid flow keeps fresh, oxygenated water flowing over the gills. Under these conditions their cuticular respiration is sufficient. Macan (1962) reported on the work of Ambühl (1959). He found that Baetis vernus was scarce when the current speed was below 10 cm sec-1 and increased in relative numbers up to 40 cm sec-1. Ephemerella ignita (Figure 60) was most common at current speeds of 10-30 cm sec-1. Movements of another type – undulating the body (Figure 81) or fanning the gills (Figure 82) – can increase the rate of oxygen movement across the gills. Undulations typically begin as oxygen levels are low and are also used for swimming, a second way to gain more oxygen. These undulations are easily seen when high-oxygen-requiring mayflies are brought to the lab and put in quiet water. Ephemerellidae species accomplish water movement over their gills by moving the gill covers (Figure 82) up and down, fanning the gills. Trichoptera (caddisflies) are able to pump water through their cases (Figure 83) to renew oxygen. Humps and projections maintain space between the larva and its case, permitting water (and oxygen)

Chapter 11-1: Aquatic Insects: Biology

movement through the case. But these activities require energy and the insects cannot sustain prolonged use of these behaviors (Hynes 1970).

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bubble (Figure 85), as done by a number of free-swimming species. However, the plastron mechanism is useful to some of the Elmidae (Figure 84), tiny beetles that clamber among the bryophytes (Arnold & Macan 1969). The plastron is much like a diving bell. The insect traps a bubble of air and carries it beneath the water surface. As the insect breathes, it exchanges its CO2 for the O2 in the plastron. Oxygen in the water will diffuse into the bubble as the oxygen is depleted, but as the nitrogen leaves the bubble, the bubble shrinks. The same mechanism applies to other types of bubbles such as the one in Figure 85. Eventually the concentration of oxygen in the bubble is too low and the insect must resurface to grab another bubble, or grab one from a photosynthesizing plant, including bryophytes. The collection of bubbles on plants under water is known as pearling (Figure 86).

Figure 81. Baetis tricaudatus naiad showing the tail and abdomen flipped up in an undulation. Photo by Bob Henricks, with permission.

Figure 84. Stenelmis crenata showing plastron (white area under ventral side). Photo by M. J. Hatfield through Creative Commons.

Figure 82. Ephemerella subvaria naiad showing four gill covers on each side. Photo by Tom Murray, through Creative Commons.

Figure 85. Lancetes angusticollis adult from South Georgia clinging to moss. Note the anal air bubble used like a diving bell. Photo by Roger S. Key, through Creative Commons.

Obtaining Food

Figure 83. Limnephilus sp. showing spacer hump just behind the thorax. Photo by Jason Neuswanger, with permission.

Most of the bryophyte dwellers do not carry oxygen in the air bubble of a plastron (Figure 84) or other form of

Feeding strategies include shredders, gatherers, scrapers, and detritus feeders. Venturing away from the protective bryophyte substrate is dangerous because the insects can easily be swept away by the current in streams. Thus, it is not any surprise that many of the insects have adapted strategies that permit them to obtain food without venturing away from their safe site. Many are detritus

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feeders, and if they have adaptations to get enough oxygen, they can live in the silt or sand. Others such as the netspinning caddisflies (Figure 87) and the blackflies (Figure 88-Figure 89) trap their food as it flows by them. The very effective anchorage permits the Simuliidae (blackflies) to hang from the rear and expose the head fans (Figure 89) into the current to trap organic particles, including diatoms, for food. Some eat their surrounding homes – the bryophytes.

Figure 89. Simuliidae larva showing head fans that are used to capture food. Photo by Bob Henricks.

Figure 86. Riccia fluitans with pearling. Photo through Creative Commons.

Others, including some of the net-spinning Hydropsychidae (Figure 87), let the bryophytes do the trapping and eat the periphyton and detritus within the bryophyte mat. I base this assumption on finding many more larvae than nets among the mosses. The Chironomidae (Figure 90) live in leaf bases where detrital matter accumulates, obtaining both protection and food. In any case, the diet of the aquatic stage is usually quite different from that of the adult.

Figure 87. Cheumatopsyche nets on Fontinalis, trapping detritus and algae that flow by. Photo by Janice Glime.

Figure 90. Coryneura sp. (Chironomidae). Photo by Stephen Moore, Landcare Research, NZ, with permission.

Who Lives There?

Figure 88. Simuliidae larva head fans closed. Photo by Bob Henricks, with permission.

Aquatic bryophytes in mountain streams typically are replete with insects, crawling about and dining on the detritus and algae in the milieu. They find themselves safely out of the torrent above and tucked away from the view of fish and other predators. It seems like they should have a pretty cushy life. When I began my studies on insects living among bryophytes in Appalachian Mountain, USA, streams, few studies were available for comparison, and most of those

Chapter 11-1: Aquatic Insects: Biology

were from Europe. Like the development of keys for bryophytes, the development of keys for aquatic insects lagged way behind what was needed. To further complicate the problem, many of the insects had been described from adults, but studies to link the immature aquatic stages to their adults were lacking for many. It was the insect version of the early Takakia classification problem. As I delved into the many more recent papers to prepare this chapter, I found many unfamiliar names of genera, only to discover that those familiar genera from nearly 50 years ago had gone through reclassification and were now represented under multiple new names, especially at the generic level. To further complicate these changes in generic concepts, the insects, like the bryophytes, comprise many microspecies. Limited dispersal distances for short-lived adult stages, mountain and land barriers, and disconnected stream or lake systems all contributed to the isolation needed for development of differences in physiology, behavior, phenology, and morphology (see for example Hughes et al. 1999; Monaghan et al. 2002). As bryologists we are well aware of these problems in classifying things separated by great distances, but for these insects the microspecies differences can be manifest over much shorter distances, a phenomenon that has been recognized in some aquatic bryophytes as well (Glime 1987; Shaw & Allen 2000). Nevertheless, there are lessons to learn from the orders, families, and even the genera as we examine who lives among the bryophytes – and why. Drozd et al. (2009) used pitfall traps to compare invertebrate inhabitants related to bryophytes in the mountain areas (384-1200 m asl) of the Czech Republic. In most cases, the Collembola were the most abundant group except for the high number of ants at Podolánky. The numbers differed by bryophyte and moisture level (Figure 92). Insects were highest in the dry litter control (within 2

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m of moss area). The lowest numbers were in wet Sphagnum fallax (Figure 91).

Figure 91. Sphagnum fallax with capsules, the species with the lowest number of Collembola among bryophytes in the mountainous areas of the Czech Republic. David T. Holyoak, with permission.

Drozd and coworkers (2009) considered several caveats in interpreting their results. Some of the invertebrates move about little and would therefore be poorly represented in the pitfall traps. Others that do move about would move easily between the bryophytes and litter, possibly only passing over the bryophytes in their search for food. Others may reside among the bryophytes as transient visitors, seeking escape from a predator or avoiding the desiccation common in more open areas, but returning to the litter habitat when that environment was safe. In any case, insects that met all their needs within the bryophyte mat would be under-represented in the pitfall traps.

Figure 92. Abundance of taxonomical groups in pitfall traps associated with several species of bryophytes in dry, moist, and wet conditions at five locations in mountains of the Czech Republic. The scale at right is for ant data (Formicoidea) from Podolánky. Redrawn from Drozd et al. 2009. Controls are litter areas

The insects found among the mosses in streams are mostly Ephemeroptera (mayflies), Plecoptera (stoneflies), Trichoptera (caddisflies), Diptera (flies), and

Coleoptera (beetles) (Needham & Christenson 1927; Wesenberg-Lund 1943; Cowie & Winterbourn 1979; Glime 1994; Gislason et al. 2001). But moving about

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among stems and leaves of mosses is not easy for the weaklegged or swimming insects in the small spaces. Hence, as already noted, most of the inhabitants are small (Thienemann 1912; Glime 1994; Amos 1999; Drazina et al. 2011). This also means that young, immature naiads of Ephemeroptera and Plecoptera are common (Stern & Stern 1969). Many species overwinter as eggs on the mosses, then begin their immature lives there. Among the Diptera, Dicranota (Figure 93), Atherix (Figure 94), and Simulium (Figure 51-Figure 53) are common at this time; likewise, young Elmidae (larvae; Figure 95) are common among the mosses (Thienemann 1912).

In a New Zealand stream, Cowie and Winterbourn (1979) found 44 species of invertebrates, mainly immature stages of insects. The moss Acrophyllum sp. (Figure 96) hosted the stonefly Austroperla cyrene (Figure 97), a species of beetle in the Helodidae (Figure 98), and a triclad, Neppia montana (Figure 99); the moss Fissidens sp. (Figure 100) hosted the stonefly Zelandoperla fenestrata (see Figure 101), the caddisfly Zelolessica cheira (Figure 102), a fly in the family Empididae (Figure 103), and several species of midges (Chironomidae; Figure 90); . The moss Cratoneuropsis (Figure 104) had only one common taxon, a terrestrial isopod, Styloniscus otakensis, suggesting that the streamside Cratoneuropsis habitat is more terrestrial than aquatic. In addition to water saturation and flow rates, the ability of mosses to trap detritus was important in determining invertebrate inhabitants.

Figure 93. Dicranota larva, a common stream moss inhabitant. Photo by Tom Murray, through Creative Commons.

Figure 96. Achrophyllum quadrifarium from New Zealand, home to beetles in Helodidae. Photo by Bill & Nancy Malcolm, with permission.

Figure 94. Atherix sp. larva, a common dweller among stream bryophytes. Photo by Jason Neuswanger, with permission. Figure 97. Austroperla cyrene from NZ. Photo by Steve Pawson, permission pending.

Figure 95. Elmidae larva, a common beetle larva among stream bryophytes. Photo by Stephen Moore, Landcare Research, NZ, with permission.

Figure 98. Helodidae adult, member of a family that lives among leaves of the moss Acrophyllum sp. Photo from , with permission.

Chapter 11-1: Aquatic Insects: Biology

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Figure 102. Zelolessica sp., an inhabitant of aquatic Fissidens in New Zealand. Photo by Stephen Moore, Landcare Research, NZ, with permission. Figure 99. Neppia, an inhabitant of the moss Acrophyllum sp. Photo by Stephen Moore, Landcare Research, NZ., with permission

Figure 103. Empididae larva, an inhabitant of aquatic Fissidens in New Zealand. Photo by Stephen Moore, Landcare Research, NZ, with permission.

Figure 100. Fissidens fontanus with Amano shrimp in an aquarium. Photo through Creative Commons.

Figure 104. Cratoneuropsis relaxa, in a genus that commonly houses isopods but few insects in New Zealand. Photo by Tom Thekathyil, with permission.

Figure 101. Zelandoperla sp., an inhabitant of Fissidens in New Zealand. Photo by Stephen Moore, Landcare Research NZ, with permission.

Suren (1988) examined faunal assemblages in New Zealand alpine streams, with the stoneflies (Plecoptera) Zelandoperla (Figure 101) and Zelandobius (Figure 105) and midge larvae (Chironomidae; Figure 90) being dominant. The mosses supported 5-15 times as many invertebrates as did the rocky habitats. In addition to these dominant insects, several non-insect invertebrates were dominant.

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And channel stability likewise determines the stability of bryophytes, hence playing a role in the bryophyte fauna. In my study of the insects inhabiting the bryophytes of mid-Appalachian Mountain streams, I identified 141 species occurring among 10 species of bryophytes in 28 streams, and that does not include the species of the Chironomidae (Figure 90), which were identified only to family (Glime 1994). The smallest of the insects occurred on the leafy liverwort Scapania undulata (Figure 106) and the largest could be found on various species of Fontinalis (Figure 107). As in many other studies, the most abundant insects were midges (Chironomidae), the stoneflies Leuctra (Figure 30) and Isoperla bilineata (Figure 108), and the blackflies (Simulium tuberosum; Figure 109).

Figure 105. Zelandobius illiesi, a stonefly genus that is common among alpine stream mosses in New Zealand. Photo by Stephen Moore, Landcare Research NZ, with permission.

One of the interesting questions about bryophyte fauna is whether any species has a unique fauna. So far we have seen little specificity among the other invertebrates. Nevertheless, differences may exist dependent upon the niches of the bryophytes themselves. Some bryophytes occupy fast flow, some occupy areas where they spend part of the year above water, some are deep, and certainly differences exist among growth forms that create differences in the protection they afford. And some Trichoptera use liverworts or mosses to construct their cases, forcing them to live with certain species. Coinciding with these differences are the kinds of food the bryophyte habitats provide, again affecting who can survive there. Paavola (2003) examined the concordance among the macroinvertebrates, bryophytes, and fish to look for possible surrogates to describe the system and its state of health. Surrogates are groups of organisms that can be used to assess suitability of a habitat for another group of organisms such as fish. When considered across drainage systems, there was strong concordance, but within a single river system that concordance was weak. Bryophyte locations in the Paavola (2003) study were mainly related to nutrient levels and in-stream complexity, whereas macroinvertebrates correlated with stream size and fish correlated with oxygen levels, depth, and substrate size. But macroinvertebrates also relate to in-stream complexity (Allan 1975; Hart 1978; Trush 1979; Wise & Molles 1979; Williams 1980; Vinson & Hawkins 1998) and to substrate texture (Glime & Clemons 1972). And bryophytes add to that complexity. Some of the genera that inhabit bryophytes are also common in leaf packs – a substrate that provides cover and detritus for food. These include Baetis (Figure 45), Leuctra (Figure 30), and Chironomidae (Figure 90) (Robinson et al. 1998). Due to differences in growing season, ice-free season, winter severity, available food, and flow regime changes from year to year, the fauna assemblage can also change from year to year. This can result in the temporary disappearance of an entire species, or even an entire order (Milner et al. 2006). This disappearance is particularly true for Plecoptera. Channel stability is important in determining faunal stability, but a normally stable channel can suffer from heavy rains or flooding during snow melt.

Figure 106. Scapania undulata, home for the smallest aquatic insects. Photo by Michael Lüth, with permission.

Figure 107. Fontinalis antipyretica, a large moss that houses the largest moss dwellers. Photo by Bernd Haynold Wikimedia Commons.

Figure 108. Isoperla bilineata, a common stream moss dweller in the Appalachian Mountains, USA. Photo by Bob Henricks, with permission.

Chapter 11-1: Aquatic Insects: Biology

Figure 109. Simulium tuberosum, a common inhabitant of bryophytes in Appalachian Mountain streams. Photo by Tom Murray, through Creative Commons.

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Figure 110. Rhyacophila invaria larva, a common freeliving caddisfly among Platyhypnidium riparioides in Appalachian Mountain, USA, streams. Photo by Donald S. Chandler, with permission.

Heino and Korsu (2008) questioned the species-area concept in aquatic systems, examining rocks and bryophyte cover in two river sites. They found only a weak speciesarea relationship on stream stones. On the other hand, bryophyte biomass was important both in supporting species richness and in increasing number of individuals of stream macroinvertebrates. They suggested that cover was important in increasing number of individuals and that the species richness was a subsequent passive response. The bryophyte biomass can be expected to increase with time, whereas the area of stones will not. The mechanisms that promote these species-area relationships need to be demonstrated experimentally. These could involve food relationships, sampling methods, niche space, flood disturbance, predation refugia, or flow regime. Specificity Many streams have only one dominant bryophyte, and others have the species intermingled. These conditions complicate any attempts to determine insect preference. Nevertheless, some specificity seems to exist, but keep in mind that it might be a preference of both insect and bryophyte for the same stream conditions. The caddisfly Rhyacophila cf. invaria (Figure 110) was present in 36% of the collections (Figure 118) of Platyhypnidium riparioides (Figure 111) in mid-Appalachian Mountain, USA, streams, but totally absent among Hygroamblystegium fluviatile (Figure 112), despite the frequent occurrence of these two mosses in the same streams, often on the same rocks (Glime 1994). Rhyacophila carolina (Figure 1) reached its greatest abundance in clumps of the leafy liverwort Scapania undulata (Figure 106; Figure 118). Less distinct preferences occurred in the elmid beetle larva Optioservus sp. (Figure 113; Figure 118) [36% of Hygroamblystegium fluviatile (Figure 112), 7% of Platyhypnidium riparioides (Figure 111)] (Glime 1994). The stonefly Pteronarcys proteus (Figure 114) occurred in 24% of the H. fluviatile, 7% of the P. riparioides, and never in any of the other species, including Scapania undulata (Figure 106), Fontinalis dalecarlica (Figure 115), and Hygrohypnum spp. (Figure 116) (Figure 118).

Figure 111. Platyhypnidium riparioides, a common moss in Appalachian Mountain, USA streams. Photo by David T. Holyoak, with permission.

Figure 112. Hygroamblystegium fluviatile, a common moss for insect fauna in Appalachian Mountain, USA, streams. Photo by Michael Lüth, with permission.

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58% of the collections and Simulium tuberosum (Figure 109) in 75% of the collections (Figure 118) of this liverwort in mid-Appalachian Mountain, USA, streams (Glime 1994). But S. tuberosum also occurred in 78% of the Fontinalis dalecarlica (Figure 115) collections (Figure 118).

Figure 113. Optioservus sp., a common beetle larva among Hygroamblystegium fluviatile and Platyhypnidium riparioides in Appalachian Mountain, USA, streams. Photo by Arlo Pelegrin, with permission.

Figure 116. Hygrohypnum luridum, a moss that is not suitable habitat for the large Pteronarcys in the streams of the Appalachian Mountains, USA. Photo by Michael Lüth, with permission.

Figure 114. Pteronarcys proteus, a stonefly that seems to have some selection in bryophytes it will inhabit. Photo by Jason Neuswanger, with permission.

Figure 117. Prosimulium hirtipes, a common blackfly on the liverwort Scapania undulata. Photo by Janice Glime. Figure 115. Fontinalis dalecarlica, a large moss but that did not house Pteronarcys proteus in Appalachian Mountain, USA, streams. Photo by Kristoffer Hylander, with permission.

The liverwort Scapania undulata (Figure 106) has a different form from that of any of the mosses. This flattened habit seems to favor the fast-water members of Simuliidae, with Prosimulium hirtipes (Figure 117) in

Diversity differs little among bryophyte species (Figure 119), although richness can be higher in the larger Fontinalis (Figure 115) species (Glime 1968, 1994). Fontinalis species are also the only ones that typically house larger insects. Scapania (Figure 106), on the other hand, housed the smallest insects in the Appalachian Mountains, USA, streams.

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Figure 118. Frequencies of insects on five bryophyte species sampled in 28 streams in the middle Appalachian Mountains, USA. Only insects with at least 10% frequency on at least one species of bryophyte are included. The bryophyte name appears by the group of species that was most abundant on that bryophyte; the name applies to all groups in that frame. From Glime 1994.

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partition niches, with different sizes of insects occupying different niches. This means that larger members of a genus or family can occupy the same moss clump as younger members of other species in that family feeding group because they have different feeding niches. In some cases this niche partitioning is done by a seasonal migration to a different substrate. Hildrew and Edington (1979; see also Muotka 1990) found that early instars of Hydropsyche siltalai (Figure 121) and H. pellucidula (Figure 122) occupied the same rocks. However, in spring H. siltalai migrates to moss beds, but H. pellucidula was totally absent among the mosses at that time.

Figure 119. Comparison of mean insect richness and Shannon diversity on a leafy liverwort (Scapania undulata) and four species of mosses in 28 mid Appalachian Mountain streams, USA. Redrawn from Glime 1994.

Perhaps the greatest specificity is among some of the case-making caddisflies (Trichoptera). Several species in the Hydroptilidae make their cases exclusively from bryophytes, including Palaeagapetus celsus from leafy liverworts (Flint 1962; Glime 1978, 1994). The flat leaves of Scapania undulata seem to be ideal for their method of cutting nearly circular pieces that they cement together for the cases, apparently causing these larvae to live almost exclusively among leafy liverworts (Glime 1978, 1994). Likewise, in the Brachycentridae Adicrophleps hitchcocki (Figure 120) uses bits of Fontinalis (Figure 107) leaves or other mosses to construct its cases (Flint 1965; Glime 1994). When it uses Hygroamblystegium fluviatile (Figure 112) it may use only costae to make the case, sometimes leaving the ends of the costae dangling from the case (Glime 1994). The Chironomidae (Figure 90), as a family, was present in 98-100% of the collections of all species (Figure 118), but these comprised multiple species that could have differed among bryophytes and streams. The acidity may affect the inhabitants, causing an appearance of bryophyte specificity. Frost (1942) found that the Plecoptera and Coleoptera were less important in the calcareous stream than in the acid stream, whereas the Ephemeroptera and Trichoptera reached their greatest density in the more calcareous stream.

Figure 120. Adicrophleps hitchcocki showing case made with Hygroamblystegium. Note costae protruding near opening. Photo by Bob Henricks, with permission.

Figure 121. Hydropsyche siltalai, a caddisfly larva that moves to moss beds as it gets older, avoiding competition with H. pellucidula. Photo by Urmas Kruus, with permission.

Seasons One reason for insects to live among bryophytes is to escape the cold of winter. To this end, some insects are more abundant in streams in the winter, but many spend the winter as pupae or eggs. Thienemann (1912) found that young fauna were especially common among mosses in summer. Seasons can

Figure 122. Hydropsyche pellucidula, a net-spinning caddisfly that avoids niche competition with H. siltalai by avoiding moss beds when the latter migrates there. Photo by Niels Sloth, with permission.

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In the Appalachian Mountain streams, the total numbers diminish in the winter (Glime 1968), but some insects, like the blackfly Prosimulium hirtipes (Figure 51Figure 53; Figure 123), hatch in late fall and spend the winter in the water, emerging as adults in spring.

Figure 124. Seasonal changes in species diversity (H') among mosses (Fontinalis spp. ▬) and liverworts (Scapania undulata --) in Appalachian Mountain, USA, streams. Figure 123. Relative abundance of the six most common insects among bryophytes in five collecting seasons in Appalachian Mountain, USA, streams. Based on Glime 1968.

In these Appalachian streams, counts do not always track diversity and richness (Figure 127; Glime 1994). What is more interesting is that Shannon diversity (following Patten 1962) and species richness do not always agree. This may be the result of the differences in counts, which are reflected in the Shannon diversity: R

H' = -Σ pi log2 pi i=1

where pi = the proportion of individuals belonging to the ith type, or count of the species divided by total count of all species R = richness, or total number of species Richness, on the other hand, is simply the number of species present. In these streams, Shannon diversity was highest in March, but richness was highest in July. It is also interesting that these seasonal differences can be different among bryophyte species (Figure 124-Figure 127). The ever-present Chironomidae (Figure 90) often peak among the mosses in winter (Frost 1942), but in the Appalachian Mountain, USA, streams the peak is midsummer (Figure 123) (Glime 1968). Whitehead (1935) suggested that this might be a behavioral attribute in which the insects seek shelter among the mosses to avoid or respond to the ravages of flooding. But clearly the insects differ among orders, families, and seasons, as seen in these Appalachian Mountain streams (Figure 125-Figure 126).

Figure 125. Relative numbers of the most abundant species (>3 occurrences) of insects per gram dry weight of bryophyte in December in Appalachian Mountain, USA, streams. Frequencies appear at right end of each bar. Based on Glime 1968.

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Figure 126. Seasonal relative numbers of the most abundant species (>3 occurrences) of insects per gram dry weight of bryophyte in Appalachian Mountain, USA, streams. Frequencies appear at right end of each bar. Based on Glime 1968.

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Table 2. Common genera of bryophyte-dwelling aquatic insects. Numbers refer to references: (1) Percival & Whitehead 1930 (UK); (2) Glime 1994 (Appalachian Mountains, USA); (3) Thienemann 1912 (North Rhine-Westphalia); (4) Suren 1988 (alpine NZ); (5) Muttkowski & Smith 1929 (Yellowstone USA); (6) Frost 1942 (UK); (7) Tada & Satake 1994 (Japan); (8) Krno 1990 (Slavakia). Only studies that included all insect groups are included; note that most studies did not identify genera of the Chironomidae. COLLEMBOLA Isotomidae – Isotoma EPHEMEROPTERA Baetidae Baetis Baetiscidae – Baetisca Caenidae – Caenis Ephemerellidae Drunella Ephemerella Torleya Heptageniidae Cinygmula Heptagenia Rhithrogena Stenacron Leptophlebiidae Habroleptoides Leptophlebia Paraleptophlebia ODONATA Gomphidae – Gomphus PLECOPTERA Chloroperlidae – Chloroperla Chloroperla Gripopterygidae Zelandobius Zelandoperla Leuctridae – Leuctra Nemouridae Amphinemura Nemoura Protonemura Perlidae Acroneuria Perlodidae Megarcys Isoperla Peltoperlidae – Peltoperla Pteronarcidae – Pteronarcys Taeniopterygidae – Taeniopteryx HEMIPTERA Veliidae – Microvelia DIPTERA Athericidae – Atherix Ceratopogonidae Bezzia Dasyhelea Chironomidae Corynoneura Cricotopus Dactylocladius Diamesa Orthocladius Tanytarsus Thienemanniella Empididae Clinocera Hemerodromia Limoniidae – Antocha Muscidae – Limnophora

2 2 1,2,3,5,6,7,8 1,2,3,6,7,8 1,2,3,6,7,8 2 6,8 1,2,5,6,7,8 5,7 1,2,6,7,8 8 1,5,7,8 7 1,5 8 2 2,6,8 8 6 2,6 2 2 1,2,4,5,6,7,8 6,7 6 4 4 4 1,2,6,8 1,2,4,5,6,7,8 6,7,8 2,8 6,7,8 5 5 2,6,7,8 7 2,6,7,8 2 2,5 2,6 2 2 1,2,3,4,6,7,8 2,3 2,8 2 2 1,2,3,4,6,7 3 3 3 3 3,7 3 3 2 6 6 7 1,3,6

Pediciidae – Dicranota Psychodidae – Pericoma Simuliidae Cnephia Odagmia Prosimulium Simulium Tipulidae Hexatoma Limnobiinae Limnophora Tipula COLEOPTERA Dytiscidae – Ilybius Elmidae Dubiraphia Elmis Esolus Limnius Optioservus Promoresia elegans Stenelmis crenata Gyrinidae – Gyrinus Hydraenidae Hydraena Limnebius TRICHOPTERA Brachycentridae Adicrophleps Brachycentrus Micrasema Hydropsychidae Arctopsyche Cheumatopsyche Diplectrona Hydropsyche Parapsyche Hydroptilidae Agapetus Agraylea Hydroptila Ithytrichia Oxyethira Paleagapetus Leptoceridae – Leptocerus Lepidostomatidae – Lepidostoma Limnephilidae Allogamus Drusus Parachiona Pseudostenophylax Philopotamidae Chimarra Dolophiloides Philopotamus Polycentropodidae – Polycentropus Psychomyiidae – Psychomyia Rhyacophilidae – Rhyacophila Uenoidae Neophylax Thremma

3,6 2,3,6,8 2,6,7,8 2 8 2,8 2,6 1,2,6,7 1,2 6 2 2,6 1,2,3,6,8 2 1,2,3,6,8 2 1,8 3,6 3,6 2 2 2 6 3 3 3 1,2,3,5,6,7,8 2,3,7,8 2 5,8 2,3,7 1,2,3,8 7 2 2 1,2,3,6,8 2 1,2 1,6 2 1,2,3,6 1,2,3,6 2,3,6 2 1,6 1,2,6 7,8 8 8 8 7 1,2,3 2,6 2 1,3 1,2,6 1,6 1,2,3,6,7,8 2,5 2 5

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invertebrates landed in the net, so I proceeded with my hand collections. These were placed in baby food jars with 95% alcohol and a few drops of glycerine added to prevent predation and decay until the jars reached the lab. I removed the insects with microforceps while systematically searching through a dissecting microscope at 10 X. Frost (1942) was one the early surveyors of bryophyte fauna. Her sample size was 200 g of wet moss. Kamler (1967) cut 10x10 cm samples under water. Maurer and Brusven (1983) were particularly careful. They surrounded the moss with a nylon organdy net of 250 µm mesh while removing the moss from the stream, then used several washes and hand picking to extract the insects. Armitage (1961) used the modified square foot sampler, similar to the Surber sampler (Figure 128) used by Gurtz and Wallace (1984), to catch insects from rocks, mosses, sticks, and under rubble in streams. However, most bryophyte dwellers are adapted to clinging to the bryophyte and require more than a little disturbance to free them. This leads to underestimates of the bryophyte fauna relative to those among the rubble of the stream bottom and also to species bias. Wulfhorst (1994) modified this method slightly, using a box sampler to cut a square of 14 cm2 to sample mosses in an acid stream. The moss samples were quantified by volume using displacement of water in a graduated cylinder. In his New Zealand studies, Suren (1988) likewise used a Surber sampler (Figure 128) with 100 µm mesh to sample 0.01 sq m. Rocky areas were sampled with a 0.02 sq m sampler that had a thick foam flange around the bottom to provide a seal with the substrate. Mosses were scraped into the sampler with a razor blade. This method permitted the same area to be sampled in both rock and moss areas.

Figure 127. Bryophyte-dwelling insect seasonal richness, species diversity, and counts from handful samples. Redrawn from Glime 1994.

Sampling Sampling of the fauna of aquatic bryophytes can be a time-consuming process. And sampling used for most terrestrial or stream habitats can introduce strong biases for these sheltered species. My own methods were to use hand grabs, then determine the dry weight of the bryophytes after the fauna had been removed. This sampling kept the internal fauna intact, and to test for surface losses, I initially placed a net just downstream from my collections. Very few

Figure 128. Surber sampler being used as drift net for winter stream drift sampling. Photo by Janice Glime.

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Preservative It is important to understand the role of the preservative. Not only does it keep the organisms from decaying and being eaten by cohabitants during the period until the sample can be examined, but it increases the extraction efficiency for flotation techniques (discussed below), at least in a sucrose solution of 1.12 specific gravity (Pask & Costa 1971). In samples preserved for 14 days in 10% formalin compared to those not preserved but examined the same day, the preserved insects had a recovery of 91% whereas those with no preservation had only an 83% recovery rate. Since any collection of bryophytes will bring significant water with it, it is necessary to use a higher concentration than that used when preserving just insects. I added 95% alcohol to my bryophyte collections (with insects), hoping to achieve a concentration of around 70%. Extraction The least bias in extraction can be achieved by careful hand picking while observing through a dissecting microscope. When I first tried to publish my Ph. D. work, the reviewer wanted to know what method I had used to "estimate" the numbers of Chironomidae, which could reach thousands in a single handful of moss. But I had removed and counted every single one of them at 10X magnification! Gurtz and Wallace (1984) also hand-picked invertebrates from the mosses at 7X under a dissecting microscope, using a count per dry weight of moss. There are simpler and less time-consuming methods for those who don't want to spend three years searching among the bryophytes with a microscope. But, these each have their biases. The Tullgren funnel (Andrew & Rodgerson 1999) creates a temperature gradient over the sample, typically with a tungsten light bulb above it. Mobile organisms will move away from the higher temperatures and fall into a collecting vessel with alcohol or mixed preservative. But not all insects move quickly, and some may die from the heat and desiccation before falling to their death in the alcohol below. Furthermore, some will die before reaching the lab due to the reduced oxygen. Fairchild et al. (1987) developed a behavioral method for extracting invertebrates from Sphagnum (Figure 91). The method includes a vertical temperature gradient coupled with dissolved oxygen gradients in a column of water containing the Sphagnum sample. They determined the overall extraction to be 85% efficient (n=4). I do have concerns about bias in the species extracted. Teskey (1969) developed a method especially for sampling the small flies of the family Tabanidae. He used a combination of a specially designed sieve with a multiple Berlese funnel (similar to the Baermann funnel in Figure 129) or by using hand searching to sample these larvae. But to identify the larvae, as in many of the aquatic taxa, they had to be reared to adults. Cochrane (1913) used sieves to collect larvae of Culicoides furensoides (Diptera: Ceratopogonidae) from Sphagnum (Figure 91).

Figure 129. Baermann funnel using moss sample and modified from the Berlese funnel setup, using water instead of air. Modified from Briones 2006.

Flotation Any flotation technique requires that the density of the flotation liquid be greater than that of the insects but less than that of the debris (Lackey & May 1971). The 1.12 specific gravity sucrose solution of Pask and Costa (1971) works well in this regard. The kerosene phase separation extracts more total individuals than those extracted by sugar flotation or the Tullgren funnel, particularly more Acari (mites) and Collembola (springtails) (Andrew & Rodgerson 1999). Fast (1970) pointed out that calling the flotation techniques "flotation" was a misnomer. While the sugar solution is important, many of the organisms remain lodged at leaf bases or caught among the leaves and stems. He preserved samples with 10% formalin. To separate the organisms, he used 360 g sucrose per liter of water and gave the samples only one immersion in the sugar solution. He then sorted at 3.5X magnification. One problem I found with the flotation method was that tiny creatures like the Chironomidae got trapped in the surface tension. They were almost impossible to pick up, so they needed to be trapped on a filter. By the time you have then picked them off the filter, you might as well sort them directly from the moss and learn about their hideouts and spatial relationships at the same time. Hribar (1990) reviewed ten methods for sampling biting midge larvae. Some of these will work for aquatic bryophytes. Hribar was successful in extracting larvae of Ceratopogonidae (Alluaudomyia, Atrichopogon, Bezzia, Culicoides, Dasyhelea, and Forcipomyia) from Fontinalis

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(Figure 107) and aquatic liverworts by using a Berlese funnel (see Figure 129). He found that sugar flotation and salt flotation provided similar results, but the sugar flotation caused less mortality. Magnesium sulfate is a slower process but results in fewer deaths than salt solutions. Nevertheless, he considered agar extraction and salt flotation to be the most effective for collecting larvae. Sieving, sieving plus salt flotation, and Berlese funnels worked well for mosses. In short, unbiased sampling to determine numbers of insects living among bryophytes requires time and patience.

mm. He found no differences in the fauna between artificial and real mosses in a New Zealand stream. The artificial mosses even had abundant periphyton growth [especially Epithemia (Figure 132) in winter and spring], but their accumulation of detritus and silt was sparse. This perhaps explains the significantly lower numbers of detritus feeders such as Acarina (mites), Collembola (springtails), Tardigrada (water bears), Dorylaimoidea (nematodes), and Ostracoda (seed shrimp) on the artificial mosses.

Artificial Mosses Several researchers have attempted to explain the role of aquatic bryophytes by using artificial mosses. Glime and Clemons (1972) used strips of plastic and bundles of string (Figure 130) as artificial mosses. The plastic permitted colonization by periphyton (attached organisms) but lacked the chambering found among mosses; only 13 species occurred on the 33 samples. The string offered a soft substrate with limited chambers; 23 species of aquatic insects occurred on the 35 samples, some of which were not present on the real mosses. The real mosses [Fontinalis novae-angliae (Figure 131) & F. dalecarlica (Figure 115) had 25 species among the 46 samples, differing little in overall richness from that of the string mosses. It appeared that density of insects was higher among real mosses, but there was no common base upon which to compare them. It is interesting that the Shannon diversity differed little among the three substrata (1.8 on moss, 1.9 on string, and 1.7 on plastic). Nevertheless, the Shannon diversity (d) on plastic was significantly different from that on mosses or strings. The lack of complexity and smaller surface area of the plastic may have accounted for the limited diversity.

Figure 131. Fontinalis novae-angliae, a moss with around 25 species of insects in a New Hampshire, USA, stream. Photo by Janice Glime.

Figure 132. Epithemia sp., a common diatom genus on mosses, on a filamentous alga. Photo by Jason Oyadomari, with permission.

Summary

Figure 130. Artificial mosses made of cotton string. Photo by Janice Glime; see Glime & Clemons 1972.

Suren (1988) used nylon twine (5 cm long, 1 mm thick) to weave squares 0.01 m2 thick with a pore size of 4

Aquatic insects are those insects that spend part of their life cycles in the water, usually as a means of escaping the harsher environment on land during one or more seasonal conditions. For most, the immature stages are those requiring such an escape. Aquatic bryophyte dwellers include the Collembola (no longer considered to be insects) that look like miniature adults when born. The hemimetabolous insects include the nymphs of Hemiptera that look like their parents from birth and simply grow larger. The naiads of Ephemeroptera, Odonata, and Plecoptera are likewise hemimetabolous, but the naiads often differ from the adults in having gills, different mouth parts, and wing

Chapter 11-1: Aquatic Insects: Biology

pads instead of wings. Their life cycle goes from egg/embryo to naiad to adult. The holometabolous insects have four distinct stages in the life cycle – egg/embryo, larva, pupa, and adult. These orders, among bryophytes, include Coleoptera, Neuroptera, Megaloptera, Trichoptera, and Diptera. Some have gills as larvae but not as adults. As an escape from unfavorable conditions, the life cycle stages often respond to environmental cues, including photoperiod, temperature, or available food. Aquatic insects are especially sensitive to temperature, and many of them are in the water for winter to escape the below-freezing temperatures in the terrestrial environment. Some overwinter as dormant eggs or pupae, others as active larvae, naiads, or adults. Structural adaptations include streamlining, small size, gills, hooks or silk for anchoring, gill covers, and cases or tubes. They move about in the bryophyte clumps to achieve the best oxygen and flow conditions, often leaving as they grow larger. Oxygen may be obtained through gills, cuticle, or a plastron that carries an air bubble from the surface or from photosynthesizing plants or algae. Bryophyte dwellers include shredders, gatherers, scrapers, and detritus feeders that prey upon smaller organisms, including periphyton, or eat the detritus gathered by the bryophytes. Some eat the bryophytes. Some make nets to trap food. A few species have a specific requirement for bryophytes for case building, but most simply need a refuge with adequate oxygen, food, and cover. Sampling is often done with nets, but is best by hand grabs and hand sorting. The faster methods such as nets are commonly used, but they have biases against interior and clinging organisms. Sorting by flotation or Berlese funnels has similar biases. Artificial mosses can sample colonizers but they may not provide the food sources needed and require somewhat lengthy colonization times.

Acknowledgments For this chapter I must thank the many students who have spent the night in the cold of winter or mosquitoes of summer to sample the streams and their bryophyte dwellers. And a special thanks to Arlene Jim, my sister Eileen Dumire, and many others who spent numerous hours staring through a dissecting microscope and pulling insects out of pickled bryophytes. And in my younger days, my parents, Mildred and Gilbert Glime, chauffeured me to streams throughout the middle Appalachians so that I could study this fascinating group of bryophyte dwellers. My sister often accompanied me on collecting trips and served as the reviewer for this chapter, giving me the perspective of a non-biologist. In my early days, Lewis Berner, Oliver Flint, Glenn Wiggins, and Ken Cummins encouraged me and helped me in identifications. Jason Neuswanger, Roger Rohrbeck, and Arlen Thomason have been helped me with updating scientific names and suggesting contacts. Jason Neuswanger and Bob Henricks have given me permission to use their large libraries of aquatic insect images. Peter Buchanan gave me permission to use the invaluable

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collection of images at Landcare Research, NZ. Throughout the insect chapters, I have constantly appreciated all the photographers, both named and anonymous, who have made their images available through Creative Commons. This project would be far less complete without Google Scholar.

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Glime, J. M. 1968. Aquatic Insect Communities among Appalachian Stream Bryophytes. Ph.D. Dissertation, Michigan State University, East Lansing, MI, 180 pp. Glime, J. M. 1978. Insect utilization of bryophytes. Bryologist 81: 186-187. Glime, J. M. 1987. Phytogeographic implications of a Fontinalis (Bryopsida) growth model based on temperature and flow conditions for six species. Mem. N. Y. Bot. Gard. 45: 154170. Glime, J. M. 1994. Bryophytes as homes for stream insects. Hikobia 11: 483-497. Glime, J. M. and Clemons, R. M. 1972. Species diversity of stream insects on Fontinalis spp. compared to diversity on artificial substrates. Ecology 53: 458-464. Golubkov, S. M., Tiunova, T. M., and Kocharina, S. L. 1992. Dependence of the respiration rate of aquatic insects upon the oxygen concentration in running and still water. Aquat. Ins. 14(3): 137-144. Gros, A. 1923. Etudes sur les premieres stades des Ephemeres du Jura francais. Ann. Biol. Lacust. 12: 49-74. Gurtz, M. E. and Wallace, J. B. 1984. Substrate-mediated response of stream invertebrates to disturbance. Ecology 65: 1556-1569. Harper, P. P. 1973. Life histories of Nemouridae and Leuctridae in southern Ontario (Plecoptera). Hydrobiologia 41: 309356. Hart, D. D. 1978. Diversity in stream insects: Regulation by rock size and microspatial complexity. Internat. Verein. Theoret. Angew. Limnol. Verhand. 20: 1376-1381. Hawkins, C. S. 1984. Substrate associations and longitudinal distribution in species of Ephemerellidae (Ephemeroptera: Insecta) from western Oregon. Freshwat. Invert. Biol. 5: 181-188. Heino, J. and Korsu, K., 2008. Testing species-stone area and species-bryophyte cover relationships in riverine macroinvertebrates at small scales. Freshwat. Biol. 53: 558568. Hildrew, A. G. and Edington, J. M. 1979. Factors facilitating the coexistence of hydropsychid caddis larvae (Trichoptera) in the same river system. J. Anim. Ecol. 48: 557-576. Hoback, W. W. and Stanley, D. W. 2001. Insects in hypoxia. J. Insect Physiol. 47: 533-542. Hodkinson, I. D. and Bird, J. M. 2004. Anoxia tolerance in high Arctic terrestrial microarthropods. Ecol. Entomol. 29: 506509. Hora, S. L. 1930. Ecology, bionomics and evolution of the torrential fauna with special reference to the organs of attachment. Phil. Trans. Royal Soc. London B 218: 171-282. Hribar, L. J. 1990. A review of methods for recovering biting midge larvae (Diptera: Ceratopogonidae) from substrate samples. J. Agric. Entomol. 7: 71-77. Hughes, J. M., Mather, P. B., Sheldon, A. L., and Allendorf, F. W. 1999. Genetic structure of the stonefly, Yoraperla brevis, populations: The extent of gene flow among adjacent montane streams. Freshwat. Biol. 41: 63-72. Hughs, D. A. 1966. The role of responses to light in the selection and maintenance of microhabitat by the nymphs of two species of mayfly. Anim. Behav. 14: 17-33. Humphries, C. F. and Frost, W. E. 1937. River Liffey Survey. The chironomid fauna of submerged mosses. Proc. Roy. Irish Acad. Ser. B 43: 161-181. Hynes, H. B. N. 1961. The invertebrate fauna of a Welsh mountain stream. Arch. Hydrobiol. 57: 344-388.

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Hynes, H. B. N. 1970. The ecology of stream insects. Ann. Rev. Entomol. 15: 25-42. Illies, J. 1952. Die molle. Faunistisch-okologische Untersuchungen an einem Forellenbach im Lipper Bergland. Arch. Hydrobiol. 46: 424-612. Illies, J. 1961. Versuch einer allgemeiner biozonotischen Gliederung zur Fliessguwasser. Internat. Rev. Gesamt. Hydrobiol. 46: 205-213. Jacob, U. and Walther, H. 1981. Aquatic insect larvae as indicators of limiting minimal contents of dissolved oxygen. Aquat. Ins. 3: 219-224. Jannot, J. E., Wissinger, S. A., and Lucas, J. R. 2008. Diet and a developmental time constraint alter life-history trade-offs in a caddis fly (Trichoptera: Limnephilidae). Biol. J. Linn. Soc. 95: 495-504. Jones, J. R. E. 1941. The fauna of the River Dovey, West Wales. J. Anim. Ecol. 10: 12-24. Jones, J. R. E. 1948. The fauna of four streams in the 'Black Mountain' District of South Wales. J. Anim. Ecol. 17: 51-65. Jones, J. R. E. 1949. A further ecological study of calcareous streams in the Black Mountain district of South Wales. J. Anim. Ecol. 19: 142-159. Jones, J. R. E. 1951. An ecological study of the River Towy. J. Anim. Ecol. 20: 68-86. Kamler, E. 1967. Distribution of Plecoptera and Ephemeroptera in relation to altitude above mean sea level and current speed in mountain waters. Polskie Arch. Hydrobiol. 14: 29-42. Knight, A. W. and Gaufin, A. R. 1966. Oxygen consumption of several species of stoneflies (Plecoptera). J. Insect Physiol. 12: 347-355. Knispel, S., Sartori, M., and Brittain, J. E. 2006. Egg development in the mayflies of a Swiss glacial floodplain. J. N. Amer. Benthol. Soc. 25: 430-443. Kolar, C. S. and Rahel, F. J. 1993. Interaction of a biotic factor (predator presence) and an abiotic factor (low oxygen) as an influence on benthic invertebrate communities. Oecologia 95: 210-219. Krno, I. 1990. Longitudinal changes in the structure of macrozoobenthos and its microdistribution in natural and moderately eutrophicated waters of the River Rajcianka (Strázovské vrchy). Acta Fac. Rer. Natur. Univ. Comen. Zool 33: 31-48. Lackey, R. T. and May, B. E. 1971. Use of sugar flotation and dye to sort benthic samples. Trans. Amer. Fish. Soc. 100: 794-797. Langton, P. H. 1998. Micropsectra silvesterae n. sp., and Tanytarsus heliomesonyctios n. sp. (Diptera: Chironomidae), two parthenogenetic species from Ellesmere Island, Arctic Canada. J. Kans. Entomol. Soc. 71: 208-215. Lencioni, V. 2004. Survival strategies of freshwater insects in cold environments. J. Limnol. 63(Suppl.): 145-155. Lillehammer, A. 1966. Bottom fauna investigation in a Norwegian river: The influence of ecological factors. Nytt Magasin for Zoologi 13: 10-29. Lindegaard, C., Thorup, J., and Bahn, M. 1975. The invertebrate fauna of the moss carpet in the Danish spring Ravnkilde and its seasonal, vertical and horizontal distribution. Arch. Hydrobiol. 75: 109-139. Linhart, J., Vlcková, S., and Uvíra, V. 2002. Moss-dwelling meiobenthos and flow velocity in low-order streams. Acta Universitatis Palackianae Olomucensis Facultas Rerum Naturalium (2001-2002) Biologica 39-40: 111-122. Macan, T. T. 1962. Ecology of aquatic insects. Ann. Rev. Entomol. 7: 261-287.

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Maurer, M. A. and Brusven, M. A. 1983. Insect abundance and colonization rate in Fontinalis neo-mexicana (Bryophyta) in an Idaho batholith stream, USA. Hydrobiologia 98: 9-15. McKenzie-Smith, F. 1987. Aquatic bryophytes as habitat for invertebrates in a Victorian upland stream. Unpubl. B.Sc. (Honors) thesis, Monash University, Melbourne Australia, 104 pp. Milner, A. M., Conn, S. C., and Brown, L. E. 2006. Persistence and stability of macroinvertebrate communities in streams of Denali National Park, Alaska: Implications for biological monitoring. Freshwat. Biol. 51: 373-387. Minckley, W. L. 1963. The ecology of a spring stream Doe Run, Meade Co., Kentucky. Wildlf. Monogr. 11: 1-126. Minshall, G. W. 1984. Substratum relationships. In: Resh, V. H. and Rosenberg, D. M. (eds.). The Ecology of Aquatic Insects. Praeger, N. Y., pp. 358-400. Monaghan, M. T., P. Spaak, C. T. Robinson, and J. V. Ward. 2002. Population genetic structure of 3 alpine stream insects: Influences of gene flow, demographics, and habitat fragmentation. J. N. Amer. Benthol. Soc. 21: 114-131. Moon, H. P. 1940. An investigation of the movements of freshwater faunas. J. Anim. Ecol. 9: 76-83. Moore, M. V. and Lee, R. E. Jr. 1991. Surviving the big chill: Overwintering strategies of aquatic and terrestrial insects. Amer. Entomol. 37: 111-118. Muotka, T. 1990. Coexistence in a guild of filter feeding caddis larvae. Do different instars act as different species? Oecologia 85: 281-292. Muttkowski, R. A. 1929. The ecology of trout streams and the food of trout stream insects. Bull. N. Y. State College Forestry, Syracuse Univ. Rossevelt Wild Life Ann. 2: 155240. Muttkowski, R. A. and Smith, G. M. 1929. The food of trout stream insects in Yellowstone National Park. Bull. N. Y. State College Forestry, Syracuse Univ. Rossevelt Wild Life Ann. 2: 241-263. Nagell, B. 1977. Survival of Cloeon dipterum (Ephemeroptera) larvae under anoxic conditions in winter. Oikos 29: 161-165. Nagell, B. and Brittain, J. E. 1977. Winter anoxia – a general feature of ponds in cold temperate regions. Internat. Rev. Gesamt. Hydrobiol. 62: 821-824. Needham, J. G. and Christenson, R. O. 1927. Economic insects in some streams of northern Utah. Bull. Utah Agric. Exper. Stat., Logan, Utah 201: 36 pp. Niesiolowski, S. 1979. Studies on the abundance, biomass and vertical distribution of larvae and pupae of black flies (Simuliidae, Diptera) on plants of the Grabia River, Poland. Hydrobiologia 75: 149-156. Nolte, U. and Hoffmann, T. 1992. Fast life in cold water: Diamesa incallida (Chironomidae). Ecography 15: 25-30. Olsson, T. I. 1981. Overwintering of benthic macro-invertebrates in ice and frozen sediment in a north Swedish river. Holarct. Ecol. 4:161-166. Paavola, R. 2003. Community structure of macroinvertebrates, bryophytes and fish in boreal streams: Patterns from local to regional scales, with conservation implications. Jyväskylä University Printing House, Finland, 35 pp. Pask, W. M. and Costa, R. R. 1971. Efficiency of sucrose flotation in recovering insect larvae from benthic stream samples. Can. Entomol. 103: 1649-1652. Patten, B. C. 1962. Species diversity in net phytoplankton of Raritan Bay. J. Marine Res. 20: 57-75.

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Percival, E. and Whitehead, H. 1926. Observations on the biology of the mayfly Ephemera danica Mull. Proc. Leeds Philosoph. Lit. Soc. Sci. Sec. 1(3): 136-148. Percival, E. and Whitehead, H. 1929. A quantitative study of the fauna of some types of stream-bed. J. Ecol. 17: 282-314. Percival, E. and Whitehead, H. 1930. Biological survey of the river Wharf. II. Report on the invertebrate fauna. J. Ecol. 18: 286-295. Pritchard, G., Harder, L. D., and Mutch, R. A. 1996. Development of aquatic insect eggs in relation to temperature and strategies for dealing with different thermal environments. Biol. J. Linn. Soc. 58: 221-244. Radford, D. S. and Hartland-Rowe, R. 1971. The life cycles of some stream insects (Ephemeroptera, Plecoptera) in Alberta. Can. Entomol. 103: 609-617. Ramløv, H. 2000. Aspects of natural cold tolerance in ectothermic animals. Human Repro. 15: 26-46. Robinson, C. T., Gessner, M. O., and Ward, J. V. 1998. Leaf breakdown and associated macroinvertebrates in alpine glacial streams. Freshwat. Biol. 40: 215-228. Sandberg, J. B. and Stewart, K. W. 2004. Capacity for extended egg diapause in six Isogenoides Klapalek species (Plecoptera: Perlodidae). Trans. Amer. Entomol. Soc. 130: 411-423. Schütz, C., Wallinger, M., Burger, R., and Füreder, L. 2001. Effects of snow cover on the benthic fauna in a glacier-fed stream. Freshwat. Biol. 46: 1691-1704. Shama, L. N. and Robinson, C. T. 2006. Sex-specific life-history responses to seasonal time constraints in an alpine caddisfly. Evol. Ecol. Res. 8: 169-180. Shama, L. N. and Robinson, C. T. 2009. Microgeographic life history variation in an alpine caddisfly: Plasticity in response to seasonal time constraints. Freshwat. Biol. 54: 150-164. Shaw, A. J. and Allen, B. 2000. Phylogenetic relationships, morphological incongruence, and geographic speciation in the Fontinalaceae (Bryophyta). Molec. Phylog. Evol. 16: 225-237. Shelford, V. E. 1945. Conditions of existence. In: Ward and Whipple. Freshwater Biology. John Wiley and Sons, NY, 1115 pp. Steinmann, P. 1907. Die Tierwelt der Gebirgsbache. Ann. Biol. lacustre 2 & Arch. Hydrobiol. Plankt. 3. Stern, M. S. and Stern, D. H. 1969. A limnological study of a Tennessee cold springbrook. Amer. Midl. Nat 82: 62-82. Suren, A. M. 1988. Ecological role of bryophytes in high alpine streams of New Zealand. Internat. Ver. Theor. Angew. Limnol. 23: 1412-1416. Suren, A. M. 1991a. Bryophytes as invertebrate habitat in two New Zealand alpine streams. Freshwat. Biol. 26: 399-418. Suren, A. M. 1991b. Assessment of artificial bryophytes for invertebrate sampling in two New Zealand alpine streams. N. Z. J. Marine Freshwat. Res. 25: 101-112. Suren, A. M. and Winterbourn, M. J. 1992a. Bryophytes as invertebrate habitat in two New Zealand alpine streams. Freshwat. Biol. 26: 327-339. Suren, A. M. and Winterbourn, M. J. 1992b. The influence of periphyton, detritus and shelter on invertebrate colonization of aquatic bryophytes. Freshwat. Biol. 17: 327-339. Tada, M. and Satake, K. 1994. Epiphytic zoobenthos on bryophyte mats in a cool mountain stream, Toyamazawa. Rikusuizatsu. [Jap. J. Limnol.] 55: 159-164.

Teskey, H. J. 1969. Larvae and pupae of some Eastern North American Tabanidae (Diptera). Mem. Entomol. Soc. Can. 63: 1-147. Thienemann, A. 1912. Der Bergbach des Sauerlandes. Internat. Rev. D. Ges. Hydrobiol. Hydrog. Biol. Suppl. 4: 22-71. Thorp, J. H. and Covich, A. P. 1991. Ecology and classification of North American Freshwater Invertebrates. Academic Press, New York, Boston, et al. Thorup, J. 1963. Growth and life-cycle of invertebrates from Danish springs. Hydrobiologia 22: 55-84. Tozer, W. 1979. Underwater behavioural thermoregulation in the adult stonefly, Zapada cinctipes. Nature (Lond.) 281: 566567. Trush, W. J. Jr. 1979. The effects of area and surface complexity on the structure and formation of stream benthic communities. Unpubl. M. S. thesis, Virginia Polytechnic Institute and State University, Blacksburg, VA, 149 pp. Ulfstrand, S. 1968b. Life cycles of benthic insects in Lapland streams (Ephemeroptera, Plecoptera, Trichoptera, Diptera Simuliidae). Oikos 19: 167-190. Verberk, W. C. and Bilton, D. T. 2011. Can oxygen set thermal limits in an insect and drive gigantism? PLoS One 6(7): e22610. Vinson, M. R. and Hawkins, C. P. 1998. Biodiversity of stream insects: Variation at local, basin, and regional scales 1. Ann. Rev. Entomol. 43: 271-293. Vlčková, S., Linhart, J. and Uvíra, V. 2002. Permanent and temporary meiofauna of an aquatic moss Fontinalis antipyretica Hedw. Acta Universitatis Palackianae Olomucensis Facultas Rerum Naturalium (2001-2002) Biologica 39-40. Ward, J. V. 1992. Aquatic Insect Ecology. 1. Biology and Habitat. John Wiley & Sons, Inc., N. Y., 438 pp. Wesenberg-Lund, C. 1943. Biologie der Süßwasserinsekten. Byldendalske Boghandel. Nordisk Forlag, Kopenhagen, 682 pp. Whitehead, H. 1935. An ecological study of the invertebrate fauna of the chalk stream near Great Driffield, Yorkshire. J. Anim. Ecol. 4: 58-78. Williams, D. D. 1980. Some relationships between stream benthos and substrate heterogeneity. Limnol. Oceanogr. 25: 166-172. Wingfield, C. A. 1939. The function of the gills of mayfly nymphs from different habitats. J. Exper. Biol. 16: 363-373. Wise, D. H. and Molles, M. C. Jr. 1979. Colonization of artificial substrates by stream insects: Influence of substrate size and diversity. Hydrobiologia 65: 69-74. Wodsedalek, J. E. 1911. Phototactic reactions and their reversal in the mayfly nymphs Heptagenia interpunctata (Say). Biol. Bull. Mar. Biol. Lab., Woods Hole 21: 265-271. Wulfhorst, J. 1994. Selected faunal elements of the hyporheos and in submerged moss clumps (bryorheal) along an acidification gradient in two brooks in the Harz Mountains, West Germany. Internat. Verein. Theoret. Angew. Limnol. Verhand. 25: 1575-1584. Zwick, P. 1996. Variable egg development of Dinocras spp. (Plecoptera, Perlidae) and the stonefly seed bank theory. Freshwat. Biol. 35: 81-100.

Glime, J. M. 2017. Aquatic insects: Bryophyte roles as habitats. Chapt. 11-2. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. Ebook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 19 July 2020 and available at .

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CHAPTER 11-2 AQUATIC INSECTS: BRYOPHYTE ROLES AS HABITATS TABLE OF CONTENTS Potential Roles .................................................................................................................................................. 11-2-2 Refuge ............................................................................................................................................................... 11-2-4 Habitat Diversity and Substrate Variability ...................................................................................................... 11-2-4 Nutrients..................................................................................................................................................... 11-2-5 Substrate Size ............................................................................................................................................. 11-2-5 Stability ...................................................................................................................................................... 11-2-6 pH Relationships ........................................................................................................................................ 11-2-6 Bryophyte Structure ................................................................................................................................... 11-2-9 Scapania undulata .............................................................................................................................. 11-2-9 Hygroamblystegium spp...................................................................................................................... 11-2-9 Platyhypnidium riparioides .............................................................................................................. 11-2-10 Fissidens grandifrons........................................................................................................................ 11-2-10 Fontinalis spp. .................................................................................................................................. 11-2-10 Flow Regimes ................................................................................................................................................. 11-2-11 Flow Rates ............................................................................................................................................... 11-2-12 Overturned Rocks .................................................................................................................................... 11-2-12 Life History and Flow .............................................................................................................................. 11-2-12 Water Level ..................................................................................................................................................... 11-2-13 Stream Drift..................................................................................................................................................... 11-2-13 Safe Sites ......................................................................................................................................................... 11-2-18 Biomass and Richness ..................................................................................................................................... 11-2-20 Food Sources ................................................................................................................................................... 11-2-21 Bryophytes as Food.................................................................................................................................. 11-2-23 Nutritional and Antifeedant Properties ............................................................................................. 11-2-24 Tracing Bryophytes in the Food Chain ............................................................................................. 11-2-26 Food when Food Is Scarce ....................................................................................................................... 11-2-28 Epiphytes and Meiofauna of Bryophytes ................................................................................................. 11-2-29 Trapping Detritus ..................................................................................................................................... 11-2-30 Detrimental Effects?........................................................................................................................................ 11-2-32 Bryophytes vs Tracheophytes ......................................................................................................................... 11-2-32 Summary ......................................................................................................................................................... 11-2-38 Acknowledgments ........................................................................................................................................... 11-2-39 Literature Cited ............................................................................................................................................... 11-2-39

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CHAPTER 11-2 AQUATIC INSECTS: BRYOPHYTE ROLES AS HABITATS

Figure 1. Habitat for stream bryophyte dwellers, Wolf Brook, NY, USA. Photo by Jason Neuswanger, with permission.

Potential Roles Ulfstrand (1967) astutely stated that aquatic insects select their habitats on the basis of factor combinations. While some minimal levels of factors are important – oxygen, temperature, space, stability – the most important factor determining location within this medley of minimums is usually food. And that food works in two directions: enough food to maintain nutrition and avoidance of becoming food themselves. To satisfy both food factors, Ulfstrand found that substrate is especially important; bryophytes are often important choices among those substrates. Bryophytes are major components in several types of ecosystems, including peatlands, mountain streams (Figure 1), high latitudes, and boreal forest floor. Many researchers have found that bryophytes are important substrata for insects (Percival & Whitehead 1929). Arnold and Macan (1969) found the greatest species richness and number of individuals among mosses, citing their role as cover and source of food by trapping particles.

Bryophytes, both mosses and liverworts, often form extensive cover in rocky and stony reaches of streams (Macan & Worthington 1951). These can have profound effects on the fauna by providing footholds against the current. Mosses with moderate thickness are suitable for the mayflies Baetis (Figure 2) and Ephemerella (Figure 3) and Plecoptera (stoneflies; Figure 20). Fish benefit as well, with the greatest production of fish-food organisms where there are either rooted plants or mosses. For example, Chironomidae (Figure 9) are in greatest numbers among thick mosses. And fish certainly eat Chironomidae (Mousavi et al. 2002). Based on gut contents, Frost (1939) considered moss-dwelling insects to be an important constituent of the diet of trout (Frost 1939) and young salmon (Frost & Went 1940) in the River Liffey, Ireland. Likewise, Minnows appear to crop the moss fauna (Frost 1942). On the other hand, Brusven et al. (1990) found that at least in the daytime when salmonid fish feed, the insects drifting in the moss-covered channel (Fontinalis neomexicana – Figure 4) did not provide any greater biomass for fish food than in channels where mosses were absent and insect faunal density was much less. Bowden et

Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

al. (1999) likewise questioned whether fish actually benefit from the increased abundance of insects in streams where bryophytes are present, citing a lack of evidence.

Figure 2. Baetis rhodani on sand, a mayfly that also lives among mosses. Photo by J. C. Schou, with permission.

Figure 3. Ephemerella dorothea on moss (Platyhypnidium riparioides) in Virginia, USA. Photo by D. N. Bennett, with permission.

Figure 4. Fontinalis neomexicana, a slightly amphibious species that provides shelter for moss dwellers. Photo by Belinda Lo, through Creative Commons.

I am aware of no study that demonstrates quantitatively that the increase in number of insects in moss mats benefits fish. It appears that insects may have evolved to drift at night precisely to avoid predation by day-feeding

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fish. Bryophytes are a difficult place for fish to locate and catch the insects, perhaps reducing the catchable food from what might have been available if rock-dwelling insects were present instead. The hypothesis that bryophyte dwellers increase available fish food needs to be tested. Corona (2010) suggested that immature insects in streams stayed together because that behavior would increase survival, a concept already suggested for vertebrates by Elgar (1986), Robinette et al. (1995), and Brown and Brown (2004). Bryophytes that provide a stable, protected habitat would facilitate such behavior. Nearly fifty years after Macan and Worthington (1951) expressed the profound contribution of bryophyte-dwelling insects, Bowden et al. (1999) summarized that bryophytes "can profoundly influence both the abundance and community structure of stream invertebrates." But they further stated that "the number of fundamentally important roles of bryophytes in stream ecosystems remain unexamined." I will attempt to pull together what various scattered studies around the world have revealed about the roles of stream bryophytes. Paddling a Kayak to gain first-hand information, Yamamura (2009) observed the adaptations of aquatic insects to various flow regimes in the rivers of Idaho, following up on studies by Rosentreter (1984). In their studies, Yamamura and Rosentreter found that aquatic insects benefit by having aquatic bryophytes because: 1. Bryophytes decrease stream velocity on the rock’s surface layer. 2. Bryophytes trap more detritus (Figure 5; product of disintegration, especially organic matter produced by the decomposition of organisms) than smooth rock (food for shredder insects). 3. Bryophytes provide hiding cover (refuges) from predators. 4. Bryophytes provide better background coloration for camouflage. 5. Bryophytes provide greater surface area, providing a greater amount of habitat area. 6. Bryophytes provide more food since algae can grow upon the greater surface area created by the three dimensions of the moss surface. 7. Bryophytes provide greater algae retention and protection when stream flow regimes are low enough to create dry surfaces. The bryophytes retain water longer than other substrata in the stream, permitting the algae to dry slowly and acclimate to the encroaching desiccation. 8. Perennial bryophytes such as Scouleria aquatica (Figure 6) can provide long-term stability to an ephemerally dry rock surface, permitting survival of algae, insect larvae, and eggs. Yamamura (2009) concluded that insect larval data support the interpretation that larvae in spring-fed streams (streams containing aquatic moss) are larger compared to those in runoff-dominated streams (streams that lacked mosses). He concurred with Rosentreter (1984) that springfed (mossy) streams have three cohorts present while most run-off (non-mossy) streams have two cohorts. This raises the question, do mosses in runoff-dominated streams benefit insects enough to produce larger larvae and another generation (cohort) per year? Perhaps the insects benefit

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Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

from the added cover of bryophytes – insect predators in Idaho streams include other insects, fish, shore birds, and the American dipper. Mosses provide cover in which to hide from all these predators.

positive effects on the size and fecundity of the adult caddisflies by reducing competition among the larvae through predation. The striking revelation of this study was that despite the detritus-based diet of these caddisflies, reduction in the number of larvae still had a positive effect on the adults of the species when compared to those in fishless streams. The adults were larger and the females had 33% more eggs, but the egg size was unchanged. Nevertheless, the increase in number of eggs did not compensate for the loss of larvae. The study by Greig and McIntosh (2008) suggests that fish have an impact on insects that typically live among the bryophytes, many of whom are detritus feeders. Thus, the bryophyte cover potentially increases the number of insects surviving and the number of adults reproducing, but we are left with the question of whether the bryophytes ultimately produce more available fish food.

Habitat Diversity and Substrate Variability Figure 5. Detritus, a common food for aquatic insects and typically accumulated at plant and leaf bases among bryophytes. Photo by James K. Lindsey, with permission.

Figure 6. Scouleria aquatica on rock near stream water. Photo by Matt Goff, with permission.

Refuge Bryophytes serve as refuges in both moving water and lentic systems such as lakes and ponds. In streams, they provide a refuge against the torrents of rapidly flowing water, permitting insects to live where they can take advantage of the higher oxygen and suspended food sources available in flowing water while remaining safely anchored within the moss or clinging to its surface. In both habitats, the bryophyte provides a hiding place from predators, especially fish, but also larger insects, crayfish, and birds. The importance of bryophyte-dwelling insects as fish food is a subject for speculation. While the bryophytes provide homes for numerous insects, there is no direct evidence that these insects are available as increased fish food. Greig and McIntosh (2008) examined the effect of brown trout (Salmo trutta) predation on the caddisfly Zelandopsyche ingens, a bryophyte dweller in New Zealand. They determined that these trout can have

Habitat diversity offers more niches, hence making the area suitable for more species. Clenaghan et al. (1998) identified ecological factors that contribute to macroinvertebrate community composition. Local ecological factors include acidic water, moss, shading, agricultural runoff, longitudinal trends in stream physicochemistry (distance from headwaters, geology, land use) and season (related to life history patterns of the invertebrates). In their study of a conifer-afforested catchment in Ireland, macroinvertebrate density and richness increased with the distance from the headwaters and the concomitant increases in pH, water hardness, and available nutrients. Douglas and Lake (1994) demonstrated that habitat diversity was important in increasing species richness in streams. Bryophytes not only add to that diversity, but increase available surface area. Based on a review of the literature, Smith-Cuffney (1987) reported that stream mosses in low order, high elevation streams have a structurally unique community. Measured as respiration rates, the communities among Fontinalis (Figure 4) had three times the rates found in the stone community and five times that of the hyporheic community. Arnold and Macan (1969) found the largest number of species and individuals of insects inhabited mosses in a Shropshire Hill stream in the UK, where the mosses provided both shelter and trapped food. Pardo and Armitage (1997) demonstrated the importance of environmental variables in the spatial distribution of aquatic insects based on eight mesohabitats. They found that water velocity and flow dynamics, together with the nature of the substrate were the major determinants of benthic (bottom) communities. Heino (2009) looked at the environmental variables somewhat differently, attempting to explain why such things as the influence of altitude varied with geography. He found pH, stream size, and moss cover were the most important variables, with functional diversity increasing with moss cover. These two approaches are not that different, with pH and water velocity both influencing moss cover and moss cover providing safe sites in areas of high flow rates. Špoljar et al. (2012) likewise found that flow velocity and pH had the greatest effect on community structure. In two springs in Papuk Nature Park, Croatia, the

Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

macroinvertebrate taxa numbered only 25. Where the bryophyte cover was dense (90% cover), the community structure was most affected by flow velocity and pH; macroinvertebrate diversity and abundance were higher than in the stream with only 50% bryophyte cover. In the latter stream, algae, protozoa, and meiofauna (minute inimals living in small spaces in soil or aquatic sediments) reached higher abundance, apparently resulting from suspended organic matter and epiphytes. Bryophyte communities exemplify the species-area relationship (Gleason 1922). Increased bryophyte cover means an increase in available substrate due to its threedimensional structure. Heino and Korsu (2008) found a strong relationship between species richness and number of individuals, and both of these were significantly related to the bryophyte biomass. They attributed the relationship to the increased cover provided by greater bryophyte coverage. Heino et al. (2005) found that despite the highest congruence between bryophytes and macroinvertebrates among the stream biological groups, that congruence was nevertheless weak. This seems to relate to differences in the stream factors that determine bryophyte locations. Bryophyte diversity followed water color, habitat stability, and stream size, in that order. Macroinvertebrate diversity instead was determined in the order of stream size, water color, and acidity.

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Figure 7. Hygrohypnum alpinum, home of many aquatic insects. Photo by Michael Lüth, with permission.

Nutrients Nutrients can affect moss growth in some cases and limit it due to competition for light by encrusting algae in others. In their study of the Kuparuk River, Alaska, USA, Lee and Hershey (2000) found that fertilization with phosphorus increased the growth of mosses (Hygrohypnum – Figure 7), but that insects did not respond as extensively as one might expect. Invasion by mosses resulted in an increased density of the mayfly Ephemerella aurivillii (Figure 8) and Chironomidae (midges; Figure 9), but had no effect on densities of the mayfly Baetis spp. (Figure 2) or Simuliidae (blackflies; Figure 22). Both Baetis and Ephemerella grew larger in fertilized areas, but Lee and Hershey suggested that this was most likely due to the increase in epiphytic diatoms. Only Ephemerella seemed to be affected by substrate type (bare rock, natural moss, artificial moss), with the greatest densities among the mosses, presumably due to increased habitat complexity. Clenaghan et al. 1998) compared several factors and found that mosses were one of the factors explaining the diversity of insects in a catchment stream in Ireland, and that both density and richness increased with moss weight. Voelz and McArthur (2000) likewise concluded that habitat complexity was one of the most important factors in determining species richness in streams. In my own culturing studies, I have found that enrichment was often detrimental to the mosses. These mosses lost their green color and were covered by algae that presumably intercepted the light – and CO2. While the bryophytes remained intact, even if dead, this enrichment could benefit the insects by increasing food sources, but such enrichment most likely would make establishment of new mosses or increased coverage by existing ones less likely.

Figure 8. Ephemerella aurivillii naiad, a species whose density increases when there are mosses. Photo by Tom Murray, through Creative Commons.

Figure 9. Chironomidae larva, an insect that increases in abundance when greater moss growth occurs. Photo by Bob Henricks, with permission.

Substrate Size The biodiversity of macroinvertebrates typically increases linearly with the substrate suitability index [suitability of sediment, periphyton (freshwater organisms attached to or clinging to plants, but also used to include other objects projecting above the bottom sediments; Aufwuchs), and benthic organic materials] (Duan et al. 2009). In large rivers in China (Yangtze River, Yellow River, East River, Juma River), Duan et al. found that the macroinvertebrate community was not dependent upon macroclimatic conditions or latitude, but rather responded to the commonality of instream habitat conditions of substrate composition and flow conditions in these rivers.

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Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

They found that taxa richness was highest on cobble covered with hydrophytes, high on moss-covered bedrock, and low on clay or cobble where there were no plants. Sandy beds were unstable and thus devoid of benthic macroinvertebrates. As in many stream studies, the EPT insects [Ephemeroptera (Figure 8, Plecoptera (Figure 20), Trichoptera (Figure 13)] dominated the cobble, gravel, and moss-covered bedrock. But contrasting with most stream studies (see Chapter 11-9, Holometabolous Insects – Diptera), the Chironomidae larvae (Figure 9) reached greatest dominance in the clay beds. But substrate size apparently does not act alone and importance differs among types of insects (see for example Ulfstrand 1967). Contrasting with other studies, Wise and Molles (1979) found that small substrates supported more insect individuals than did the larger stones. And mixed sizes supported numbers between the small and large sizes.

(Figure 13) was absent at sites with S. undulata and N. compressa, but present in streams with Fontinalis squamosa (Figure 14).

Stability I love the expression "A rolling stone gathers no moss," because it so perfectly describes the situation of stability. This expression can be traced to Erasmus' Adagia, first published around 1500, and has since taken on wide usage with somewhat conflicting interpretations. Nevertheless, in the context of a stream, its meaning is clear. Bryophytes themselves indicate a stable substrate (Yamamura 2009). Such stable areas are present due to stream channel geometry. Rapids can focus the ice scraping at the center of the river, away from the sides where bryophyte populations are able to grow. Hence, some invertebrates may live in those mossy areas simply because they, too, only survive where the substrate is stable and the water has a reduced shearing effect. Stability is most important for eggs and many pupae that cannot move to a more favorable location when the need arises. Bryophytes will only become well established on stable rocks and boulders, so they signal a stable habitat. Furthermore, as water levels recede, bryophytes maintain water content well beyond the time that a rock can do so, creating a moisture stability. And when the young insects hatch from the eggs, these tiny animals are not only easy prey for larger animals, but they are poor swimmers unable to navigate in the flowing water. The bryophytes provide cover and protection in their small-chambered labyrinth that prevents entry to predators such as fish and large insects and that reduces the flow to near-pool conditions (Glime 1978).

Figure 10. Scapania undulata, a leafy liverwort that can serve as food for the mayfly Ecdyonurus. Photo by Michael Lüth, with permission.

Figure 11. Nardia compressa, a leafy liverwort that can be eaten in some streams by the mayfly Ecdyonurus sp. Photo by Des Callaghan, with permission.

pH Relationships The depauperate (lacking in numbers or variety of species) fauna of some bryophytes may relate more to the preferred habitats of the bryophytes than to the bryophytes themselves. For example, in Wales, Ormerod et al. (1987) found that in streams with low pH the bryophytes [liverworts Scapania undulata (Figure 10) and Nardia compressa (Figure 11)] had few insects; 60% of the S. undulata sites had fewer than 20 macroinvertebrate taxa. The pH where Ormerod et al. found these liverworts growing was 5.2-5.8. On the other hand, less than 5% of the sites with the red alga Lemanea (Figure 12) (pH 5.58.5) were so impoverished. In particular, Hydropsyche

Figure 12. Lemanea sp. covered with blackflies. Photo by Janice Glime.

Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

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Other factors may affect the choices of many insects to avoid colonizing these acid-tolerant bryophytes. For example, one possibility that Ormerod et al. (1987) considered was that the diatom Eunotia (Figure 16) that grows in the leaf axils of leafy liverworts (acid-loving) is inaccessible to grazing Baetis (Figure 2), whereas the diatom Cocconeis (Figure 17) grows on the leaf lamina of the moss Hygrohypnum (Figure 7; growing at a higher pH) where it is easily grazed (Sutcliffe et al. 1986).

Figure 13. Hydropsyche larva, a net-spinning caddisfly that frequents Fontinalis antipyretica (Figure 18) and Platyhypnidium riparioides. Photo by Guillaume Doucet , with permission.

Figure 16. Eunotia sp., a diatom that grows in leaf axils of leafy liverworts where Baetis is unable to reach it. Photo by Janice Glime.

Figure 14. Fontinalis squamosa above and below water on rocks, home to several stonefly genera. Photo by Janice Glime.

Research by Willoughby and Mappin (1988) suggests that the insect avoidance of the two leafy liverworts that Ormerod et al. (1987) observed may not have been a response to pH, but rather the result of the liverwort terpenes and terpene alcohols in the oil bodies. On the other hand, some insects such as the mayfly Ecdyonurus (Figure 15) feed on such acid-tolerant bryophytes as S. undulata (Figure 10), but are unable to live in the acid streams at the lower end of the pH tolerance range of this liverwort. Ormerod and coworkers (1987) considered that these mayflies are therefore physiologically restricted from acid streams.

Figure 15. Ecdyonurus venosus naiad, a mayfly genus in which some members feed on Scapania undulata (Figure 10) when the pH is not too low. Photo by Guillaume Doucet , with permission.

Figure 17. Cocconeis placentula, an epiphytic diatom that cements itself to aquatic bryophyte leaves. Photo by Ralf Wagner at , with permission.

Heino (2005) likewise found that functional richness of macroinvertebrates increased with increased pH, with total nitrogen, water color, and substrate particle size also varying with moss cover in 111 boreal headwater streams in Finland. The functional structure depended on these same variables with its dominant pattern being related to increase of shredder-sprawlers and decrease of scraperswimmers in acidic conditions. Frost (1942) compared the fauna on the mosses in acid and alkaline streams in her survey of River Liffey, Ireland. Chironomidae (Figure 9) constituted 40-54% of the fauna in these streams. In the carboniferous limestone sites,

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Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

Fontinalis antipyretica (Figure 18) and Platyhypnidium riparioides (Figure 19) dominated in a pH range of 7.4 to 8.4. The stonefly fauna of these mosses was comprised of predominantly Isoperla (Figure 20). The dominant caddisfly genus was Hydropsyche (Figure 13). Mayflies included Ephemerellidae (Figure 8) (mean 533 per sample of 200 g wet weight), Baetis (Figure 2), and Caenis (Figure 21). The blackfly Simulium (Figure 22) was common. In the acid streams (peat bog drainage), the pH ranged 4.4-6.8, and the bryophytes were dominated by Fontinalis squamosa (Figure 14) with a small coverage by the leafy liverwort Scapania undulata (Figure 10). The stonefly fauna was comprised of Protonemura (Figure 104), Amphinemura (Figure 105), Leuctra (Figure 49), and Chloroperla (Figure 23). Polycentropus (Figure 24) was the predominant caddisfly.

Figure 18. Fontinalis antipyretica, home to the stonefly Isoperla and net-spinning caddisfly Hydropsyche. Photo by Andrew Spink, with permission.

Figure 21. Caenis youngi naiad, member of a genus that sometimes inhabits Fontinalis antipyretica (Figure 18) and Platyhypnidium riparioides. Photo by Bob Newell, with permission.

Figure 22. Simulium (blackfly) larvae showing the large numbers that can occupy one rock – or moss. Photo by F. Christian Thompson, through USDA public domain.

Figure 19. Platyhypnidium riparioides, home to the stonefly Isoperla and net-spinning caddisfly Hydropsyche. Photo by Andrew Spink, with permission.

Figure 23. Chloroperlidae naiad, a detritus inhabitant, including mosses. Photo by Bob Henricks, with permission.

Figure 20. Isoperla similis naiad, member of a genus that inhabits Fontinalis antipyretica (Figure 18) and Platyhypnidium riparioides. Photo by Donald S. Chandler, with permission.

Figure 24. Polycentropus larva, a dominant caddisfly among Fontinalis in acid streams. Photo by Jason Neuswanger, with permission.

Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

In a similar study, Willoughby and Mappin (1988) found that growth of the mayfly Serratella ignita (Figure 25) was similar when fed on food from acid or alkaline streams. In acid streams they fed on the leafy liverwort Nardia compressa (Figure 11) with the filamentous alga Klebsormidium subtile (Chlorophyta; see Figure 26), whereas in the alkaline streams they ate the moss Platyhypnidium riparioides (Figure 19) with the epiphytic diatom Cocconeis placentula (Figure 17). But if the alga Klebsormidium subtile was absent in the acid streams, they were unable to subsist on the liverworts alone.

Figure 25. Serratella ignita naiad, a mayfly species that can subsist in both acid and alkaline streams, feeding on bryophytes and associated algae. Photo by J. C. Schou, with permission.

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Scapania undulata This is a leafy liverwort whose chemical components of terpenoids have already been mentioned. Its growth form is somewhat layered (Figure 27), and its leaves are conduplicate (Figure 28). That is, the leaf is folded over so that the smaller portion is on top. This fold provides a protected area where several small insects such as the stoneflies Leuctra (Figure 49) and Nemoura (Figure 40) like to hide (Glime 1968). Its layered effect makes it somewhat more open to the water, permitting predators to penetrate more deeply in search of prey, a problem that is avoided by the small insects that can hide within the folds of the leaves.

Figure 27. Scapania undulata showing layered effect. Photo by Hermann Schachner, through Creative Commons.

Figure 28. Scapania undulata showing folded leaves with smaller lobes on top. Photo by Florent Beck, through Creative Commons. Figure 26. Klebsormidium flaccidum, a green alga associated with Nardia compressa in acid streams, providing food for Serratella ignita. Photo by Sarah Kiemle, with permission.

Bryophyte Structure Not all bryophytes are created equal, despite their frequent treatment as one entity in ecological studies. Their structures can differ greatly, and this has a strong influence on which organisms can live there. This structure is seldom considered in describing the habitat and the influences of the bryophytes on the inhabitants. Let's consider a few and the differences they offer.

Hygroamblystegium spp. This genus, including Hygroamblystegium fluviatile and H. tenax, forms thick mats on rocks (Figure 29). Its extensive branching provides an array of spaces within the mat, affording protection from both the current and most larger insects and fish. The leaf has a strong costa (Figure 30) that is used by some caddisflies in the construction of their cases (to be discussed later in the Trichoptera subchapter). Its small leaves and branches afford small spaces unavailable to larger insects, thus limiting the species and life stages that can live there.

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Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

Figure 29. Hygroamblystegium tenax in a dry stream bed. Photo by Janice Glime.

Figure 30. Hygroamblystegium fluviatile showing cupped leaves and strong costa used by some caddisflies in construction of their cases. Photo by Hermann Schachner, with permission.

Platyhypnidium riparioides

Figure 31. Platyhypnidium riparioides, home to many kinds of aquatic insects. Photo by Michael Lüth, with permission.

Figure 32. Platyhypnidium riparioides, showing leaves where many kinds of insects are able to hide. Photo by John Hribljan, with permission.

This species occurs in many of the same streams as those of Hygroamblystegium fluviatile (Figure 30). It is a widespread species that forms a chambered mat. It has somewhat larger leaves than H. fluviatile but creates a similar habitat with many species in common. It is not unusual to find these two species on the same rock, often intermixed. Platyhypnidium riparioides (Figure 31-Figure 32) affords somewhat larger spaces within the mat. Its costa is reduced and much thinner than that of Hygroamblystegium species and does not seem to be particularly useful for case building. Fissidens grandifrons Fissidens grandifrons (Figure 33) tends to prefer alkaline streams. It is a large moss with flat branches that are layered somewhat like those of Scapania undulata (Figure 27-Figure 28), an inhabitant of acid streams. It occurs in very cold water and waterfalls, both conditions that provide it access to more CO2 than would be available in un-aerated warmer water. I never searched this moss for insects, but my collections of it did not reveal any conspicuous fauna. It is a stiff moss and its preference for torrential water may discourage them.

Figure 33. Fissidens grandifrons showing the flat branches and accessible spaces between them. Photo by Janice Glime.

Fontinalis spp. Fontinalis species are large mosses (Figure 34). They have a streamer growth form in which all stems dangle in the same direction as the flow of water, at least where there is a distinct flow. The end portions of the stems are

Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

exposed, harboring Simuliidae. The leaf structure varies among species, thus providing differing suitability for the insects. Fontinalis antipyretica (Figure 35) has large, keeled leaves that form a 3-sided branch with well protected interior space. However, this space may be somewhat difficult for many insects to enter due to the close appression (state of being pressed close to) of leaves. Fontinalis hypnoides (Figure 36) has narrow, more or less flat leaves that do not provide much enclosed space. In between these two extremes are various degrees of enclosure and access to that enclosure. The flat surface of the branch of F. antipyretica would be ideal for blackfly larvae, but this Fontinalis species is often not successful in the very fast flow needed by these larvae. If the moss is in fast flow, the keel is easily worn away and the leaves become tattered. However, in cool streams there is usually sufficient oxygen for both the moss and blackflies to survive.

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Figure 36. Fontinalis hypnoides showing flattened, narrow leaf. Photo from Dale A. Zimmerman Herbarium, Western New Mexico University, with permission.

Fontinalis squamosa (Figure 37), a European species, is one of the several intermediate species. Its leaves are concave and provide hiding places within the concavities. Like all Fontinalis species, it lacks a costa. This species has been indicated as home to numerous insects in many European stream studies.

Figure 34. Fontinalis dalecarlica, a refuge for invertebrates during low water levels. Photo by Kristoffer Hylander, with permission. Figure 37. Fontinalis squamosa showing concave leaves. Photo by Michael Lüth, with permission.

Flow Regimes

Figure 35. Fontinalis antipyretica demonstrating the folded, overlapping leaves that give little accessibility to the interior leaf space. Photo by David T. Holyoak, with permission.

Flow regimes provide another limitation for bryophyte inhabitants. Many bryophytes live in areas of high flow that is too abrasive for the establishment of tracheophytes (plants with lignified vascular tissue, i.e., all plants that are not bryophytes). At the same time, many insects require protection from the rapid flow. Furthermore, insects drift in streams for various reasons – searching for food, making a false move that puts them in the current, overpopulation, finding a site for pupation, and dislodgment due to changes in flow. Baker et al. (1996) found that the hydraulic stability of streams over multiple years determined whether a site was dominated by periphyton, bryophytes, or tracheophytes. Variations within the year can control periphyton biomass, with low velocities favoring both periphyton and tracheophytes that serve as additional substrate for them. Bryophytes, on the other hand, are often restricted to areas of high velocity; these same high velocities restrict colonization and accumulation of detritus.

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Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

Bryophytes modify the internal flow of water. The arrangement of sedimentary deposits and fauna below the leaves of submerged stream bryophytes supports this concept of internal current modification (Devantery 1995). Using Platyhypnidium riparioides (Figure 19) and colored liquid, Devantery was able to demonstrate that a single leaf of this moss caused symmetrical twirling behind it. Between the leaves he observed a retrocurrent in the direction of the leaf. This current was slowed progressively and directed the water toward the leaf insertion, explaining the accumulation of detritus there. The same hydrodynamics also occurred in a second species of bryophyte that had a different leaf morphology. Certain insects take advantage of refugia, especially during periods of high flow (Lancaster & Hildrew 1993). Bryophytes are able to provide such refugia and are likely to be especially important for such species as Nemurella pictetii (Figure 38) and larger naiads of Leuctra nigra (Figure 39), both stoneflies known from bryophytes. Lancaster and Hildrew found that seasonal flow conditions affected the distribution of these two species in streams after high-flow events, but that these seasonal differences in flow seemed to have little effect on the Chironomidae or the young instars (instar is developmental stage between molts of an insect) of Leuctra nigra.

Figure 38. Nemurella pictetii naiad, a species that uses bryophytes as refugia. Photo by Urmas Kruus, with permission.

Macan and Worthington (1951) suggested that mosses can "profoundly influence the fauna by providing a foothold for animals which otherwise could be swept away by the current." Devantery (1987) reminds us of the importance of flow in contributing to the accumulation of food resources in the bryophyte mat. With regard to the moss Platyhypnidium riparioides (Figure 19), Devantery considers that the moss increases the spatial uniformity, a perspective that seems to be in contrast with those who consider the moss to increase the complexity of the habitat (Dražina et al. 2011). The flow serves as an antagonist with the danger that it can dislodge the bryophytes. Flow rates approaching the bryophytes influence the insects that make those bryophytes home. The Chironomidae (Figure 9) are reduced by higher flow velocities associated with Fontinalis antipyretica (Figure 18), whereas the smallest of the Simuliidae larvae (Figure 22) are positively influenced (Linhart et al. 2002a, b). This may relate to available food, with the Simuliidae trapping fine particles with their head fans and Chironomidae living among the detritus that has been trapped by the moss. Overturned Rocks The famous statement, "a rolling stone gathers no moss," applies in its literal sense as well as the figurative. Bryophytes cannot grow under an overturned rock, and rolling is abrasive, damaging new stems and knocking off older clumps. For stream ecosystems, these dangers prevail. Englund (1991) found that 16.7% of the mosscovered stones in North Swedish woodland streams had been overturned in the last few years. Small stones rarely had mosses (See also Slack & Glime 1985), a factor most likely related to their instability. But when stone size exceeded more than 12 cm, mosses were abundant even on rocks that were not embedded into the substrate. Englund (1991) experimented on the effects of overturning not only on the mosses, but also on their invertebrate fauna. Overturning, as expected, reduced both diversity and abundance of fauna as well as reducing the dry weight of mosses. Nevertheless, 3 out of 16 invertebrate taxa increased, predominantly on the mosscovered underside. For the remaining taxa, peak densities occurred on the upper moss-covered sides of control stones, and these densities decreased on the overturned stones. Despite the introduction of insects through stream drift (see below), recovery was still weak 14 months later, probably because of the slow recovery of the mosses. Life History and Flow

Figure 39 Leuctra nigra naiad, a species that uses bryophytes as refugia. Photo by J. C. Schou, with permission.

Flow Rates One possible role of bryophytes as a habitat for insects and other invertebrates is their ability to provide a refuge with multiple current velocities (Madaliński 1961; Elliott 1967a; Gurtz & Wallace 1984; Suren 1992a, b; Glime 1994). Hence, organisms can migrate within the bryophyte mass to locate the current velocity that meets their needs.

For insects living in streams, the habitat is likely to be too fast at times and too dry at others. Yamamura (2009) concluded that the variability of the flow regime can limit the distribution and the life history traits of aquatic insects. Some have solved this transient habitat problem by life cycle stages that either are dormant or that do not require water. Among these, the egg stage is a suitable stage for surviving drought in some stoneflies, mayflies, and dipterans (Ward 1992). In the case of the stonefly Nemoura (s.l.) (Figure 40), a common moss dweller, in a Welsh stream, the adults emerge at the end of the drought (Hynes 1958; Ward 1992). In their short adult life stage,

Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

they may take advantage of newly formed pools in the stream for oviposition before the stream returns to normal flow.

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invertebrates for food. But those that feed on periphyton, and most likely on high quality detritus, may depend on the chambered bryophyte clumps for their dinner. Fortunately, these bryophytes help to provide both hydration and food for herbivores and detritus feeders. As the water level decreases, bryophytes can act like a filter to trap detrital matter from the slow water. In a Québec, Canada, stream, Cattaneo et al. (2004) found that many of the invertebrates moved to or remained among mosses (Fontinalis dalecarlica; Figure 34) at low water levels. Water depth explained 50-80% of the variation in the invertebrate biomass among the mosses and the biomass was lower on shallow mosses that had more frequent exposure. Grazers were more common in the moss habitat than in the gravel, but carnivores such as Plecoptera and Odonata were in the gravel.

Stream Drift Figure 40. Nemoura naiad, a common bryophyte dweller. Photo by Bob Henricks, with permission.

Mosses may often play an important role in providing moist sites for the aquatic insects during fluctuating conditions, but their role at such times has scarcely been investigated. In a Welsh mountain stream, severe flooding transported large quantities of gravel (Hynes 1968). Gravel-dwelling insects were greatly reduced, and the moss cover was reduced by 80%. But the fauna living among the remaining mosses was not significantly decreased. The stoneflies, caddisflies, and Elmidae (riffle beetles; Figure 41) recolonized the area before any reproduction could have contributed to their recovery. Hynes hypothesized that these insects migrated to deep within the benthic zone (away from abrasion) during the flood and then reappeared after the water level returned to normal.

Figure 41. Elmidae adult, a rapid colonizer of bryophytes. Photo by Stephen Moore, Landcare Research, NZ, with permission.

Water Level Water level changes bring problems of not only hydration but also food availability for aquatic insects. Open-water carnivores can easily move and will most likely still have access to smaller insects and other

Stream drift is a natural occurrence among stream fauna, especially insects (Anderson & Lehmkuhl 1968). Waters (1972) emphasized that this is an episodic event and not a continuous phenomenon. The drift organisms are bottom and vegetation organisms. When stream discharge is reduced by seasonal events, catatrostrophic drift can occur. Two primary organisms in such drift in Oregon, USA, are Simulium sp. and Baetis tricaudatus, both bryophyte dwellers (Corrarino & Brusven 1983). Catastrophic drift (Minckley 1964) occurs from a physical disturbance such as flooding, anchor ice (ice anchored to bottom) (O'Donnell & Churchill 1954), pollution (Coutant 1964), drought, and high temperatures (Wojtalik & Waters 1970; Reisen & Prins 1972 for Simulium - Figure 22). Behavioral drift occurs at a particular time of day or night; it may result from crowding, competition, need for food, predation, making a new case, or attempting to reach land at emergence time (Waters 1972). Constant drift is comprised of small numbers that are always present as organisms move about and become dislodged from their substrates (Waters 1972). Most drift occurs at night (Bishop 1969; Elliott 1965, 1968; Holt & Waters 1967), and it always moves the drifters downstream, at least initially. This night-time drift typically has two peaks: one just after darkness begins and one just before dawn (Waters 1972). But in some species, younger individuals may drift in the daytime and older, larger individuals at night (Anderson & Lehmkuhl 1968). Light often suppresses drifting in night drifters (Holt & Waters 1967); a full moon on a clear night can suppress it (Anderson 1966; Bishop & Hynes 1969). Brusven (1970) found that the riffle beetle Optioservus seriatus (Figure 42) was much more likely to drift as an adult compared to its larval form. This species demonstrated the complexity of the drift phenomenon, with drift relating closely to density in one stream but not in the other in this study. Larimore (1974) studied a very different kind of stream in the Salt Fork Basin, Illinois, USA. This stream ran through farmland where farm runoff was common and rooted macrophytes and bryophytes were absent. Only Chironomidae (Figure 9) among the drift organisms matched those found in cooler streams with rocky bottoms discussed above.

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Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

Some insects enter the drift to avoid or escape from predators. In experiments the net-spinning caddisfly Ceratopsyche bronta (Figure 44) moved from one area to another in an artificial stream when the predator stonefly Acroneuria lycorias (Figure 45) was present (Michael & Culver 1987). However, it did not exhibit the same drift response to the predator megalopteran Corydalus cornutus (Figure 46). Michael and Culver suggested that the caddisfly might have been unable to detect the megalopteran.

Figure 42. Optioservus seriatus adult, an insect more likely to drift as an adult than as a larva. Photo from ISUInsects.org, through Creative Commons.

Drift distances are usually not far. McLay (1970) found that the maximum drift in a New Zealand stream was 45.7 m, with a mean of only 10.7 m. Waters (1965) found that Baetis tricaudatus (Figure 43) travelled 50-60 m, but Elliott (1971a) showed that this strong swimmer was also capable of dropping out of the drift rapidly. Elliott (1967a) found that when dense macrophyte vegetation was present the maximum drift distance was only about 10 m. Nevertheless, this is sufficient to redistribute the insects and reduce local population competition.

Figure 44. Ceratopsyche bronta larva, an insect that drifts in response to the presence of the predator stonefly Acroneuria lycorias. Photo by Bob Henricks, with permission.

Figure 45. Acroneuria lycorias naiad, predator on the caddisfly Ceratopsyche bronta larvae. Photo by Tom Murray, through Creative Commons. Figure 43. Baetis tricaudatus naiad, a drifter that can travel 50-60 m in the drift, or drop out rapidly. Photo by Bob Henricks, with permission.

Many of the species enter the drift as young naiads and larvae, permitting them to disperse and to reduce population competition (Anderson 1967; Elliott 1967a, b; Waters 1969). But more frequently it is the larger stages later in the life cycle that enter the drift (Anderson 1967; Elliott 1967a; Müller 1966; Ulfstrand 1968). While drifting permits macroinvertebrates in streams to seek a more favorable location and to colonize new habitats, it poses its own set of threats (Brittain & Eikeland 1988). The insects may fall prey to predatory fish or fail to stop at a favorable habitat before reaching a quiet area of the stream where drift can no longer help them to relocate.

Figure 46. Corydalus cornutus larva, a stream predator. Photo by Alan Cressler, with permission.

Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

Researchers were curious about how the upstream positions got repopulated. Elliott (1971b) marked insects and found that some immature insects were able to move upstream on the stream bottom, especially small naiads of stoneflies and mayflies, small larvae of true flies, and beetle larvae. In winter, upstream movement was about 30% of downstream drift; in spring and summer it fell to only 7-10%. Madsen et al. (1973) examined upstream movement in adult mayflies and stoneflies and found that the representative of the common moss-dwelling stonefly genus Nemoura (Figure 40) did not move upstream, whereas the mayflies Caenis rivulorum (Figure 47), Baetis rhodani (Figure 2), B. vernus (Figure 48), and Serratella ignita (Figure 25) all moved upstream; all three of these mayfly genera are known from bryophytes. Furthermore, females migrated upstream more than males.

Figure 47. Caenis rivulorum naiad, a mayfly whose adults move upstream to lay eggs. Photo by Urmas Kruus, with permission.

Figure 48. Baetis vernus adult, a species in which females fly upstream to lay eggs. Photo by Walter Pfliegler, with permission.

Elliott (1971a) divided the drift invertebrates into three groups based on their ability to return to a substrate. The first group apparently had no control over their return to a substrate and did so at the same rate as dead organisms. This group included the Chironomidae (Figure 9). The second group includes several bryophyte dwellers, including Leuctra (Figure 49) and Simulium (Figure 22). These insects travelled shorter distances and were able to return to the substrate more quickly than dead ones at low velocities (10-12 cm sec-1) but not at faster velocities (≥19 cm sec-1). The third group, which included bryophyte dwellers such as Serratella ignita (Figure 25), Hydropsyche spp. (Figure 13), and Baetis rhodani (Figure 2), returned to the substrate significantly faster and drifted significantly shorter distances at all velocities tested; Baetis and Simulium are usually the insects with the highest numbers in the drift (Waters 1972). Caddisflies with cases fall out of the drift very quickly.

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Figure 49. Leuctra sp naiad. Photo by Guillaume Doucet , with permission.

Elliott (2003) examined dispersal in nine genera of aquatic invertebrates, most of which occur among bryophytes. He found that dispersal of invertebrates in the streams was not density dependent. Rather, it was a constant percentage of the initial number of each species. The most rapid dispersers, with 70-91% dispersing within 24 hours, were the carnivores Perlodes (Figure 50), Rhyacophila (Figure 116), and Isoperla (Figure 20), travelling up to 13.5 m per day. Protonemura (Figure 104) and Rithrogena (Figure 51) exhibited about 50% dispersal within 24 hours and travelled only about 8 m per day. The third group, Ecdyonurus (Figure 15), Hydropsyche (Figure 13), Gammarus (Figure 52), and Baetis (Figure 2, Figure 48), only had about 33-40% dispersal in 24 hours and travelled only 5.5-7 m per day. All of these genera dispersed upstream. These examples do not answer the question of why drift, but they suggest that some of that downstream drift is compensated by upstream movement.

Figure 50. Perlodes microcephala naiad, a genus in the high dispersing insects of Elliott 2003. Photo by Niels Sloth, with permission.

Figure 51. Rhithrogena impersonata naiad, a genus with 50% dispersal in 24 hours. Photo by Donald S. Chandler, with permission.

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Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

Figure 52. Gammarus pulex, a genus in the dispersing invertebrates of Elliott 2003. Photo by Niels Sloth, with permission.

Lehmkuhl (1969) found that the six mayfly species in his study, including the sometimes moss-dweller Baetis tricaudatus (Figure 43), were displaced by winter flooding. He found that in these species drift was not related to habitat. Two of the species that were abundant in the riffle areas were scarce in the drift. In the lab, drift rate did not correlate with ability of a species to hold to its substrate. Lehmkuhl and Anderson (1972) demonstrated that drift of individual species is seasonal. Within the four species of Ephemeroptera studied, some species had peak drift in October and others in May. Winter floods accounted for lesser peaks in drift. Periods of low drifting occur when a species is in its egg state, suggesting that life cycle stages are among the determinants of who is drifting. Some insects enter the drift at the time of emergence, not by choice, but because they must at that time break through the water-air interface and penetrate the surface tension. If there is no suitable emergent rock or vegetation, this becomes a nearly impossible task. Bryophyte-covered rocks can afford a better place to climb out than a smooth rock. However, there is thus far no study to determine if any insect group might seek out bryophytes as opposed to just rocks for this dangerous endeavor. The behaviors of the Hydropsyche spp. (Figure 13) are worthy of note. This net-spinning caddisfly must live near the water surface where it can trap food in its nets (Edington 1968). When released into the water, larvae would swim with side-to-side movements toward the surface (Edington 1965; Elliott 1971a). When the velocity was slow, they returned to the bottom (Elliott 1971a). When they encountered mosses in swift-flowing areas they made "firm contact." It appears that bryophytes may have a role in catching these drifters. Elliott (1967a) suggested that aquatic plants served as a natural net for drifting insects. Previously Elliott (1965) examined invertebrate drift in a Norwegian mountain stream where bryophytes formed a dense bottom cover. He did not show a direct link between the bryophyte fauna and drift, but did list the dominant insects in both. Using 400 cm2 samples, he found Baetis sp. (Figure 2), Simulium spp. (Figure 22), Rhyacophila sp. (Figure 79), Polycentropidae (Figure 24), and Plecoptera (Figure 49). When he

calculated those insects in the water column above a square meter of bottom at any time, he found that the values were extremely low, although all the insects among the top taxa in the mosses except Polycentropidae were also in the drift. At least some of the bryophyte dwellers are drift organisms, including Simulium (Figure 22), Isoperla (Figure 20), and Ephemerella (s.l.) (Figure 8) (Minshall & Winger 1968). In these three genera, the drift is suppressed by light, including that of a full moon on a clear night. Density may play a role in the number of individuals entering the drift, as in Capniidae (Figure 109), Ephemerella sp., and Hydropsyche sp. (Figure 13) in a South Carolina, USA, stream (Reisen & Prins 1972; see also Waters 1962, 1966). And, to my surprise, Minshall and Winger (1968) found that reductions in flow cause an increase in drift. The latter may relate to the need for a new location to gain suspended food or oxygen. To this end, Simulium larvae may drift at least 100 m (Carlsson 1967). Elliott (2002) calculated the rate of drift and found that most of the organisms had a very constant amount of time spent in a drifting event. For Serratella ignita (Figure 25) the mean drift time was 28.8 s, whereas for Baetis rhodani (Figure 2) it was 9.4 s, the same drift time as for the amphipod Gammarus pulex (Figure 52). For the blackfly Simulium it was only 6.4 s, with their choice of rapid water accounting for the 100 m drifting they can accomplish. In Oregon, USA, Anderson and Lehmkuhl (1968) likewise found known moss dwellers in the drift: the mayflies Paraleptophlebia (Figure 53) and Baetis (Figure 2), the stoneflies Nemoura (Figure 40), Capnia (Figure 109), and possibly Leuctra (Figure 49) (small Capnia and Leuctra are difficult to distinguish), dipterans Chironomidae (Figure 9) and Simuliidae (Figure 22). Dendy (1944) likewise found Baetis, Nemoura, Simuliidae, Chironomidae, and Hydropsychidae (Figure 13) in the drift in a stream in Michigan, USA, but added significant numbers of the mayfly Ephemerella (s.l.) (Figure 8) and caddisfly Brachycentrus americanus (Figure 54) to those found by Anderson and Lehmkuhl. To these, Reisen and Prins (1972) added the stoneflies Isogenus (probably now Isogenoides; Figure 55) and Isoperla (Figure 20).

Figure 53. Paraleptophlebia bicornuta naiad, a mossdweller genus that enters the drift. Photo by Bob Newell, with permission.

Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

Figure 54. Brachycentrus americanus larva, moss dweller that enters the drift. Photo by Donald S. Chandler, with permission.

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would support such loss by forcing at least some individuals to the smooth rock as the bryophyte itself becomes overpopulated. This would seem to eventually provide a selection factor against those organisms that did not do their nightly foraging among the mossy safe site. Is there really a selection factor involved in moss-seeking behavior? Glime and Clemons (1972) set out to determine the relative importance of bryophytes in catching such insects and constructed artificial mosses to determine how the new colonizers compared to the organisms in the drift. Clemons (unpubl data; Glime & Clemons 1972) used string mosses to determine the use of substrata similar to mosses as a catching net for drifting organisms and compared this substrate to that of real mosses and Visqueen (polyethylene plastic sheeting) strips. In the 24 hours following the placement of 7 of these artificial mosses, insects were found on the strings. These included the mayfly Baetis sp. (Figure 2), stoneflies Amphinemura nigritta (=Nemoura venosa) (Figure 56) and Leuctra sp. (Figure 49), blackflies Cnephia sp. (Figure 57) and Prosimulium mixtum (Figure 58), midges Chironomidae (Figure 9), and the caddisfly Lepidostoma sp. (Figure 59) occurring in more than one of the string habitats. The Visqueen strips had a smaller and less diverse fauna. While this experiment provides evidence that insects can settle on such substrates rather quickly from the drift, much more study is needed to determine the importance of bryophytes in providing safety nets for drifting insects. Gurtz and Wallace (1984) found that following a major disturbance that dislodged many of the insects, it was moss-covered rock faces that increased in insect density more than any other substrate. Furthermore, they considered that the mosses may enhance the stability of the substrate on which they reside.

Figure 55. Isogenoides frontalis larva, a moss-dweller that enters the drift. Photo by Donald S. Chandler, with permission.

Bryophytes may provide safe sites for drifting organisms, primarily insects. There is a periodicity in stream drift, with light, even strong moonlight, suppressing activity (Albrecht 1968). Numerous organisms, particularly stoneflies, mayflies, and caddisflies, become detached from their substrate and join the water current (Bishop & Hynes 1969). Diptera are day-active and contribute significant numbers to daytime drift. Lest they travel ultimately to a lake or even the distant sea, these drifting organisms must find a suitable substrate where they can cling against a sometimes raging current. Furthermore, it is during these excursions that they are most visible and vulnerable to predation by birds and especially fish. Bryophytes would seem to provide an ideal location for regaining their composure and taking a more leisurely approach to locating a suitable settling place. The 3-d surface of the bryophyte provides numerous "handles" for hanging on in the current and gives the insects either an instant home or one that can be traversed while maintaining a safe hold to something permanent. On the other hand, one theory for the cause of drift is to decrease population numbers (Müller 1954; Waters 1961, 1962; Pearson & Franklin 1968; Bishop & Hynes 1969). If such is the case, a rock with both smooth surface area and bryophyte cover

Figure 56. Amphinemura nigritta naiad, a rapid bryophyte colonizer. Photo by Donald S. Chandler, with permission.

Figure 57. Cnephia adult, a genus that sometimes lives among bryophytes and enters the drift. Photo by Sam Houston, with permission.

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Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

Figure 58. Prosimulium mixtum larva, a blackfly that lives among bryophytes and enters the drift. Photo by Tom Murray, through Creative Commons.

Figure 59. Lepidostoma larva, a drifting caddisfly that sometimes lives among bryophytes. Photo by Jason Neuswanger, with permission.

In alpine streams the drift pattern may differ. Hieber et al. 2003) found no night-day differences in these streams. They found that Chironomidae (Figure 9) were the dominant drifting organisms, so one might look at this group in alpine streams as creating more food for fish in streams with mosses than in those without. The complex structure of bryophytes may not only catch drift, but it may also deter stream drift. Holomuzki et al. (1999) found that resettlement choices after drifting by hydropsychid caddisfly larvae depended on the complexity of the algal community. Drift entry of hydropsychids due to stonefly predation increased on rocks with a biofilm, but not on rocks with a thick periphyton mat or macroalgae such as Cladophora (Figure 60), with drift inversely related to the amount of Cladophora on the rocks. Since bryophytes are even more complex in structure, it is reasonable to assume that they reduce drift.

It is interesting that when Perić et al. (2014) sampled the invertebrate drift in a moss-rich karst (landscape underlain by limestone that has been eroded by dissolution, producing characteristic landforms) stream system, they did not find the Chironomidae (Figure 9) (3.9%) to be the most abundant. Rather, the most abundant insects were the beetles in Elmidae (Figure 41) (13.2%) and blackflies Simuliidae (Figure 58) (12.2%). So let's revisit the possibility that other bryophyte dwellers besides Chironomidae do not enter the drift as readily as insects on other substrates. Brusven et al. (1990) found that in a channel of the South Fork Salmon River, Idaho, USA, the 20% moss-covered portion (Fontinalis neomexicana, Figure 4) had 1.6-7.2 times the diversity of the moss-free channel and 1.4-6.1 times the biomass. But the mossy portion did not have any greater numbers in the drift than did the moss-free channel. This, however, does not offer us much on which to base a conclusion because the study only included daytime drift. Their drift organisms were more than 50% Chironomidae (Figure 9), a group that drifts equally in day and night (Anderson & Lehmkuhl 1968). The implications for fish are that the bryophytes do not benefit them because the food organisms they house do not increase the daytime drift, at least in this one example.

Safe Sites For many insects, the mosses offer a safe site, a poollike environment in which they can forage for food without danger of being swept away by rapidly flowing water. Beetles (Coleoptera), scuds (Gammarus; Figure 52) and mites occupy only sheltered niches and mosses in the Welsh Dee (Badcock 1949). On vertical faces of waterfalls, the dipteran Limnophora (Figure 61) can be found only in moss (Badcock 1949).

Figure 61. Limnophora larva, sometimes a bryophyte dweller. Photo by Stephen Moore, Landcare Research, NZ, with permission.

Figure 60. Cladophora crispata, a filamentous alga that keeps Hydropsychidae from entering the drift in the presence of predatory stoneflies. Photo by Yuuji Tsukii, with permission.

In aquatic habitats, fish are a major predator on insects. The result is that fishless lakes have a higher insect species richness and diversity than lakes inhabited by fish, as demonstrated for chironomids (midge larvae) (Mousavi et al. 2002). Bryophytes are typically inhabited by many Chironomidae (Figure 9) and when present in lakes or streams they can provide safe sites with loads of detrital food.

Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

Several studies have alluded to the possibilities of bryophytes in providing a refuge, a location in the stream where the small organisms can escape predation by larger ones. For example, Parker et al. (2007) found twice as many insects on Fontinalis novae-angliae (Figure 62) as on Podostemum ceratophyllum (Figure 62). One possibility is that the insects are avoided because the moss provides an unpalatable location – an enemy-free space. Parker et al. (2007) remind us that a number of studies have shown that small herbivores that use plants as both a habitat and a food source may be protected by living on hosts that are chemically defended against wood-be insect consumers. Aquatic mosses may be just such safe sites. To test this hypothesis, Parker and coworkers observed the feeding habits of the Canada goose (Branta canadensis, Figure 63-Figure 64) and a crayfish (Procambarus spiculifer, Figure 65). In a riverine system where both the riverweed Podostemum ceratophyllum (Figure 62) and the moss Fontinalis novae-angliae (Figure 62) occurred, both animals consumed riverweed in preference to the moss. This was despite the fact that the moss comprised 89% of the plant biomass. At the same time, there were twice as many macroinvertebrates among the mosses as associated with the riverweed. Examination of the moss chemistry revealed the presence of C18 acetylenic acid, octadeca-9,12dien-6-ynoic acid, a compound that deterred the crayfish from eating it. Some invertebrates, on the other hand, had different connoisseurial preferences; the amphipod Crangonyx gracilis (Figure 66) and the isopod Asellus aquaticus (Figure 67) rejected the riverweed, but consumed significant quantities of Fontinalis novaeangliae. For periphyton-consuming insects, the same chemical deterrents could protect them without affecting their food source.

Figure 62. Podostemum ceratophyllum (red) and Fontinalis novae-angliae, the latter protecting invertebrates from grazing by geese. Photo by John Parker, with permission.

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Figure 63. Canada Goose (Branta canadensis) searching for food. Photo by Eileen Dumire, with permission.

Figure 64. Canada Geese (Branta canadensis) grazing on Podostemum ceratophyllum. Photo by John Parker, with permission.

Figure 65. Procambarus spiculifer eating Egeria. Photo by John Parker, with permission.

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Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

demonstrates that bryophytes with different mesh sizes could provide differential refugia for insects during periods of high flow rates.

Biomass and Richness

Figure 66. Crangonyx sp., an amphipod Fontinalis consumer. Photo from Biodiversity Institute of Ontario, through Creative Commons.

Many insects hang out among the riffles, taking advantage of the flowing water that brings food and oxygen. Dodd (2011) found that in a river community 516 out of 521 individuals collected occurred among riffles and mosses. These are the sites where biomass and richness usually reach their peaks. Clenaghan et al. (1998) concluded that macroinvertebrate density and richness increased with moss weight. Wulfhorst (1994) compared the biomass of insects among mosses with those in the interstitial spaces of the substrate (Figure 68). In general, they were orders of magnitude higher (100's of times) in biomass among the mosses. These included Ephemeroptera, Plecoptera, Trichoptera and Coleoptera (EPTC).

Figure 67. Asellus aquaticus, an isopod Podostemum avoider and Fontinalis consumer. Photo by Niels Sloth, with permission.

But bryophytes are not always selected for their provision of shelter. Using experimental reduction of bryophyte stem density in New Zealand alpine streams, Suren and Winterbourn (1991b) found that only two out of 22 taxa of invertebrates selected the bryophytic home based on shelter as the primary factor in the shaded site; none of them selected it based primarily on its offer of shelter in the sunny site. Rather, periphyton or detrital biomass were the primary influencing factors. Winterbottom et al. (1997) cleverly tested the importance of refugia against the effect of reduction of shear stress during periods of peak flow by creating artificial refugia using cages of different mesh sizes to restrict the flow within cages. They compared a 1.1 mm mesh size that created a reduced flow within the cage with that of a 15 mm mesh size that did not restrict flow. They found that during periods of high flow the invertebrates accumulated more in the flow-restricted refugia than they did there during low-flow periods or in the unrestricted cages. By contrast, in a second stream with lower flow rates generally and during the experimental period, the number of invertebrates did not increase in the refugia during natural spates of increased flow (but less flow than in the first stream), suggesting that the reduced flow in the 1.1 mm mesh cages enabled them to serve as refugia in the first stream during periods of rapid flow. However, the researchers were unable to determine if the accumulation of invertebrates was by active movement to the refugia or by passive collection. Nevertheless, this experiment

Figure 68. Combined biomass (mg L-1) of Ephemeroptera, Plecoptera, Trichoptera, and Coleoptera at six stations of two brooks in the Harz Mountains in mosses and interstitial spaces of the hyporheic zone at 10 and 20 cm depth. Bars show 95% CI. N = 14 for mosses, 28-36 for interstitial spaces. Redrawn from Wulfhorst 1994.

Linhart et al. (2002a, b) examined the meiobenthos (meiofauna; between .1 mm and 1 mm in size) of two loworder streams (i.e., small feeder streams) and found that these bryophytes harbored ten times as many organisms as the surrounding mineral bed. In this case, the Chironomidae (midge larvae, Figure 9) were the dominant organisms, but a number of other aquatic insects and other invertebrates call this location home, at least in the early stages of their lives. Brusven et al. (1990) studied the effect of bryophyte biomass on macroinvertebrate density in the South Fork of the Salmon River, Idaho, USA. They compared the insect densities on sand, pebbles, cobbles, and the moss Fontinalis neomexicana (Figure 4). Insect densities in moss clumps were 4-18 times as great as those in adjacent mineral substrata. Although mosses occupied only 20% of the channel, insect density was 1.6 to 7.2 times as great, with 1.4 to 6.1 times as much insect biomass as the mossfree channel, thus accounting for nearly 50% of the insects in the stream. Midges (Chironomidae, Figure 9) typically comprised over 50% of the insect community, whereas

Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

annelids were the primary non-insect invertebrates. The moss seemed to provide a safe site, at least during the day, because despite the greater number of insects present, daytime drift was not greater. Hence, the salmonid fish that feed primarily on drifting invertebrates during the day derive little benefit from the increased numbers in the bryophytes. On the other hand, Tada and Satake (1994) found that in a cool mountain stream in Japan macroinvertebrates from Platyhypnidium riparioides (Figure 19) had 11-13 taxa (species), whereas bare rock bottoms had 13-14. Nevertheless, the caddisfly Micrasema sp. (Figure 69) exceeded 100,000 individuals per m2 of mosses in November, a level that ranged 2.8-16.3 times as high as that on the bare rock bottom.

Figure 69. Micrasema charonis larva, a common genus on bryophytes. Photo by Robert G. Henricks, with permission.

Chantha et al. (2000) found that the invertebrate communities of bryophytes and algae in a Quebec, Canada, stream were dominated by Chironomidae (especially Orthocladiinae; Figure 9). The algae and invertebrates formed stable communities during the summer, even sustaining during strong mid-summer flooding. Like many other northern streams, the Ephemeroptera and Coleoptera were important components. The relative importance of the various taxa changed with the seasons as sizes and life cycle stages changed. Moss biomass explained 43% of the algal spatial variation, but surprisingly the periphyton did not increase proportionally with increase in moss biomass. The epiphytes were less dense per unit of bryophyte biomass as the bryophyte biomass increased in density. Insects in this system became more abundant, but smaller, as the moss biomass increased, with a net result of little change in insect biomass per moss biomass. This may be a function of decreased light for algal growth and decreased oxygen for insects in deeper parts of the moss mat. Matthaei et al. (2006) found that runoff from land use could reduce both aquatic mosses and invertebrate density. The greatest decrease in richness occurred in Ephemeroptera, Plecoptera, and Trichoptera, the three most abundant moss-dwelling orders that move among the open spaces of the bryophyte mats.

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Food Sources Bryophytes harbor a wide variety of invertebrates that can serve as food for the larger members of the bryophyte fauna. Dražina et al. (2011) reported 100 taxa of meiofauna among bryophytes in a European study. For example, rotifers averaged 219 individuals per cm3. Bryophytes are usually predominate in the upper reaches of streams where the flow rate is greater and the stream is shaded. Shredders likewise predominate among the bryophytes in these reaches. Hawkins and Sedell (1981) found that functional groups characterized different stretches of the river continuum. Upstream in shaded reaches the shredders were dominant. Scrapers were most important in the intermediate sections. Collectors increased in importance progressively downstream. Predators were represented equally throughout the stream. Mosses seem to afford ideal feeding locations for some kinds of insects. In particular, filterers and scrapers can be more common there than elsewhere in streams, showing a positive correlation with such habitats, whereas shredders are negatively correlated, i.e., are moss avoiders (Ely 2005). On the other hand, Zalewski et al. (2001) found a significant correlation between CPOM (coarse particulate organic matter), bryophytes, and shredders. Smith-Cuffney (1987) found that mosses in streams of a clearcut community supported collector-gatherers, whereas in the forested streams the shredders formed a much larger proportion of the moss fauna. Cattaneo et al. (2004) found that in a Québec stream grazers were more abundant in mosses than among gravel, suggesting that they used the periphyton. The reduction of periphyton when shallow water mosses are exposed may explain why deeper mosses might house more invertebrates. Wallace et al. (1988) found that the mosses retained large amounts of detritus, providing abundant food for collector-gatherers. Like Ely, they found that scrapers reached greatest abundance on cobbles and pebbles that were free of mosses. Smith-Cuffney (1987) found that in a southern Appalachian Mountain stream, mosses in a clearcut community of a forested watershed supported predominantly collector-gatherers with shredders as a minor component. Shredders were a much larger component in the stream that drained the clearcut. Scrapers were more common in the clearcut system where periphyton were abundant. Collector-filterers such as Parapsyche cardis (see Figure 70) benefited from the physical environment provided by the mosses. Although aquatic mosses are seldom eaten by their inhabitants (Haefner & Wallace 1981), they can provide a rich food source through the other inhabitants. Fontaine and Nigh (1983) considered the periphyton (Figure 71) on bryophytes to be an important food source. In New Zealand, periphyton and detritus were primary food sources (Suren 1993). Unfortunately, bryophytes tend to be shade plants and periphyton tends to prefer the sun, so the periphyton is not at its max. Nevertheless, invertebrate densities were higher among mosses containing periphyton than among those with detritus, most likely reflecting the higher food quality of periphyton. Ogbugu and Akinya (2001) likewise found that mosses in Nigeria provided a suitable substrate for periphytic algae, especially diatoms.

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Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

scrapers reached their maximum. Gregg and Rose (Gregg 1981; Gregg & Rose 1985) found that among the tracheophytes (plants with lignified vascular tissue, i.e., all plants that are not bryophytes), shredders, scrapers, and predators were the primary guilds in the autumn and that all guilds had their highest abundances in spring. Bryophytes offer the advantage of being present year-round, and their extensive periphyton growths provide a good winter food source for those insects that remain active in the winter. It is interesting that Gregg found that Hydropsyche (Figure 13), Simulium (Figure 22), Baetis tricaudatus (Figure 43), Glossosoma velona (Figure 73), and Helicopsyche borealis (Figure 74) avoided macrophytes, whereas all of these genera are known from bryophytes (though Helicopsyche is rare there). One problem for these insects was that the tracheophytes reduced the velocity, creating problems for these high-oxygen taxa. The advantage in the presence of tracheophytes seemed to be that of increasing heterogeneity, an advantage also offered by bryophytes.

Figure 70. Parapsyche apicalis larva, member of a genus known to seek shelter in bryophytes. Photo by Donald S. Chandler, with permission.

Figure 72. Drepanocladus exannulatus, a less desirable food source than Fontinalis for insect scrapers. Photo by Michael Lüth, with permission. Figure 71. Stream mosses in Tucquan Creek, Lancaster County, Pennsylvania, USA, laden with a detrital-periphyton complex. It is likely that the schist bedrock is contributing to the light color. Photo by Keith Williams, with permission.

McWilliam-Hughes et al. (2009) found Fontinalis sp. (Figure 4) abundant in headwater streams and Drepanocladus (s.l.) sp. (Figure 72) abundant in low-order streams. The scrapers living in low-order streams seemed to depend more on Fontinalis as a food source than did scrapers in high-order streams depend on Drepanocladus (s.l.). They suggested that in low-productivity, nutrientlimited rivers primary consumers might switch to marginal food sources such as bryophytes when more preferred food is limited or unavailable. The feeding guilds change with the seasons. Habdija et al. (2004) found that current velocity and food supply affected the composition of insects inhabiting bryophytes in karst streams. Those inhabiting the bryophytes were predominantly small forms of oligochaetes, Diptera (Figure 58), and Coleoptera (Figure 41), comprising 64.198.7% of the total macroinvertebrate individuals. Collector-gathers dominated in spring and summer, whereas in autumn it was collector-filterers, and in winter

Figure 73. Glossosoma sp. larvae, a tracheophyte avoider that lives among bryophytes. Photo by Jason Neuswanger, with permission.

Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

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generic or higher levels. In his study of four streams in northeastern France he found that two species in the same genus with very similar mouthparts had different diets, one feeding on bryophytes and the other on detritus, including leaf litter.

Figure 74. Helicopsyche sp. larva & case. Helicopsyche borealis avoids tracheophytes, but the genus is known from bryophytes. Photo by Stephen Moore, Landcare Research, NZ, with permission.

Thus, we have seen that the reduced water velocity within a bryophyte mat (Devantery 1987; Suren 1991) makes the bryophytes suitable safe sites not only for insects, but also for the periphyton and detrital food components, as shown in New Zealand (Suren 1991), as well as for the insect prey species, as shown in the North Temperate Zone (Elliott 2005). Bryophytes as Food Early reports indicated that bryophytes were ingested, but the food value remained in question. Nevertheless, Fontinalis (Figure 4) was found in gut contents (Gaevskaya 1969). Jones (1949, 1950) found Fontinalis in the guts of the stoneflies Amphinemura (Figure 105), Chloroperla (Figure 23), Dinocras (Figure 75), Leuctra (Figure 49), and Protonemura (Figure 104), the mayflies Ecdyonurus (Figure 15) and Ephemerella (s.l.) (Figure 8), as well as in the caddisflies Hydropsyche (Figure 13) and Philopotamus and the beetle Oreodytes (Figure 76).

Figure 76. Oreodytes septentrionalis, a genus including bryophyte consumers. Photo by Brian Eversham, with permission.

Caddisflies Pycnopsyche guttifera (Figure 77) and Philocasca alba both feed on mosses. In an interesting study, Mutch and Pritchard (1984) found that the late-instar larvae of Philocasca alba had significantly higher growth rates if their diet of detritus or leaf litter was supplemented with mosses.

Figure 77. Pycnopsyche guttifera larva, a consumer of mosses. Photo by Donald S. Chandler, with permission.

Figure 75. Dinocras cephalotes naiad, a stonefly genus that eats mosses. Photo by Guillaume Doucet , with permission.

Jones (1951) considered Fontinalis antipyretica (Figure 18) to be one of the main foods for herbivorous insects in his study of the River Towy, Wales. But Dangles (2002) cautions us against categorizing food habits by

Tada and Satake (1994), working with insects on mats of the moss Platyhypnidium riparioides (Figure 19) in a cool mountain stream in Japan, found the mayflies Baetis (Figure 43) and Ephemerella (s.l.) (Figure 8), the stoneflies Acroneuria (Figure 45) and Isoperla (Figure 20), and the caddisflies Micrasema (Figure 69), Rhyacophila (Figure 79), and Palaeagapetus rotundatus not only live among the bryophytes, but also feed on the leaves of the leafy liverwort Chiloscyphus polyanthos (Figure 78) and Scapania undulata (Figure 10). Interestingly, they do not feed on leaves of the moss Platyhypnidium riparioides, suggesting the possibility of antifeedant compounds in that species.

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Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

Figure 78. Chiloscyphus polyanthos in the fluctuating water level zone where several kinds of insects eat the leaves. Photo from , with permission.

Even the free-living carnivore caddisfly Rhyacophila dorsalis (Figure 79) apparently eats mosses (Slack 1936). One out of nine had Fontinalis antipyretica (Figure 18) leaves in the gut. For the mayfly Ephemerella (s.l.) (Figure 8), Fontinalis is a common food (Jones 1949). Ephemerella (s.l.) feeds on the green alga Ulothrix when it is available, but feeds on the ever-present moss when the alga is scarce or absent (Jones 1949). On the other hand, in a different study, Jones (1950) found that beetles and mayflies did not eat Fontinalis (Figure 18), but the moss was in the gut of Chloroperla (Figure 23), Leuctra (Figure 49), Protonemura (Figure 104), and Amphinemura (Figure 105), all stoneflies, and in the gut of the netspinning caddisfly Hydropsyche (Figure 13) – a genus that traps its food with a net. In addition to using the moss for housing, the caddisfly Micrasema (Figure 69) eats mosses and associated periphyton (Chapman & Demory 1963; Decamps & Lafont 1974). Chapman and Demory (1963) found that in its preferred food was Platyhypnidium riparioides (Figure 19). It is possible that many insects eat the mosses primarily for their associated periphyton, but for Micrasema it appears that the primary target is the mosses themselves. Even the filter-feeding blackflies such as Simulium tuberosum (Figure 80) will feed on aquatic mosses (Jones 1949), but we need to check to see if they are really digested.

Figure 80. Simulium tuberosum larva, known to have mosses in its gut. Photo by Tom Murray, through Creative Commons.

Most members of the caddisfly genus Rhyacophila (Figure 79) are carnivores, although some of these bryophyte dwellers eat bryophytes. Perhaps more importantly is their ability to hide among the mosses to ambush their prey at dusk and dawn [e.g. Baetis (Figure 43), Gammarus (Figure 52)]. Elliott (2005) found most of the Rhyacophila dorsalis (Figure 79) among clumps of the leafy liverwort Scapania (Figure 10) and the mosses Platyhypnidium riparioides (Figure 19) and Fontinalis antipyretica (Figure 18). Although most of the Rhyacophila species are carnivores, most of their guts had fragments of bryophytes, but these appeared to be undigested, exhibiting chlorophyll. Older individuals fed primarily at night and diatoms occurred in 29% of the guts of 4th instars; bryophytes occurred in 25%. However, in the 5th instar, only 9% contained diatoms and 7% contained bryophytes. The Rhyacophila larvae would disappear into the moss colony to search for food, then return to the bryophyte surface to eat it. These observations suggest that the bryophytes may have been eaten inadvertently when capturing prey. The inadvertent consumption of bryophytes by carnivores is a likely occurrence in a number of insects. For example, Jones (1950) found Fontinalis (Figure 14) in the guts of Plecoptera [Chloroperla (Figure 23), Leuctra (Figure 49), Protonemura (Figure 104), Amphinemura (Figure 105)] and Trichoptera (Hydropsyche, Figure 13), but these could have resulted from bits of the moss mixed in with their typical food. Hydropsyche is a filter feeder, spinning its own nets to trap food, but bits of drifting moss may get trapped in the net. Nevertheless, Jones did not find any Fontinalis in guts of either Coleoptera (beetles) or Ephemeroptera (mayflies) in these same collections. Nutritional and Antifeedant Properties

Figure 79. Rhyacophila dorsalis larva, a moss consumer. Photo by Walter Pfliegler, with permission.

Few protein values are published for aquatic mosses, so we cannot judge if any relationship to protein content is typical. However, it has been a common view among biologists that mosses are avoided as food because of their low food value, among other reasons. Nevertheless, Winterbourn and co-workers (1986), using C13 ratios, found bryophytes to be important sources of carbon for the benthic fauna in two British rivers.

Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

Suren and Winterbourn (1991a) examined the gut contents of 23 invertebrate taxa that dwell among bryophytes in two New Zealand alpine streams. Fourteen of these taxa had bryophytes in the guts, but the researchers found that only the tipulid larvae of Limonia hudsoni (Figure 81) and caddisfly larvae Zelandopsyche ingens (Figure 82) and Oeconesus similis (Figure 83) regularly consumed the bryophytes. They found that the bryophytes contained more refractory and indigestible compounds than other riparian plants and were thus less nutritious for the animals. They suggested that the bryophytes might also contain antifeedant compounds (compounds that discourage herbivory). Such compounds do exist in aquatic bryophytes, including Fontinalis (Liao 1993; LaCroix 1996). But we must keep in mind that modifications of digestive systems and their pH and enzymes make these "indigestible" foods digestible to some specialists (see discussion in Chapter 10-3 on Asellus).

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artificial mosses that became colonized with periphyton. But separating assimilation of moss tissue vs periphyton is a challenging endeavor.

Figure 83. Oeconesus larva head; O. similis frequently eats bryophytes. Photo by Stephen Moore, Landcare Research, NZ, with permission.

Figure 81. Limonia larva; some species are regular consumers of bryophytes. Photo courtesy of State Hygienic Laboratory, University of Iowa, with permission.

Modern methods have made it somewhat easier to determine the diets of aquatic insects. Using Δ13C, Winterbourn et al. (1986) demonstrated the importance of bryophytes as important food sources. It is surprising that so many invertebrates eat aquatic mosses. Pritchard and Berté (1987) found that the aquatic moss Leptodictyum (Figure 84) had the lowest protein content of the five foods tested (wheat flakes, alder, burreed, willow leaves, Leptodictyum. Wheat flakes and alder had the most, burreed and willow leaves were next. Nevertheless, Pritchard and Berté (1987) found that despite the low nutritional value in Leptodictyum, the caddisfly Limnephilus externus (Figure 85) chose mosses second out of the five choices, and the caddisfly Nemotaulius hostilis (Figure 86) chose mosses third among these choices. As the larvae grew, they increased their intake of moss, preferring it over alder or willow. Their preference for burreed over moss varied and was sometimes equal. Nevertheless, N. hostilis grew more slowly on mosses than on alder or burreed.

Figure 82. Zelandopsyche larva & case; some species include bryophytes in their regular diet. Photo by Stephen Moore, Landcare Research, NZ, with permission.

But sometimes the consumed mosses carry with them associated periphyton that might be the real food source, as in Micrasema (Figure 69) of the Pyrénées (Decamps & Lafont 1974). Dudley (1988) likewise considered that the real food might be the associated periphyton. Suren (1988) similarly concluded that the mosses were not an important food source, citing the similarity of faunal communities on

Figure 84. Leptodictyum riparium, an aquatic moss with lower protein content than several tracheophytes, but still eaten by the caddisfly Limnephilus externus. Photo by David T. Holyoak, with permission.

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Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

Figure 85. Limnephilus externus larvae, consumers of the moss Leptodictyum. Photo by Bob Newell, with permission.

Figure 87. Calliergon cordifolium, a moss in which acetylenic fatty acids comprise 6.6% of the triacylglycerols. Photo by Michael Lüth, with permission.

Figure 86. Nemotaulius hostilis larva in case. This species chooses mosses third compared to tracheophyte choices. Photo by Donald S. Chandler, with permission.

Tracing Bryophytes in the Food Chain If identification of assimilated bryophytes is a challenge, the identification of the role of bryophytes in the food chain is an even greater challenge. To what degree is the assimilated carbon from bryophytes passed upward to predators and top carnivores? Or is it simply stored in the insect tissues and unavailable to them? Or is it mostly lost through egestion (process of ridding the body of undigested or waste material; defecation; not to be confused with elimination of nitrogenous waste such as that in urination)? Identification of unique acetylenic fatty acids in bryophytes, including Fontinalis antipyretica (Figure 18) (Anderson & Gellermann 1975; Dembitsky & Rezanka 1995; Sushchik et al. 2007), has enabled us to use these fatty acids as markers. These unique acetylenic fatty acid markers are absent in tracheophytes, algae (e.g. Sushchik et al. 2007), and bacteria, providing us with a tool to trace bryophytes in their consumers (Dembitsky & Rezanka 1995). When testing five aquatic bryophytes, Dembitsky and Rezanka determined that acetylenic fatty acids occurring in the triacylglycerols of bryophytes comprised from 6.6% of the fatty acids in the moss Calliergon cordifolium (Figure 87) to 80.2% in the thallose liverwort Riccia fluitans (Figure 88). Identification of these unique acetylenic fatty acids opened the possibility of determining if the bryophytes were actually assimilated into tissues of their consumers (Kalachova et al. 2011).

Figure 88. Riccia fluitans, a thallose aquatic liverwort that contains 80.2% acetylenic fatty acids in its triacylglycerols. Photo by Jan-Peter Frahm, with permission.

Torres-Ruiz et al. (2007) used fatty acid content to identify the food groups eaten by several aquatic invertebrates. They found the aquatic primary producers had a higher EFA content for 18:2ω6 and 18:3ω3 in green algae, 20:5ω3 in diatoms, and 20:4ω6 in bryophytes. Furthermore, they identified specific markers for diatoms (20:5ω3 [eicosapentaenoic acid], 16:1ω7, 16:ω4s, 16Cpolyunsaturated FAa [PUFAa]), green algae (18:3ω3 [αlinolenic acid], 18:2ω6 [linoleic acid], 16C-PUFAb), and bryophytes (20:4ω6, 20:3ω3), permitting them to identify aquatic primary producers as the primary food source for the moss-dwelling mayfly Ephemerella (s.l.) (Figure 3, Figure 8) and caddisfly Hydropsyche (Figure 13). Gladyshev et al. (2012) used stable isotope composition of fatty acids to trace a food web from periphyton and mosses, to consumers, including Trichoptera, and finally to the secondary consumer fish, the grayling, in the Yenisei River in Siberia. Kalacheva et al. (2009) and Kalachova et al. (2011) used similar logic to determine the use of Fontinalis antipyretica (Figure 18) as a food source in the Yenisei River. In addition to the differences among fatty acids listed above by Torres-Ruiz et al. (2007), green algae and Cyanobacteria synthesize high amounts of α-linolenic acid

Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

(18:3ω3); bacteria synthesize odd-numbered, branched fatty acids (Kalacheva et al. 2009; Kalachova et al. 2011). Bryophytes differ from these and from tracheophytes not only by having highly specific acetylenic fatty acids, but also the levels in the bryophytes maintain a high level of these fatty acids throughout the year (Kalacheva et al. 2009). Kalacheva et al. (2009) used fatty acid and stable isotope analyses in a 4-year study on the food sources of macroinvertebrates in the Yenisei River. Using the highly specific biomarkers of acetylenic acids in Fontinalis antipyretica (Figure 18), they determined that the lipids of gammarids, Ephemeroptera, Trichoptera, and Chironomidae (Figure 9) all demonstrated the presence of these acetylenic acids in their fatty acids. In some cases, these were seasonal. For example, the amphipod Eulimnogammarus viridis exhibited maximum levels of the F. antipyretica biomarker in winter and minimum levels in summer. In particular, Serratella ignita (Figure 25) and S. setigera had the highest level of acetylenic acids A18 and A20 when analyzed. On the other hand, the Chironomidae Prodiamesa olivacea (Figure 89) and Pseudodiamesa branickii (Figure 90) and Trichoptera Apatania crymophila (Figure 91) had the lowest. The researchers concluded that for most of the aquatic insects the Fontinalis antipyretica in the Yenisei River played only a minor role in assimilation. On the other hand, the aquatic insects seemed to have a more depleted 13C content than the biofilms, an indication that the consumption of F. antipyretica, which likewise has a lower δ13C value than biofilms, contributed to their assimilation. Although the moss was consumed as a minor supplement year-round, consumption in general increased in winter when food sources such as epilithic biofilms were greatly reduced.

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into the tissues of the mayfly Serratella (Figure 25) species and others, a conclusion that cannot be supported by gut analysis alone. This line of research is worth pursuing further in other systems to determine the importance of bryophytes in the food web.

Figure 90. Pseudodiamesa branickii, a species that had low levels of bryophyte-derived acetylenic fatty acids, indicating little or no consumption of bryophytes. Photo from , with online permission.

Figure 91. Apatania crymophila larva, a caddisfly with low levels of acetylenic acid. Photo from Omnilexicon, through Creative Commons.

Macroinvertebrates can be flexible in their choices of food. In four acid streams of northeastern France, only 2436% of the biomass consumed by shredders was comprised of leaf fragments; 44% of their diet was benthic algae and bryophytes (Dangles 2002). Some taxa such as the stonefly Brachyptera seticornis (Figure 92) and caddisfly Chaetopterygopsis maclachlani (Figure 93), specialized on benthic algae and bryophytes. Even though the caddisfly Pycnopsyche guttifera (Figure 77) is a classical shredder, it eats algae and is known to eat even terrestrial mosses (Williams & Williams 1982).

Figure 89. Prodiamesa sp. larva. Prodiamesa olivacea had low levels of bryophyte-derived acetylenic fatty acids, indicating little or no consumption of bryophytes. Photo by Peter Cranston, with permission.

Kalachova and coworkers (2011) raised the question of whether the moss was consumed directly or transferred up the food pyramid by consumption of invertebrates that had eaten it. They concluded that it was direct consumption because of lack of the marker fatty acids in the invertebrates lower in the food pyramid. Perhaps the most important conclusion is that these mosses were assimilated

Figure 92. Brachyptera seticornis naiad, stonefly that specializes in eating algae and bryophytes. Photo from .

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Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

Figure 95. Tipula larva, a genus that is common among bryophytes and leaf litter and is known to feed on both mosses and liverworts. Photo by J. C. Jones, through Creative Commons. Figure 93. Chaetopterygopsis maclachlani adult. The larvae specialize on bryophytes as food. Photo from Biodiversity Institute of Ontario, through Creative Commons.

Few preference experiments have been done with aquatic mosses as a choice. Leberfinger and Bohman (2010) gave detritivores Limnephilus bipunctatus (caddisfly, Figure 94) and Nemoura sp. (stonefly; Figure 40) the choice of shrubby cinquefoil, birch, Swedish whitebeam, dead and fresh grass, aquatic moss, and algae. Both insects preferred leaves of shrubby cinquefoil; Nemoura sp. also ate algae. The dead grass was the least preferred food. The shrubby cinquefoil had the highest nutritional value among the detritus choices. Leberfinger and Bohman considered the high carbon to nitrogen content of the fresh foods to be a contributing factor in their choice.

Figure 96. Rhizomnium punctatum, food for Tipula opezoides. Photo by Jan-Peter Frahm, with permission.

Food when Food Is Scarce Bryophytes are often considered to be emergency foods for aquatic insects (Dangles 2002; McWilliamHughes et al. 2009; Kalachova et al. 2011). They can be particularly important as a winter food source when other foods become scarce (Kalachova et al. 2011). Even within the growing season, the abundance of insects changes and this changes their impact on the bryophytes they consume (Figure 97) (Dangles 2002).

Figure 94. Limnephilus bipunctatus larva in case, a species that preferred aquatic mosses over grass, but less than shrubby cinquefoil.. James K. Lindsey, with permission.

The Tipulidae (craneflies) are known from both terrestrial and aquatic habitats. In the terrestrial realm they typically live in wet habitats such as cedar swamps. Tipula oropezoides (Figure 95) is one such species. And it feeds on both mosses and liverworts. Wyatt and Stoneburner (1989) observed the larvae feeding on the moss Rhizomnium punctatum (Figure 96). It would strip the one-cell-thick lamina from the thick costa and leaf borders.

Figure 97. Density and biomass of insect shredders feeding on bryophytes in four streams in four replicate study streams (shown by 4 different symbols and lines) in Vosges Mountains (northeastern France). Modified from Dangles 2002.

Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

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Bryophytes can also serve as food in areas of a stream where other food sources are scarce (McWilliam-Hughes et al. 2009). Based on δ13C values, McWilliam-Hughes and coworkers determined that scrapers in low-order streams were more dependent on Fontinalis sp. (Figure 23) than scrapers in high-order streams depended on the Drepanocladus sp. (Figure 72) that was dominant there. In fact, 98% of the scraper δ13C values were enriched relative to bryophyte δ13C values and those two measures correlated well (r=0.53). When the values from pool habitats were removed, the correlation increased to r=0.76. McWilliam-Hughes and coworkers suggested that in lowproductivity rivers, primary consumers might switch to alternative marginal food sources such as Fontinalis sp. Epiphytes and Meiofauna of Bryophytes In aquatic habitats, bryophytes are typically covered with periphyton. This periphyton coating can serve as food for many kinds of insects. The most common of these are diatoms (Ward 1994; pers. obs.). Amos (1999) found diatoms, desmids, and filamentous algae associated with Fontinalis (Figure 62). In New Zealand, Suren (1988) found that as day length increased the mosses were covered with flocculent masses of the diatom Diatoma sp. (Figure 98) and the filamentous green alga Ulothrix sp. (Figure 99) Cyanobacteria included Placoma (Figure 100), Tolypothrix (Figure 101), and Chamaesiphon (Figure 102). Suren (1992b) found that the bryophytes provided an abundant and persistent food source for invertebrates, one that was more stable than that on plain tiles. The bryophytes grew a high biomass of the filamentous diatom Diatoma hiemale (Figure 98) in the unshaded site and the crustose diatom Epithemia sorex (Figure 103) at the shaded site. The masses of filamentous diatoms were of short duration because they were easily washed away.

Figure 99. Ulothrix, a filamentous green alga that covers stream mosses as days grow longer in spring. Photo by Yuuji Tsukii, with permission.

Figure 100. Placoma sp., a member of Cyanobacteria that covers stream mosses as days grow longer in spring. Photo by Stephen Moore, Landcare Research, NZ, with permission.

Figure 98. Diatoma hiemale, a common diatom on bryophytes at unshaded sites in New Zealand. Photo from Proyecto Agua, with permission.

Figure 101. Tolypothrix tenuis, a member of Cyanobacteria that covers stream mosses as days grow longer in spring. Photo by Yuuji Tsukii, with permission.

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Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

Figure 102. Chamaesiphon sp., member of Cyanobacteria that covers stream mosses as days grow longer in spring. Photo by Stephen Moore, Landcare Research, NZ, with permission.

meiofauna, including such small insects as the Chironomidae (Figure 9) (Aguila-S. 1998). Trapping is possible due to the reduced flow within the bryophyte mat, and this same slower flow provides a refuge from flow for stream insects (Madaliński 1961; Elliott 1967a, b; Gurtz & Wallace 1984; Suren 1992a, b; Glime 1994). Huryn and Wallace (1987) found that in mountain stream areas where bedrock outcrops are covered with mosses, collector-gatherers consume the FPOM (fine particulate organic matter) collected by the moss colony. Some probably also eat the dung that accumulates there from the many inhabitants (Fisher & Gray 1983). Cherchesova et al. (2012) suggested that small and medium stoneflies living among mosses and other locations where detritus (Figure 5) is common probably eat detritus. These include Protonemura aculeata (see Figure 104), Amphinemura trialetica (Figure 105), Taeniopteryx nebulosa (Figure 106), Taeniopteryx caucasica, Brachyptera transcaucasica (see Figure 107), Chloroperla sp. (Figure 23), Nemoura cinerea (Figure 108), Capnia nigra (Figure 109), Leuctra fusca (Figure 110), and Leuctra hippopus (Figure 111), all in genera that commonly live among mosses.

Figure 103. Epithemia sorex, a common inhabitant on bryophytes in shaded streams of New Zealand. Note the puncta (holes) in the cell wall. Photo by Ralf Wagner, with permission.

Diatoms (Bacillariophyta) at first appear to be indigestible boxes with glass shells of SiO2. However, Ogilvie and Clifford (1986) reported that insects can digest the cytoplasm of diatoms through the tiny holes (puncta; Figure 103) in the cell wall. Diatoms and detritus are important foods for the tiny insect inhabitants of bryophytes. But meiofauna, intolerant of high water velocity (Winner 1975), can also reside there, seeking refuge from the high velocity of water on rocks and other substrata in the area. As already noted, Chantha et al. (2000) found that as the moss biomass increased in a Quebec, Canada, stream, the invertebrates became more abundant but smaller. Clumps of moss with greater depth provided more spaces for invertebrates, but the algae did not increase proportionally, presumably due to diminishing light deeper into the mat. Both the algal biomass (5-fold) and invertebrate density (10-fold) was much greater on mosses compared to the nearby rocks, but the overall invertebrate biomass was similar on these two substrates because of the much greater area of bare rock.

Figure 104. Protonemura meyeri naiad, seen here amid a bed of detritus. Photo by James K. Lindsey, with permission.

Trapping Detritus The ability of bryophytes to trap detritus (Butcher 1933; Cowie & Winterbourn 1979; Gurtz & Wallace 1984; Suren & Winterbourn 1992a, b) as well as other food resources (Devantery 1987) undoubtedly plays an important role in feeding many kinds of inhabitants. Bryophytes trap CPOM (coarse particulate organic matter), FPOM (fine particulate matter), and UFPOM (ultra fine particulate organic matter) (Habdija et al. 2004). The fine particulate matter may to be particularly important for the

Figure 105. Amphinemura naiad, a stonefly that blends well with detritus. Photo by Bob Henricks, with permission.

Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

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Figure 108. Nemoura cinerea naiad, a moss and detritus dweller. Photo by James K. Lindsey, with permission.

Figure 109. Capnia sp. naiad, a detritus dweller. Photo by Jason Neuswanger, with permission.

Figure 106. Taeniopteryx nebulosa naiad, a detritus dweller. Photo by Niels Sloth, through Creative Commons.

Figure 110. Leuctra fusca, a probably detritus feeder. Photo by Louis Boumans, through Creative Commons.

Figure 107. Brachyptera risi naiad. Photo by Guillaume Doucet , with permission.

Figure 111. Leuctra hippopus naiad, a probably detritus feeder. Photo by Niels Sloth, with permission.

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Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

Suren (1992b) found that bryophytes increase the periphyton and detritus through increased habitat stability, acting much like debris jams in forested North American streams, but on a micro scale. The variation of periphyton among the bryophytes was much lower than that of plain tiles, suggesting that this food source is more stable than that on rocks. Thus the bryophyte periphyton and detritus provide persistent food sources for the bryophyte inhabitants. Linhart et al. (2002a, b) found that Chironomidae (Figure 9) and rotifers responded negatively to flow velocity, but correlated positively with the fine detrital matter trapped within the moss clump. Within Fontinalis antipyretica (Figure 18), the amount of trapped fine matter was dependent on the local flow velocity. Egglishaw (1969) found that detritus was the most important factor determining the structure of the community. The moving waters provide a continuous renewal of organic particles that serve as food items. This permits the filter feeders among the Trichoptera and the Simuliidae to form large populations there (Galdean et al. 2001). Macan and Worthington (1951) found that the fauna on different bryophyte growth forms differed. The not-sothick moss housed the mayflies Baetis (Figure 43) and Ephemerella (s.l.) (Figure 8), stoneflies, and the scud Gammarus (Figure 52). Thick mosses supported great numbers of Chironomidae (Figure 9) (75% of the individuals). These mosses support some of the greatest productivity of the fish food organisms. In Appalachian headwater streams, Wallace et al. (1988) found that thick mats of moss on the bedrock were important in retaining large amounts of organic matter. This seemed to account for the 48% collector-gatherers (insects) in the stream with dense mosses compared to 31% in the one with mostly cobbles and pebbles that were free of mosses. Habdija et al. (2000) found a positive correlation between flow velocity and the deposition rate of CPOM in moss mats, the location where most of the CPOM was deposited in an alkaline stream. Miliša et al. (2006) found similar relationships in the Plitvice Lakes of Croatia. It is interesting that some folks in the UK have proposed that the increase of aquatic bryophytes downstream of sheep-dip (insecticide & fungicide mix) or heavy metal mines may be evidence that invertebrates are a major factor controlling aquatic bryophyte abundance (Richard Lansdown, Bryonet 13 January 2008). I wonder if the metals, at least, reduce the growth of periphyton, reducing competition and permitting higher productivity among the bryophytes. On the other hand, it is possible that sheep feces provide a food source, as suggested by Fisher and Gray (1983) in regard to macroinvertebrates living in a moss matrix in a desert stream. Seasonal fluctuations in water level can present a challenge to stream macroinvertebrates. Wood et al. (2016) examined the role of trapped organic matter among the inundated clumps of the leafy liverwort Porella pinnata in the Middle Oconee River, GA, USA. This liverwort is generally above the water level, but during periods of high flow it becomes inundated. They found a significant increas in macroinvertebrate biomass, insect density, and organic matter among the P. pinnata than on adjoining

bare rock. Thus, the presence of bryophytes explained the additional organic matter, insect biomass, and density. Among these opportunistic insects were the Diptera and Plecoptera as the most abundant. I would suggest that additionally, the liverworts may have provided "landing sites" for insect that were caught up in the high-water flow.

Detrimental Effects? But the encroachment of bryophytes is not good for all members of the stream community. Bryophytes displace epilithic algae that would otherwise occupy the rocks. These diatoms and other algae serve as food for the scrapers, some of whom cannot carry out the same feeding strategy on the bryophytes. The soft structure and irregular surface of bryophytes sometimes requires a different scraping apparatus from that used on a rock. Slavik et al. (2004) found that added phosphorus in an Alaskan stream increased epilithic algae initially, but that after eight years of fertilization the bryophytes replaced the diatoms as primary producers. This increased moss growth altered ammonia uptake rates, benthic gross primary productivity, habitat structure, insect abundance, and faunal species composition. The detrimental effects of bryophyte encroachment was apparent in a South African stream when managers chose to transplant Fontinalis (Figure 18) into the stream to increase habitat for insects and ultimately increase fish production (Richards 1947). While the idea sounded good, the mosses took over the rock surfaces that had been inhabited by scrapers and insects adapted to clinging to smooth rock surfaces and displaced the native fauna. Unfortunately, I don't know the long-term outcome, which may indeed have increased the number of insects once the bryophyte-adapted species were able to colonize.

Bryophytes vs Tracheophytes It is clear that bryophytes house numerous aquatic insects. And we know that aquatic insects serve as fish food. But do the insects that live among the bryophytes achieve that role? Bowden et al. (1999) found that such a role was unclear. As will soon be seen, bryophytes serve as safe sites for the insects. On the other hand, tracheophytes usually provide a more open habitat than the small chambers of bryophytes. And the tracheophytes can house larger individuals, sheltering fish that seek food there. Macroinvertebrate biomass, insect density, and organic-matter content were significantly greater in patches of P. pinnata than on adjacent bare rock. Bryophyte biomass explained additional variation in organic matter, insect biomass, and density. The most abundant insects in P. pinnata patches were Dipterans and Plecopterans. A legitimate comparison between the bryophyte fauna and that of tracheophytes is difficult because these two plant groups tend to occupy different habitats. In lakes the bryophytes are able to extend into deeper water where there is less light than that needed to support the more rapidly growing tracheophytes. The greater depth furthermore coincides with lower temperatures and less temperature fluctuation. Nutrients and dissolved O2 also differ. And the meshlike nature of the bryophyte more easily traps detritus that can serve as a food source.

Chapter 11-2: Aquatic Insects: Bryophyte Roles as Habitats

In streams, most tracheophytes are unable to tolerate the rapid flow regime that bryophytes can withstand. Since bryophytes occupy greater flow, their surface interface can have a higher oxygen concentration. And since the bryophytes tend to occupy upstream reaches that are steeper and more rocky, they coincide with a different group of insects adapted to faster water, sometimes lower temperatures, some drying in summer, and different species of predators, especially fish. With such limitations on the comparisons, it should be no surprise that studies designed to compare the inhabitants between bryophytes and tracheophytes are rare. Harrod (1964) found that in a UK chalk stream four aquatic tracheophytes [Ranunculus fluitans (Figure 112), Callitriche platycarpa (Figure 113), Veronica beccabunga (Figure 114), and Carex sp. (Figure 115) had some inhabitants, present on all four species, that are also known bryophyte inhabitants: Baetis rhodani (mayflies; Figure 2) (Frost 1942), Rhyacophila dorsalis (free-living caddisflies; Figure 116) (Slack 1936), and Chironomidae (midges; Figure 9) (Hynes 1961). Hydropsyche sp. (net-spinning caddisflies; Figure 13) and Ephemerella (s.l.) spp. (mayflies; Figure 3, Figure 8, Figure 25) preferred C. platycarpa. Simulium ornatum (blackflies; Figure 117) dominated both Carex sp. and R. fluitans (Harrod 1964).

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Figure 114. Veronica beccabunga with flowers. Photo by Jacopo Werther, through Creative Commons.

Figure 115. Carex hystricina with flowers, a species where Simulium ornatum is dominant. Photo by Dale A. Zimmerman Herbarium, Western New Mexico University, with permission.

Figure 112. Ranunculus fluitans with flower, a species where Simulium ornatum is dominant. Photo by Rasbak, through Creative Commons.

Figure 116. Rhyacophila dorsalis larva, a bryophyte inhabitant that also occurs on aquatic tracheophytes. Photo by Walter Pfliegler, with permission.

Figure 113. Callitriche platycarpa, a preferred substrate for Hydropsyche and Ephemerella. Photo by J. C. Schou, with permission.

Krecker's (1939) model contends that invertebrate abundance varies with macrophyte biomass, but also with plant species. Those plants with finely dissected leaves have more inhabitants than do plants with broad leaves. Cyr and Downing (1988) tested this assumption with macrophytes and found that the dissected Myriophyllum spp. (Figure 118) harbored significantly (p 0.1% of the total invertebrate density in ten studies on invertebrate fauna of stream bryophytes: 1) Percival & Whitehead 1929 from a) thin moss & b) thick moss; 2) Percival & Whitehead 1930; 3) Frost 1942; 4) Egglishaw 1969; 5) Stern & Stern 1969); 6) Glime & Clemons 1972; 7) Lindegaard et al. 1975; 8) Cowie & Winterbourn 1979; 9) McKenzie-Smith 1987; 10) Smith-Cuffney 1987 from a) unshaded and b) shaded streams; 11) Suren 1991a from a) unshaded and b) shaded streams; 12) Vlčková et al. 2002; - = not reported with abundances > 0.1% total density. (from Suren 1993). The last two columns indicate the number of studies presented here in which the taxon was represented by >0.1% and the average percent of the community the taxon represented. References 8

1a

1b

2

3

4

5

6

7

Turbellaria Nematoda Oligochaeta 3.6 Tardigrada Amphipoda 1.2 Copepoda Ostracoda Isopoda Hydracarina 3.3 Collembola Ephemeroptera 15.9 Plecoptera Diptera 1.3 Chironomidae 54.3 Coleoptera 6.2 Trichoptera 4.0

3.3 1.1 3.0 6.5 1.5 40.9 4.2 0.3

24.1 0.1 57.8 3.6 0.1 9.2 3.6 0.1

0.4 0.4 2.5 1.0 4.0 2.3 83.0 2.0 3.7

4.2 44.6 2.3 34.1 1.4

0.3 4.8 1.9 0.1 2.6 2.9 77.9 0.1 9.1

2.0 5.7 12.6 71.7 2.9 3.4

10.3 6.9 6.3 4.2 16.7 33.2 0.7 -

Summary Bryophytes increase the number of niches for occupancy by aquatic insects. They increase surface area, culture algae, collect detritus, provide high prey density, and provide a refugium against the current. At the same time they permit the insects to live in the greater oxygen provided by the rapid flow, saving them ventilation energy. Feeding groups of these insects include collector-gatherers, scrapers, shredders, collector-filterers, and engulfers, with collectorgatherers typically being most abundant. Altitude and latitude are important determinants of both the bryophytes and the associated fauna. Thickness of the moss mat also is important in determining the fauna, with thicker mats creating more niches. The most common orders of moss dwellers in streams are Ephemeroptera (mayflies), Plecoptera (stoneflies), Trichoptera (caddisflies), and Diptera (flies). Streams in the Arctic and alpine habitats lack most of the Trichoptera (caddisflies), but otherwise have similar order representation among stream bryophytes, with even more Chironomidae. The associations of insects with the species of bryophytes may be a consequence of both needing similar conditions, as exemplified by the similarities of insect

22.5 21.2 33.7 23.6

9

10a

10b

11a

11b

2.9 42.5 5.46 5.4 21.6 2.3 13.4

1.6 1.8 6.0 2.7 1.2 15.2 3.1 1.1 54.0 6.2

2.8 1.4 4.0 7.0 1.8 8.2 6.1 53.0 7.9

22.1 2.4 9.0 -2.8 1.1 2.1 1.5 57.7 -

12.5 1.5 0.7 5.9 2.5 7.7 63.4 -

No. Av % 12 Studies Comp 0.26 14.65 0.57 0.59 0.47 0.13 0.73 0.88 0.01 1.96 33.81 0.15 0.29

3 4 8 1 6 5 2 1 10 2 9 10 11 14 8 11

0.2 2.8 5.8 0.2 4.0 5.3 0.3 0.2 2.4 0.9 4.1 11.4 5.2 49.1 1.5 5.2

communities on the moss Fontinalis dalecarlica and the liverwort Scapania undulata, two species that often occur side-by-side. Nevertheless, bryophytes do not make good surrogates for the stream inhabitants, correlating primarily with nutrient levels and habitat heterogeneity, whereas insects correlate more with stream size, pH, and water color. In fact, clumps of string and other artificial mosses seem to attract communities similar to those on real mosses. On the other hand, the presence of bryophytes will usually indicate a high density of insects. The bryophytes may serve as a refuge for insects in winter when non-bryophyte plants are absent and the bryophytes are common in fast water where freezing is less common. The bryophytes furthermore serve as a location of collected detritus and a site for winter diatoms. Within the clump of bryophytes of a stream one can find a detritus zone with little or no flow, a water zone within the moss clump, and a madicolous zone just above the water surface but where the bryophytes are still wet. And at the surface of the moss, but submerged, the highest water velocity and therefore the most oxygen exist. Waterfalls may have specialists that live among the wet mosses, avoiding the torrent itself. Springs often have dense bryophyte cover. Chironomidae here

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Chapter 11-3: Aquatic Insects: Bryophyte Habitats and Fauna

respond to temperature; many insects also respond to nutrient concentrations or pH. Depth of streams, pools, and springs can influence insect community composition, in part because of temperature and oxygen gradients. Bogs and fens have both pool and dry hummock conditions, contributing a wide range of niches that differ in moisture, temperature, and light. Consequently, there is a wide variety of insects, and even flying adults make use of the mosses for egg deposition, mating, and resting. More Collembola (springtails) are found in bogs and fens than in most aquatic habitats. Coleoptera (beetles) and Odonata (dragonflies and damselflies) likewise are common in these habitats. Hymenoptera (ants, bees) are absent from streams and lakes, but in bogs and fens ants build nests from the Sphagnum. Little seems to be published about insects associated with lake bryophytes. Some of the beetles are associated with floating Riccia fluitans and Ricciocarpos natans in shallow lakes. In one case, the latter is inhabited by the leaf miner Phytoliriomyza mesnili. Disturbance immediately reduces the number of invertebrates, but if mosses remain or are replaced, they are quickly recolonized by remaining drifting organisms or from egg-laying. Attempts at restoration can cause the bryophytes to break loose and reduce the insect fauna. If one compares the bryophyte fauna around the world, differences in relative abundance of the orders are apparent. These differences are often the result of evolutionary and distributional differences. For example, the families of the insects are different in Australia and New Zealand from those in North America.

Acknowledgments My gratitude goes to my sister, Eileen Dumire, for her candid suggestions for improvement of this chapter. I appreciate all the photographers who have kindly given me permission to use their images and to those who have contributed their images to Creative Commons.

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concordant across multiple taxonomic groups? Freshwat. Biol. 48: 1912-1923. Paavola, R., Muotka, T., Virtanen, R., Heino, J., Jackson, D., and Mäki-Petäys, A. 2006. Spatial scale affects community concordance among fishes, benthic macroinvertebrates, and bryophytes in streams. Ecol. Appl. 16: 368-379. Parker, S. M. and Huryn, A. D. 2006. Food web structure and function in two Arctic streams with contrasting disturbance regimes. Freshwat. Biol. 51: 1249-1263. Percival, E. and Whitehead, H. 1929. A quantitative study of the fauna of some types of stream-bed. J. Ecol. 17: 282-314. Percival, E. and Whitehead, H. 1930. Biological survey of the river Wharf. II. Report on the invertebrate fauna. J. Ecol. 18: 286-295. Plitt, C. C. 1907. Webera sessilis and ants. Bryologist 10: 54-55. Reichle, D. E. 1966. Some pselaphid beetles with boreal affinities and their distribution along the postglacial fringe. Syst. Zool. 15: 330-344. Reichle, D. E. 1967. The temperature and humidity relations of some bog pselaphid beetles. Ecology 48: 208-215. Richards, O. W. and Davies, R. G. 1977. Imm's General Textbook of Entomology, 10th edn. Chapman and Hall, London. Richardson, D. H. S. 1981. The Biology of Mosses. Chapter 8, Mosses and micro-organisms. John Wiley & Sons, Inc., New York, pp. 119-143. Robinson, C. T., Uehlinger, U., and Hieber, M. 2001. Spatiotemporal variation in macroinvertebrate assemblages of glacial streams in the Swiss Alps. Freshwat. Biol. 46: 16631672. Rosa, B. F., Silva, M. V. da, Oliveira, V. C. D., Martins, R. T., and Alves, R. D. G. 2011. Macroinvertebrates associated with Bryophyta in a first-order Atlantic Forest stream. Zoologia (Curitiba) 28: 351-356. Sahlén, G., Bernard, R., Rivera, A. C., Ketelaar, R., and Suhling, F. 2004. Critical species of Odonata in Europe. Internat. J. Odonatol. 7: 385-398. Scotland, M. B. 1934. The animals of the Lemna association. Ecology 15: 290-294. Slavik, K., Peterson, B. J., Deegan, L. A., Bowden, W. B., Hershey, A. E., and Hobbie, J. E. 2004. Long-term responses of the Kuparuk River ecosystem to phosphorus fertilization. Ecology 85: 939-954. Smirnov, N. N. 1961. Food cycles in sphagnous bogs. Hydrobiologia 17: 175-182. Smith-Cuffney, F. L. 1987. Ecological interactions in the moss habitat of streams draining a clearcut and a reference watershed. Unpublished Ph.D. thesis, University of Georgia, 174 pp. Spencer, K. A. 1990. Division Bryophyta. In: Host Specialization in the World Agromyzidae (Diptera) series Entomologica, vol. 45. Kluwer Academic Publishers, Springer, Netherlands, pp. 1-3. Stern, M. S. and Stern, D. H. 1969. A limnological study of a Tennessee cold springbrook. Amer. Midl. Nat 82: 62-82. Suren, A. M. 1987. The ecological role of bryophytes in high alpine streams of New Zealand. In: Sladecek, V. (ed.). Proceedings of the Conference (23) Congress in New Zealand, Hamilton, NZ, 8 Feb 1987. Suren, A. M. 1988. Ecological role of bryophytes in high alpine streams of New Zealand. Internat. Ver. Theor. Angew. Limnol. 23: 1412-1416. Suren, A. M. 1991a. Bryophytes as invertebrate habitat in two New Zealand alpine streams. Freshwat. Biol. 26: 399-418.

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Suren, A. M. 1991b. Assessment of artificial bryophytes for invertebrate sampling in two New Zealand alpine streams. N. Z. J. Marine Freshwat. Res. 25: 101-112. Suren, A. 1993. Bryophytes and associated invertebrates in firstorder alpine streams of Arthur's Pass, New Zealand. N. Z. J. Marine Freshwat. Res. 27: 479-494. Suren, A. M. and Winterbourn, M. J. 1992. The influence of periphyton, detritus and shelter on invertebrate colonization of aquatic bryophytes. Freshwat. Biol. 17: 327-339. Tarras-Wahlberg, N. 1952-53. Oribatids from the Åkhult-Mire. Oikos 4: 166-171. Thienemann, A. 1936. Alpine chironomiden. Arch. Hydrobiol. 30: 167-262. Thomas, A. G. B. 1980. Dipteres Torrenticoles Peu Connus 7. Les Cecidomyiddae Porricondylinae du Sud-Ouest de la France (Nematocera). [Poorly known torrential Diptera. 7. Cecidomyiidae Porricondylinae (Nematocera) from SouthWest of France.]. Ann. Limnol. 16(3): 225-231. Thorup, J. 1963. Growth and life-cycle of invertebrates from Danish springs. Hydrobiologia 22: 55-84. Thorup, J. and Lindegaard, C. 1977. Studies on Danish springs. Folia Limnol. Scand. 17: 7-15. Usinger, R. L. 1974. Aquatic Insects of California. University of California Press, Berkeley. Uvarov, B. 1977. Grasshoppers and Locusts, a Handbook of General Acridology, Vol. 2. Centre for Overseas Pest Research, London. Vickery, V. R. 1969. Two species of Pteronemobius previously unreported in Quebec (Orthoptera: Ensifera: Grylloidea: Nemobiinae). Ann. Soc. Entomol. Quebec 14: 22-24. Vinson, M. R. and Hawkins, C. P. 2003. Broad-scale geographical patterns in local stream insect genera richness. Ecography 26: 751-767.

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Virtanen, R., Ilmonen, J., Paasivirta, L., and Muotka, T. 2009. Community concordance between bryophyte and insect assemblages in boreal springs: A broad-scale study in isolated habitats. Freshwat. Biol. 54: 1651-1662. Vlčková, Š., Linhart, J., and Uvíra, V. 2002. Permanent and temporary meiofauna of an aquatic moss Fontinalis antipyretica Hedw. Acta Univ. Palacki. Olomuc 39/40: 3140. Vuori, K.-M. and Joensuu, I. 1996. Impact of forest drainage on the macroinvertebrates of a small boreal headwater stream: Do buffer zones protect lotic biodiversity? Biol. Conserv. 77: 87-95. Wallace, J. B. and Ross, H. H. 1971. Pseudogoerinae: A new subfamily of Odontoceridae (Trichoptera). Ann. Entomol. Soc. Amer. 64: 890-894. Ward, J. V. 1986. Altitudinal zonation in a Rocky Mountain stream. Arch. Hydrobiol. Suppl. 74: 133-199. Ward, J. V. and Dufford, R. G. 1979. Longitudinal and seasonal distribution of macroinvertebrates and epilithic algae in a Colorado springbrook-pond system. Arch. Hydrobiol. 86: 284-321. Wehr, J. D. and Whitton, B. A. 1983. Accumulation of heavy metals by aquatic mosses. 3. Seasonal changes. Hydrobiologia 100: 285-291. Williams, D. D. 1980. Some relationships between stream benthos and substrate heterogeneity. Limnol. Oceanogr. 25: 166-172. Winterbourn, M. J., McDiffett, W. F., and Eppley, S. J. 2000. Aluminium and iron burdens of aquatic biota in New Zealand streams contaminated by acid mine drainage: Effects of trophic level. Sci. Total Environ. 254(1): 45-54.

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Glime, J. M. 2017. Aquatic Insects: Hemimetabola – Collembola and Ephemeroptera. Chapt. 11-4. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. Ebook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 19 July 2020 and available at .

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CHAPTER 11-4 AQUATIC INSECTS: HEMIMETABOLA – COLLEMBOLA AND EPHEMEROPTERA TABLE OF CONTENTS COLLEMBOLA....................................................................................................................................................... 11-4-2 Isotomidae ......................................................................................................................................................... 11-4-6 Bog Springtails .................................................................................................................................................. 11-4-6 HEMIMETABOLA ................................................................................................................................................. 11-4-9 EPHEMEROPTERA – Mayflies .............................................................................................................................. 11-4-9 Suborder Furcatergalia .................................................................................................................................... 11-4-12 Leptophlebiidae – Prong-gilled Mayflies ................................................................................................. 11-4-12 Caenidae – Small Squaregill Mayflies ..................................................................................................... 11-4-13 Neoephemeridae ...................................................................................................................................... 11-4-13 Ephemerellidae – Spiny Crawlers ............................................................................................................ 11-4-14 Seasons ............................................................................................................................................. 11-4-17 Food .................................................................................................................................................. 11-4-18 Ephemerella ...................................................................................................................................... 11-4-19 Serratella .......................................................................................................................................... 11-4-20 Teloganopsis ..................................................................................................................................... 11-4-22 Cincticostella .................................................................................................................................... 11-4-22 Drunella ............................................................................................................................................ 11-4-23 Caudatella......................................................................................................................................... 11-4-24 Attenella ............................................................................................................................................ 11-4-24 Torleya .............................................................................................................................................. 11-4-24 Leptohyphidae – Little Stout Crawler Mayflies ....................................................................................... 11-4-24 Suborder Pisciforma ........................................................................................................................................ 11-4-25 Ameletidae – Combmouthed Minnow Mayflies ...................................................................................... 11-4-25 Baetidae – Blue-winged Olives ................................................................................................................ 11-4-25 Siphlonuridae – Primitive Minnow Mayfly ............................................................................................. 11-4-28 Heptageniidae – Clinger Mayflies ........................................................................................................... 11-4-28 Isonychiidae ............................................................................................................................................. 11-4-29 Oligoneuriidae – Brushleg Mayflies ........................................................................................................ 11-4-29 Suborder Carapacea ........................................................................................................................................ 11-4-29 Baetiscidae – Armored Mayflies .............................................................................................................. 11-4-29 Summary ......................................................................................................................................................... 11-4-30 Acknowledgments ........................................................................................................................................... 11-4-30 Literature Cited ............................................................................................................................................... 11-4-30

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Chapter 11-4: Aquatic Insects: Hemimetabola – Collembola and Ephemeroptera

CHAPTER 11-4 AQUATIC INSECTS: HEMIMETABOLA – COLLEMBOLA AND EPHEMEROPTERA

Figure 1. Serratella ignita, a common moss dweller. Photo by J. C. Schou, with permission.

COLLEMBOLA – Springtails This group was traditionally considered to be one of the insect orders, but more recently they have been classified in the class Entognatha. Collembola are quite small and lack wings. They have three pairs of legs, like insects, but have only six abdominal segments (Thorp & Covich 1991). The young (nymphs) resemble the adults, changing to adults by breaking their outer covering (exoskeleton) and discarding it, then expanding while the new exoskeleton is still soft.. They are unique in having a furcula (Figure 3-Figure 5) that forms the spring and a collophore (cylindrical ventral tube; Figure 3, Figure 6). When at rest, the furcula bends forward under the abdomen and is held in place by the tenaculum (Figure 3), a midventral structure that clasps the furcula. The springtail accomplishes rapid distance movement by releasing the furcula, which springs backward, propelling the springtail forward several centimeters. This can be used even on the

water surface. Some can be seen bouncing around on the snow in winter.

Figure 2. Podura aquatica moulting; note split in outer skeleton. Photo by Jan van Duinen, with permission.

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the springtail. It is these bacteria that control the parthenogenesis in the colonized species. That is, they feminize the springtails.

Figure 3. Collembola external anatomy. Modified from Cooperative Extension illustration, University of Missouri.

Figure 6. Isotoma (springtail) showing collophore (arrow). Photo by U. Burkhardt, through Creative Commons.

Figure 4. Arthropleona oruarangi showing furcula. Photo by Stephen Moore, Landcare Research, NZ, with permission.

Figure 7. Collembola eggs. Photo by Jan van Duinen, with permission.

Figure 5. Dicyrtomina ornata ventral side showing furcula. Photo by Jan van Duinen, with permission.

Collembola can be sexual or parthenogenetic. Sexual males deposit spermatophores in clusters or individually. Females stimulate this deposition by producing pheromones (Waldorf 1974). But among many of the soil Collembola, presumably including bryophyte dwellers, females lay eggs (Figure 7-Figure 8) that have not been fertilized, i.e., are produed parthenogenetically. Since few reproductive studies exist, I cannot generalize of aquatic bryophyte dwellers. What makes this reproduction so interesting is the role of symbiotic bacteria in the genus Wolbachia (Werren et al. 1995). These bacteria live in and reproduce in the female reproductive organs and eggs of

Figure 8. Sminthurides eggs in duckweed. Photo by Jan van Duinen, with permission.

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Chapter 11-4: Aquatic Insects: Hemimetabola – Collembola and Ephemeroptera

The Collembola are predominately moist terrestrial organisms, but some can hop on the water surface (Figure 9) or live among wet mosses. Waltz and McCafferty (1979) considered only 10 species as semiaquatic and five as riparian (relating to bank of river or other moving water). The waxy cuticle (Chang 1966), coupled with small size, permits them to float on water. The collophore (ventral tube) serves a double function: absorption of water and respiration. The Collembola seem to be particularly responsive to drawdown and drainage (Silvan et al. 2000). On older drained sites their numbers were up to 100 times as high compared to pre-drawdown. Other invertebrates were typically about ten times as high. The Collembola occurred mostly in the top 4 cm of the drained land.

collections using insect nets. Others may have "sprung" away from surface locations as the collector approached.

Figure 10. Odontella cf. incerta; O. lamellifera is a springtail that occasionally occurs among stream bryophytes in the Appalachian Mountains, USA. Photo by Andy Murray, through Creative Commons.

Figure 9. Collembola (springtails) on water where they can jump about on the surface tension. Photo by Janice Glime.

In my search for information on the bryophytedwelling springtails, I was surprised to find so little that related to aquatic habitats. In my own studies in the Appalachian Mountain, USA, streams, I found representatives of eight families, albeit not frequently. The species in these collections were Odontella lamellifera (Figure 10) (Brachystomellidae), Entomobrya griseoolivata (Figure 11) and Orchesella quinquefasciata (Figure 12) (Entomobryidae), Hypogastrura armatus (see Figure 13), and Schotella glasgowi (Hypogastruridae), Hydroisotoma schaefferi (Figure 14), Isotoma violacea, Isotoma viridis (Figure 15), and Isotomurus palustris (Figure 16) (Isotomidae), Pseudachorutes lunatus (Neanuridae; see Figure 17), Onychiurus subtenius (Onychiuridae), Sminthurides aquaticus (Figure 18) (Sminthuridae), and Tomocerus flavescens (Figure 19) (Tomoceridae). Of these taxa, only Isotomurus palustris was present in more than two collections. Nevertheless, I recorded Orchesella quinquefasciata in North America for the first time (Toliver Run, Garrett County, MD) (Richard Snider, pers. comm.). The Hydroisotoma schaefferi was an atypical blind form from Little Bennett Creek,. Montgomery Co., MD. Snider also found this species (not blind) in ponds surrounded with mosses in Michigan, USA (Snider 1967). It is likely that some of these springtails were living at the surface of emergent mosses. But the tiny size of these insects suggests they may have been missed in

Figure 11. Entomobrya griseoolivata, a springtail that sometimes occurs among Appalachian Mountain stream bryophytes. Photo by Domingo Zungri, through Creative Commons.

Figure 12. Orchesella quinquefasciata, a springtail that sometimes occurs among Appalachian Mountain stream bryophytes. Photo by Malcolm Storey, through DiscoverLife Creative Commons.

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Figure 13. Hypogastrura nivicola; H. armatus is a springtail that sometimes occurs among Appalachian Mountain stream bryophytes in eastern USA. Photo by Scott Justis, with permission.

Figure 16. Isotomurus palustris, an aquatic springtail that keeps its offspring together for two days after birth. Photo by Scott Justis, with permission.

Figure 17. Pseudachorutes sp.; Pseudachorutes lunatus lives among mosses in mountain streams. Photo by Jan van Duinen, with permission. Figure 14. Hydroisotoma schaefferi, a springtail that sometimes occurs among Appalachian Mountain stream bryophytes. Photo by Tom Murray, through Creative Commons.

Figure 15. Isotoma viridis, a springtail that sometimes occurs among Appalachian Mountain stream bryophytes. Photo by Kyron Basu, through Creative Commons.

Figure 18. Sminthurides aquaticus, a springtail that sometimes occurs among Appalachian Mountain stream bryophytes. Photo by Andy Murray, through Creative Commons.

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Chapter 11-4: Aquatic Insects: Hemimetabola – Collembola and Ephemeroptera

Figure 19. Tomocerus flavescens, a springtail that sometimes occurs among Appalachian Mountain stream bryophytes. Photo by Royce Bitzer, through Creative Commons.

Isotomidae The family Isotomidae was most frequently (almost exclusively among springtails) represented in the publications I found regarding bryophyte fauna. Among these, Isotomurus palustris (Figure 16) is most typically considered to be aquatic, although a few other species, including Sminthurus aquaticus (Figure 18), have names that suggest they are aquatic. Isotomurus palustris (Figure 16) is able to float on the water because of their non-wetting waxy epicuticle composed of a lipid monolayer that is extremely impermeable to water (Beament 1960). But Noble-Nesbitt (1963) provided evidence that the presence of wax gives it hydrofuge (shedding water) properties. A cementing substance contributes to this hydrofuge ability. The cuticle, combined with surface hairs, provides this springtail with a protective air layer that both makes these springtails unwettable (repelling water) and makes them float. Springtails also are very sensitive to desiccation, so the protection by the cuticle is important. The collophore is wettable (doesn't repel water) and doubles as both a respiratory and water-taking organ (Noble-Nesbitt 1963). The air layer on the surface also behaves as a plastron (breast plate breathing apparatus). These springtails also take water by mouth and this may additionally supply dissolved oxygen. I wonder if they ever get hiccups! This tubule, combined with their small size, would permit them to drink water from the leaves of emergent mosses. But it appears that the cuticle may also play an important role in their locomotion on the water surface (Noble-Nesbitt 1963). In the water, the furcula is used as a spring, much as it is on land. On the water surface the insect actually walks, using only its limbs. Isotomurus palustris (Figure 16) is viviparous, producing one egg at a time (Chang 1966). These eggs are carried internally and hatched inside the female with the nymph emerging from the genital pore. The female arches its body to permit the emerging nymph to reach the water surface. In observations on newborns of Isotomurus palustris (Figure 16) and Folsomia fimetaria (Figure 20), Chang found that the newborns stayed close to the mothers for the first two days. The young are able to float, walking on the surface tension with their non-wetting (repelling water) claws, but if they are forced to submerge they will sink. The cuticle does not develop until they spend time above water.

Figure 20. Folsomia fimetaria, a springtail whose newborns stay close to the mother for two days. Photo by Andy Murray, through Creative Commons.

Antennae are important in assessing the environment in both Isotomurus palustris (Figure 16) and Folsomia fimetaria (Figure 20). They are the sensory organ, often in consort with the post-antennal organ, that recognizes light intensity, wind direction, and heat. When one or the other of these organs is removed or cauterized, the springtails move about aimlessly or not at all, whereas those with both organs intact wiggle their antennae and exhibit a directional movement in response to the stimulus. Some Collembola like it cold – Anurida frigida (Neanuridae) occurs under mosses on stones and on stones by melt-water brooks in the high alpine of Swedish Lapland (Fjellberg 1973). The greatest numbers of these were located under mosses that were wet by ice-cold meltwater. In the Nordic countries, Agrenia riparia prefers wet mosses, especially on lowland stream banks (Fjellberg 2007b) Bog Springtails These tiny creatures seem often to be overlooked, but a treatment of Collembola in Michigan, USA, indicates that many species can occur in bogs (Snider 1967): Hypogastrura nivicola (Onychiuridae; Figure 21) Isotoma viridis (Isotomidae; Figure 15) Lepidocyrtus cyaneus (Entomobryidae; Figure 32) Lepidocyrtus lignorum (Entomobryidae; Figure 22) Lepidocyrtus unifasciatus (Entomobryidae) Lepidocyrtus violaceous (Entomobryidae; Figure 23) – in Sphagnum Neelus minutus (Neelidae; see Figure 24) Orchesella ainsliei (Entomobryidae) Orchesella albosa (Entomobryidae) Pseudobourletiella spinata (Sminthuridae; Figure 25) Sminthurides aquaticus (Sminthuridae; Figure 18) – in Sphagnum Sminthurides lepus (Sminthuridae) Sminthurides malmgreni (Sminthuridae; Figure 26) – semi-aquatic habitats Sminthurides occultus (Sminthuridae) Sminthurides penicillifer (Sminthuridae; Figure 27) Sminthurinus aureus (Sminthuridae; Figure 28) Sminthurinus bimaculatus (Sminthuridae; Figure 29) Tomocerus flavescens (Tomoceridae; Figure 19) – in Sphagnum

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Figure 24. Neelus murinus carrying eggs; Neelus minutus is a bog dweller. Photo by Frans Janssens, with permission.

Figure 21. Hypogastrura nivicola on snow. Charley Eiseman, through Creative Commons.

Photo by

Figure 22. Lepidocyrtus lignorum, a bog inhabitant. Photo by Jan van Duinen, with permission. Figure 25. Pseudobourletiella spinata, a bog inhabitant. Photo by Tom Murray, through Creative Commons.

Figure 23. Lepidocyrtus violaceus, a bog Sphagnum dweller. Photo by Jan van Duinen, with permission.

Figure 26. Sminthurides malmgreni, a bog inhabitant. Photo by Andy Murray, through Creative Commons.

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Chapter 11-4: Aquatic Insects: Hemimetabola – Collembola and Ephemeroptera

pseudassimilis in boreal Sphagnum bogs and smaller lakes, boreal; Sminthurides parvulus uncommon in bogs, wet meadows, and shores of lakes; Neelides minutus uncommon in bogs; Arrhopalites cochlearifer and Arrhopalites principalis (common) in bogs; Isotomurus unifasciatus (Figure 33) in forest bogs; Isotomurus balteatus in boreal bogs and wetlands; Dicyrtomina minuta and Dicyrtoma fusca (Figure 34) common in bogs; Heterosminthurus insignis in wet meadows and bogs.

Figure 27. Sminthurides nr. penicillifer female, a bog inhabitant. Photo by Andy Murray, through Creative Commons.

Figure 30. Desoria olivacea, a species of acidic forest bogs. Photo by Jan van Duinen, with permission.

Figure 28. Sminthurinus aureus, a bog dweller. Photo by Andy Murray, through Creative Commons.

Figure 31. Desoria blufusata, a common species in bogs and wet meadows. Photo by Arne Fjellberg, through Creative Commons.

Figure 29. Sminthurinus bimaculatus, a bog dweller. Photo by Andy Murray, through Creative Commons.

In his treatment of the Collembola of Fennoscandia and Denmark, Fjellberg (2007a) included Maristoma canaliculata as a species usually found in Sphagnum and Maristoma tenuicornis in Sphagnum bogs. The treatment for Nordic Collembola (Fjellberg 2007b) includes Marisotoma canaliculata in Sphagnum ponds; Marisotoma tenuicornis in boreal Sphagnum bogs; Desoria olivacea (Isotomidae; Figure 30) common in acidic forest bogs; Desoria blufusata (Figure 31) in bogs and wet meadows; Lepidocyrtus cyaneus (Entomobryidae; Figure 32) common in humid habitats including Sphagnum/Salix bogs; Sminthurides schoetti common in bogs and damp meadows; Sminthurides

Figure 32. Lepidocyrtus cyaneus, a species of Sphagnum bogs. Photo by Steve Hopkin, with permission.

Chapter 11-4: Aquatic Insects: Hemimetabola – Collembola and Ephemeroptera

Figure 33. Isotomurus unifaciatus, a species of boreal bogs and wetlands. Photo by Jan van Duinen, with permission.

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The immature mayflies, known as naiads, are all aquatic (Thorp & Covich 1991). They can be distinguished by their three (two in some) long caudal filaments that are also present in the adults. They are most similar to the stoneflies (Plecoptera – see subchapter on Plecoptera in this chapter), but differ in having abdominal gills (lacking in middle abdominal segments of stoneflies) and typically three tails (caudal filaments), which always number two in stoneflies. Most of the naiads are herbivores and some eat bryophytes. The mayfly naiads are largely night-active and appear most often in the night-time drift (Elliott 1967). Adult mayflies emerge from the naiad first as a sub-imago (also known as a dun; Figure 35-Figure 40), a stage that often becomes a nuisance to motorists (Figure 36) in the area because of the large numbers that meet their demise (Figure 37) on the windshields. To complete emergence they must climb so they can pump fluids into their new wings (Figure 41). The adult does not eat – in fact lacking mouthparts – and typically lives for only a few days.

Figure 35. Baetis male subimago emerging to adult. Photo by Jason Neuswanger at , with permission.

Figure 34. Dicyrtoma fusca, a species common in bogs. Photo by Jan van Duinen, with permission.

Greenslade et al. (2006) suggests that Mesaphorura macrochaeta may have been introduced to the Southern Hemisphere by human importations of soil and moss peat.

HEMIMETABOLA The hemimetabolous insects are those with incomplete metamorphosis. Instead of a larva, they have a nymph or naiad stage that resembles the adult except for having reduced wings or only wing pads. They lack a pupa stage and pass directly from the nymph or naiad stage to the adult stage. Most of the aquatic Hemimetabola have a stage with gills and wing pads and are distinguished as naiads.

EPHEMEROPTERA – Mayflies As in most of the names of insect orders, optera refers to wings. In the Ephemeroptera, ephemera refers to short-lived. Hence, these are insects that are short-lived in the winged, or adult, stage.

Figure 36. Adult mayflies on emergence day. Photo by Jeff Reutter, through Ohio Sea Grant public domain.

In my own studies in the Appalachian Mountain streams, USA (Glime 1968, 1994), the Ephemerellidae was by far the most abundant of the mayflies. Frost (1942) reported the importance of the mayflies Ephemerella (s.l.) (Figure 45) and Baetis (Baetidae; Figure 35-Figure 40) among aquatic mosses, where they feed mostly on algae, but occasionally on bryophytes (Hynes 1961; Chapman & Demory 1963). Frost (1942) found about 530 mayfly nymphs per 200 g of mosses in Ireland. In a cool mountain stream of central Japan, Tada and Satake (1994) found that Baetis thermicus (Figure 38) and Ephemerella (s.l.) sp.

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Chapter 11-4: Aquatic Insects: Hemimetabola – Collembola and Ephemeroptera

were more abundant among the moss Platyhypnidium riparioides (Figure 39) than in bare rock areas.

Figure 40. Baetis sub-imago showing huge eyes. Photo by Jason Neuswanger at , with permission.

Figure 37. Mayflies that met their end on a travelling car during an emergence in August in Michigan, USA. Photo by Eileen Dumire, with permission.

Figure 41. Emerging Ephemeroptera. Mayflies live their immature lives as naiads in the water of streams and lakes. When they emerge as adults, they must climb, like these naiads, so they can pump up their wings once they have exited the naiad exuvia. Photo by Jason Neuswanger at , with permission.

Figure 38. Baetis thermicus naiad, a common moss dweller of the moss Platyhypnidium riparioides in Japan. Photo from Shiiba Research Forest. Permission requested.

With such a dwarfed lifespan, finding a mate quickly is paramount. This is accomplished by flying in giant swarms, facilitated by coordinated emergence time. At this time, they are a nuisance for motorists and a feast for birds (Figure 42). Those females that survive deposit their eggs, often among mosses.

Figure 39. Platyhypnidium riparioides partially submersed at the edge of a waterfall. Photo by Michael Lüth, with permission.

Figure 42. Hermit thrush (Catharus guttatus) with mayfly subimago in its beak, enjoying the brief period of emergence. Photo by Bob Armstrong, with permission.

Chapter 11-4: Aquatic Insects: Hemimetabola – Collembola and Ephemeroptera

Increased biomass of bryophytes may increase some insects while having no effect on others. Lee and Hershey (2000) found that a dense growth of the moss Hygrohypnum (Figure 43-Figure 44) following stream fertilization in Alaska increased the density of the mayfly Ephemerella aurivillii (Figure 45) but not Baetis (Figure 46). In the fertilized zone, these mayflies both grew larger, a fact Lee and Hershey attributed to the greater growths of epiphytic diatoms. Furthermore, although the density of Ephemerella increased with increased moss density, the highest drift ratios were in the unfertilized zone with lower moss density. In enclosure experiments, they found that bare rock, mosses, and artificial mosses had no effect on any taxa except Ephemerella. They considered that the Ephemerella benefitted from the increased complexity of the moss habitat.

Figure 43. Hygrohypnum ochraceum, home for a variety of stream insects. Photo by Michael Lüth, with permission.

Figure 44. Close view of Hygrohypnum ochraceum, home for a variety of insects. Photo by Michael Lüth, with permission.

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Figure 45. Ephemerella aurivillii naiad, a mayfly that increased with increased coverage of Hygrohypnum in Alaska. Photo by Tom Murray, through Creative Commons.

Figure 46. Baetis naiad, a bryophyte inhabitant in many streams. Photo by Bob Henricks, with permission.

Jones (1950) did extensive gut analysis of insects from the River Rheidol. Among the Ephemeroptera, none of the five species examined had fragments of the common moss Fontinalis antipyretica (Figure 47) in the gut. Detritus was the most common food. Gilpin and Brusven (1970) found six mayfly species with Fontinalis sp. in their guts, but these all amounted to less than 1% of the gut contents.

Figure 47. Fontinalis antipyretica, a moss found in the guts of some mayflies in the River Rheidol. Photo by Kristian Peters, with permission.

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Chapter 11-4: Aquatic Insects: Hemimetabola – Collembola and Ephemeroptera

It is surprising to find such flattened, rock-adapted genera as Heptagenia (Figure 48) among mosses, but Muttkowski and Smith (1929) did find it several times among mosses in trout streams of Yellowstone National Park, USA.

Macan (1957) found Leptophlebia (Figure 50) among mosses in Ford Wood Beck, UK. Berner (1959) described this genus as one that would live in submerged mossy banks and other quiet areas. The genus is negatively phototactic (movement of organism toward or away from source of light), explaining their presence in the secluded shade of streambank mosses. When it is time for the naiads to emerge into adults, they become positively phototactic and crawl upward onto sticks, logs, or other protruding structure, probably including emergent bryophytes. Vuori et al. (1999) considered Leptophlebia marginata (Figure 50) to be among the dominant moss dwellers in the Tolvajärvi region of the Russian Karelia. Bengtsson (1981) found that L. marginata demonstrated a steady growth rate throughout winter, permitting it to thrive in such northern regions.

Figure 48. Heptagenia dalecarlica naiad, a flattened species adapted for smooth rocks, but that occasionally visits mosses. Photo by Urmas Kruus, with permission.

Suborder Furcatergalia Leptophlebiidae – Prong-gilled Mayflies This is a family that lives in freshwater streams and lakes where the naiads eat detritus and algae (Leptophlebiidae 2013). Their length is up to 20 mm; they are nocturnal (active at night) and are poor swimmers, generally clinging to rocks. Only a few seem to live among bryophytes. Paraleptophlebia (Figure 49) was a minor component of the bryophyte communities in my own Appalachian, USA, stream studies (Glime 1968). Maurer & Brusven (1983) found Paraleptophlebia heteronea (Figure 49) frequently in the clumps of Fontinalis neomexicana (Figure 79) in an Idaho stream. In their study of four Appalachian streams, Woodall and Wallace (1972) found this genus where there was moderate or slow current among decaying leaves, bark, and wood. Its food is predominately detritus (Chapman & Demory 1963).

Figure 50. Leptophlebia marginata naiad on waterweed. Photo by Niels Sloth, with permission.

Figure 49. Paraleptophlebia sp. naiad, a frequent dweller among Fontinalis neomexicana. Photo by Jason Neuswanger, with permission.

Figure 51. Sphagnum affine, member of a genus that contributes H+ ions, lowering the pH of bogs and their outflow waters. Photo by Michael Lüth, with permission.

One advantage enjoyed by some members of this family is tolerance of somewhat low pH. Mayflies in general are indicators of fresh, unpolluted water. They do not generally tolerate extremes, low pH included (Raddum & Fjellheim 1988; Raddum et al. 1988; Braukmann 1992; Lingdell & Engblom 1995). Thus the streams that drain Sphagnum fens and bogs (Figure 51) are generally depauperate (lacking in numbers or kinds of species) of mayflies. However, this habitat is suitable for a few, including Leptophlebia vespertina (Figure 52) (Bauernfeind & Moog 2000). This intolerance of low pH may explain its relative rarity among bryophytes in the mid-Appalachian Mountain streams (Glime 1968).

Chapter 11-4: Aquatic Insects: Hemimetabola – Collembola and Ephemeroptera

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In his study of the River Rajcianka, Krno (1990) found a genus I have not encountered elsewhere – Habroleptoides. Habroleptoides modesta (Figure 55) is a bryophyte dweller in the river, but like many of the mayfly genera, it is unable to live among the wet mosses above the water level.

Figure 52. Leptophlebia vespertina adult, a species whose naiads can inhabit the acid outflows of acid bog lakes. Photo by Niels Sloth, with permission.

In New Zealand Austroclima sepia (see Figure 53) frequently lives among mosses in small waterfalls (Winterbourn & Gregson 1981). Similarly, Towns (1987) reported this species along with A. jollyae and Mauiulus luma (Figure 54) as 72%, 13%, and 9%, respectively, of the fauna from mosses in rapid flow (where only 4 insect species lived!) on the Great Barrier Island, New Zealand.

Figure 55. Habroleptoides modesta naiad, a mayfly that sometimes lives among bryophytes in rivers. Photo by Alfeo Busilacchio, with permission.

Caenidae - Small Squaregill Mayflies The Caenidae are small sprawlers in quiet and sometimes stagnant water as well as streams (Caenidae 2014). They are adapted to the relatively low oxygen of silt. Caenis (Figure 56) seems to prefer loose mosses (Percival & Whitehead 1929). Frost (1942) found that it was most likely to occur among mosses that had accumulated considerable silt. In the River Rajcianka in Slovakia, Caenis beskidensis (Figure 56) lives among submerged bryophytes but is not found, like some mayflies, among the wet emergent bryophytes (Krno 1990). In the Appalachian Mountain, USA, streams naiads of Caenis were among the lesser of the moss inhabitants, appearing mostly among Fontinalis dalecarlica (Figure 69). Figure 53. Austroclima naiad, a genus with moss dwellers in New Zealand. Photo by Stephen Moore, Landcare Research, NZ, with permission.

Figure 56. Caenis lactea naiad, a mayfly that prefers loose mosses. Photo by Niels Sloth, with permission.

Neoephemeridae Figure 54. Mauiulus luma naiad, a mayfly that lives among mosses in small waterfalls in New Zealand. Photo by Stephen Moore, Landcare Research, NZ, with permission.

The rare genus Neoephemera (Figure 57) sometimes lives deep within submerged moss mats in rapid water in eastern North America (Berner 1959), including

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Chapter 11-4: Aquatic Insects: Hemimetabola – Collembola and Ephemeroptera

Neoephemera compressa (Figure 57) among mosses on submersed parts of trees (Berner 1956). The naiad moves slowly, but when it bends its 3 tails over its abdomen, then suddenly lashes them back, this action propels it forward (see Figure 60).

Figure 57. Neoephemera compressa, an inhabitant of mosses on submersed parts of trees. Photo by Dana R. Denson, Florida Association of Benthologists, with permission.

Figure 58. Ephemerella subvaria naiad gill covers, closed over gills. Photo by Tom Murray, through Creative Commons.

In Australia, Neoephemera (Figure 57) naiads live in protected parts of streams with slow to moderate flow where they hide among debris, plant roots, and mosses (Edmunds et al. 1976). These naiads are difficult to dislodge from the mosses, partly because they grip the mosses. The membranous respiratory gills are fragile and they need the protection that is provided by the fused, sclerotized opercula (gill covers) (Notestine 1994). This genus relies heavily on these gills for respiration. Ephemerellidae – Spiny Crawlers This family occurs throughout North America as well as the United Kingdom (Ephemerellidae 2014). These collector-gatherers occur where there is moving water, including lake shores subject to wave action, but seem to require reduced flow. They are able to live in fast water by accepting the protection of bryophytes. When these mayfly naiads are threatened by a predator, they raise their three tails like a scorpion, arching them up and over their backs, making them appear larger (Ephemerellidae 2014). They will then project the tails forward to poke the enemy. Spines on the back of the abdomen (Figure 58) may contribute to their protection. One suggestion is that the spines help the mayflies hold their positions when attacked from behind by a predator. This family takes advantage of the protection of the bryophyte habitat while modulating the oxygen and keeping its tuft of gills clean with its gill covers. When oxygen concentrations become too low, the Ephemerellidae move the gill covers (Figure 58) up and down to keep fresh water circulating across the gills (Figure 59) (Ephemerellidae 2014). Their bodies are somewhat flattened dorsiventrally and are adapted to crawling among the chambers of their mossy habitat. When they are in open water and need to move quickly, mayflies in this family flip their tails upward over their backs and down to act like a paddle (Figure 60), thrusting them forward.

Figure 59. Drunella sp. naiad with gill covers up to expose the tufts of gills. Photo by Bob Henricks, with permission.

Figure 60. Ephemerella subvaria naiad in a swimming position with its tails flipped upward. Photo by Bob Henricks, with permission.

Berner (1959) described some members of this family as living on the tops of rocks, deep within the moss. Arnold and Macan (1969) found that Ephemerellidae (Figure 58-Figure 64) were common among mosses in a

Chapter 11-4: Aquatic Insects: Hemimetabola – Collembola and Ephemeroptera

Shropshire Hill stream in the UK. In a study of the McKenzie River, Oregon, USA, Hawkins (1984) reported that 5 species [Serratella teresa, C. hystrix (Figure 61), Caudatella cascadia (now a synonym of C. hystrix), C. edmundsi (Figure 62), and Drunella spinifera (Figure 63)] out of 12 Ephemerellidae species were common among mosses, including Fontinalis sp. (Figure 79) and others. Gilpin and Brusven (1970) likewise found C. edmundsi among clumps of Fontinalis. Hawkins (1984) found those restricted to mosses were usually at upstream locations where the mosses were abundant. However, two moss dwellers [Caudatella edmundsi (100% moss usage - found only on Fontinalis), Drunella spinifera (54%)] were most abundant downstream, living among mats of the moss Fontinalis sp. For other species with more than 5% use of bryophyte habitats he found Serratella teresa (85%), Caudatella cascadia (46%), and Caudatella hystrix (22%). Brittain and Saltveit (1989) found that river impoundments had "profound" effects on the Ephemerellidae (Figure 58-Figure 64) living there. Changes in temperature, discharge, flow patterns, food availability, and predator density all contribute to changes in living conditions for the mayflies. Increased growth of mosses and additional available substrata for periphyton below the dams often favor some of the Ephemerellidae while reducing suitable habitat for Heptageniidae (Figure 48). The mayflies living under these changeable regimes often have flexible life cycles or shorter periods of rapid growth with a long period of egg development that permit them to survive unsuitable periods.

Figure 61. Caudatella hystrix naiad, a common moss dweller in the McKenzie River, Oregon, USA. Photo by Bob Newell, with permission.

Figure 62. Caudatella edmundsi naiad, a common moss dweller. Photo by Bob Newell at , with permission.

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Figure 63. Drunella spinifera naiad. Photo by Bob Newell at , with permission.

Percival and Whitehead (1929) considered mosses and algae to be the main food of the Ephemerellidae (Figure 58-Figure 64). Woodall and Wallace (1972) found Eurylophella funeralis (=Ephemerella funeralis, Figure 64) to be the most abundant Ephemerella species among mosses in the southern Appalachian Mountains, USA, and I found a similar relationship for E. funeralis and E. temporalis in the middle Appalachian Mountain streams (Glime 1968). The members of Ephemerella tended to avoid the heavily shaded hardwood stream where mosses and algae were scarce.

Figure 64. Eurylophella funeralis, a common mayfly among mosses in the southern Appalachian Mountain, USA, streams. Photo by Donald S. Chandler, with permission.

Brittain and Saltveit (1989) found that growth of mosses and associated periphyton below dams favored presence of Ephemerellidae (Figure 58-Figure 64). They reasoned that flexible life cycles permitted them to survive adverse conditions, including rapid nymphal growth and long period of egg development. Eggs typically form a ball (Figure 65). Percival and Whitehead (1929) found Eurylophella funeralis (=Ephemerella funeralis) (Figure 64) to be the most abundant species of the Ephemerella genus group in their study of UK streams. The main foods of Ephemerella species are algae and mosses (Percival & Whitehead 1929; Jones 1949, 1950; Gerson 1969). This is convenient because this genus is common among mosses, but it also occurs on the pebbles on the bottom. Jones (1949, 1950) found that Ephemerella s.l. fed primarily on Fontinalis (Figure 47) and the alga Ulothrix (Figure 66) in calcareous (having dissolved chalk or limestone) streams of South

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Chapter 11-4: Aquatic Insects: Hemimetabola – Collembola and Ephemeroptera

Wales. Among 14 specimens examined on 14 July the moss was the primary food, but they concluded that Ephemerella feeds on Ulothrix when it is abundant but switches to Fontinalis antipyretica (Figure 47) when the Ulothrix becomes scarce.

The family Ephemerellidae (Figure 58-Figure 64) seems to have bryological preferences, or preferences that match those of the bryophytes. They reach extremely high numbers among Hygroamblystegium fluviatile (Figure 68) in mid-Appalachian streams, but are nearly absent in Fontinalis dalecarlica (Figure 69) and Scapania undulata (Figure 70) in different streams (Glime 1968).

Figure 65. Ephemerella egg mass with debris stuck to it. Photo by Jason Neuswanger at , with permission.

Figure 68. Hygroamblystegium fluviatile, home to large numbers of Ephemerellidae. Photo by Michael Lüth, with permission.

Figure 66. Ulothrix, food for Eurylophella funeralis. Photo by Yuuji Tsukii, with permission.

Reproduction in the mayflies involves swarming, a behavior that maximizes contact of males and females that typically live for only one day as adults. In Serratella ignita (Figure 67) this swarming occurs in the late afternoon and evening (Elliott & Humpesch 1980). The egg mass is a greenish ball. Once fertilized, eggs are laid in turbulent water, usually where there are mosses. The female flies upstream to deposit the eggs on the water surface. She then usually falls on the surface and is vulnerable to fish predation. The egg mass separates when it enters the water and each egg attaches to the substrate with its polar anchoring cap.

Figure 67. Serratella ignita naiad. Photo by J. C. Schou, through Creative Commons.

Figure 69. Fontinalis dalecarlica, a stream moss that houses some of the larger insects. Photo by Jan-Peter Frahm, with permission.

Figure 70. Scapania undulata, a leafy liverwort that has few of the typical moss-dwelling Ephemerellidae. Hermann Schachner, through Creative Commons.

Chapter 11-4: Aquatic Insects: Hemimetabola – Collembola and Ephemeroptera

D. N. Bennett (pers. comm. 19 April 2011) described her field experience with an aquatic entomologist, Bob Henricks. Henricks was attempting to distinguish between mosses and grasses, so she began looking at the inhabitants of the mosses. When the moss-covered rocks were removed from the stream, the insects began moving about and became more noticeable. There were often 40-50 Ephemerellidae naiads on a single moss-covered rock – determined to be Hygroamblystegium, probably H. tenax (Figure 71-Figure 72). The moss grew on and "under" the rock, and it was the submersed "under" portion that housed the many mayflies. She observed the naiads rolling up the algae from the moss leaf surface, starting at the leaf tip and moving to the stem.

Figure 71. Hygroamblystegium tenax in a dry stream bed. Photo by Janice Glime.

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For example, Serratella ignita (Figure 1) has an annual cycle with the eggs spanning the winter in a dormant state, hatching in April and May in the River Endrick in Scotland (Maitland 1955). The naiads develop quickly, emerging in July and August, and adults typically lay eggs within 24 hours of emergence. These eggs are often laid among mosses in abundance (Percival & Whitehead 1928). The eggs are laid in evening light and are caught by Platyhypnidium riparioides (Figure 39) and Fontinalis species (Figure 47) where they adhere as a greenish gelatinous mass. In a Shropshire Hill stream in the UK, Arnold and Macan (1969) found that the longest stage in Serratella ignita (Figure 1) was the egg, a stage that remained from late summer one year to late spring the next year, hence overwintering as an egg (Elliott 1967). Rosillon (1988) found that completion of naiad development on a diatom diet required about 950 degree-days above a temperature of 3.5°C (range 9.5-18°C). [Degree days for insect development can be calculated by adding the minimum and maximum temperature of the day and dividing by 2. The minimum required for development is subtracted from that number to determine how many degree-days have been added that day. (Townsend et al. 2010)]. Those reared on detritus rarely achieved adult stage. Rosillon suggested that poor food quality would reduce fecundity (reproductive rate) of females. Furthermore, it appears that under ideal conditions Serratella ignita could have a bivoltine (2 broods per year) life cycle. Emergence patterns can be gleaned from the stages of the naiad development of mayflies in samples. Based on such sampling, Gurtz & Wallace (1984) estimated that in a stream in the southern Appalachian Mountains, USA, the moss inhabitants Ephemerella catawba (Figure 73) probably emerged from May to July, E. hispida from April to June, E. excrucians (Figure 81) in May and June, and Drunella tuberculata (Figure 74) from June to September. Both Ephemerella catawba and Ephemerella invaria occurred among mosses in the acidic mid Appalachian streams in my own studies (Glime 1968). Ephemerella invaria (Figure 75) increased in Big Hurricane Branch following a clearcut, but no specimens with fully developed wing pads were ever collected, suggesting that nymphs of this species might complete their development farther downstream in Shope Creek (Gurtz & Wallace 1984).

Figure 72. Hygroamblystegium tenax, home to many kinds of stream insects, including Ephemerellidae. Photo by Jan-Peter Frahm, with permission.

Seasons Seasonal differences in the life cycle stages spent in the water are often the key to success for these species. Timing differences in emergence times and hatching times can separate realized niches in closely related species. In the Ephemerellidae (Figure 58-Figure 64), the life cycle is typically one year with one brood per year (univoltine).

Figure 73. Ephemerella catawba, a moss inhabitant as a naiad that emerges May to July in the southern Appalachian Mountains, USA. Photo by Biodiversity Institute of Ontario, through Creative Commons.

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Chapter 11-4: Aquatic Insects: Hemimetabola – Collembola and Ephemeroptera

contributed to the improved growth rates, with the mosses serving as traps for seston (swimming or floating living organisms and non-living matter) being released from the reservoir. Both of these species occur among bryophytes in streams of the mid Appalachian Mountains, USA (Glime 1968).

Figure 74. Drunella tuberculata, a summer emerger. Photo by Bob Henricks, with permission.

Figure 76. Ephemerella subvaria naiad. Photo by Donald S. Chandler, through Creative Commons.

Food

Figure 75. Ephemerella invaria naiad. Henricks, with permission.

Photo by Bob

Ephemerella invaria (Figure 75) occurred both above and below a hydroelectric plant on the Sturgeon River in northern Michigan, USA, with similar abundance and growth (Mundahl & Kraft 1988). Ephemerella subvaria (Figure 76) naiads were 4x as abundant below the plant (136 m-2 below vs. 33 m-2 above), but grew more slowly there. Nevertheless, the growth rate increased with distance downstream from the power plant for nearly 10 km. Extensive beds of Fontinalis (pers. obs.) may have

The Ephemerellidae (Figure 58-Figure 64) are the most commonly reported mayflies among the bryophyte consumers (Table 1). Caudatella hystrix (as C. cascadia; Figure 61) varies its diet depending on the site (Coffman et al. 1971; Hawkins 1985). Detritus is important in its diet, but the proportion decreases when that of moss increases (Hawkins 1985). The naiads of Caudatella edmundsi (Figure 62, Figure 101) feed primarily on diatoms, but also include detritus and mosses in their diet. Hawkins found that as size increased in the Ephemerellidae, especially in Caudatella edmundsi and Ephemerella dorothea infrequens (Figure 80), the consumption of both animal matter and mosses increased. Hawkins found that eight species demonstrated a correlation between moss consumption and size. López-Rodríguez et al. (2008) likewise found that the proportion of mosses in the diet increases in Ephemerellidae as naiads age. Several researchers (Hynes 1941; Chapman & Demory 1963; Gaevskaya 1969) found that mosses are eaten by members of this family more often than other aquatic macrophytes (not including algae). But it is not clear if the moss is eaten for its own food value or for the attached periphyton. Percival and Whitehead (1929) found that two species in this family ingested large amounts of moss, suggesting that the moss itself was an important food source. Among the members of Ephemerellidae studied by Hawkins (1985), Caudatella edmundsi, C. heterocaudata, C. hystrix, and Serratella teresa were moss shredders. Others living among the mosses and ingesting them were detritus shredders, including Attenella margarita (Figure 77), Ephemerella dorothea infrequens, E. excrucians (Figure 81), E. velmae, Serratella tibialis (Figure 84), and Timpanoga hecuba (Figure 78). Drunella pelosa is a diatom scraper, permitting it to eat the many diatoms adhering to the moss leaves.

Chapter 11-4: Aquatic Insects: Hemimetabola – Collembola and Ephemeroptera

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Table 1. Correlations between size (mm) and percent composition of major food items in the gut. Values are correlation coefficients (r). * = P larva > pupa > adult. These insects typically spend only part of the life cycle in the water. Some lay their eggs near water and larvae develop in the water. Some have eggs, larvae, and pupae in the water, but their emerging adults break through the water surface and climb onto land to emerge. For most, adult life and mating occur on land.

mosses (Elliott et al. 1996) near water, laying about 30 eggs either singly or in pairs. Larvae leave the egg site within 1-3 days to burrow into mosses. Larvae may live in or out of water, but pupation is on land, lasting 7-18 days. If the larvae are submersed, they crawl out of the water (Ward 1965). If the moss is submersed, they burrow deeply into it, but within 8-28 days of submersion they die. Adults live two weeks to three months, depending on species and location.

NEUROPTERA – Net-winged Insects Neuroptera literally means nerve wings, so-named because of the prominent wing veins of the adults. This order is not well represented among bryophytes, and only the larvae are associated with aquatic habitats.

Osmylidae On continents other than North America a small family, the Osmylidae (Figure 2-Figure 6), occurs among mosses and organic matter in and near streams (Flint 1977). Osmylus fulvicephalus (Figure 2) is the only species known in the UK, likewise living among mosses of streambanks (Elliott et al. 1996) and seeking food there (NatureSpot 2015). The adults (Figure 3; 25 mm long including wings) don't stray far from water but are not aquatic. The females lay their eggs on overhanging plants, tree trunks, or stones (Osmylidae 2014), and especially on

Figure 2. Osmylus fulvicephalus larva, a species that lives among mosses on streambanks and feeds there. Photo by Walter Pfliegler, with permission.

Chapter 11-8: Aquatic Insects: Holometabola – Neuroptera and Megaloptera

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Like Osmylus fulvicephalus (Figure 2-Figure 5), Kempynus sp. (Figure 6) in the Southern Alps of New Zealand is somewhat amphibious, living at the edge between water and land (Cowie & Winterbourn 1979). In springbrooks it lives in clumps of the mosses Acrophyllum quadrifarium (=Pterygophyllum quadrifarium; Figure 7) and Cratoneuropsis relaxa (Figure 8).

Figure 3. Osmylus fulvicephalus adult that lays its eggs on overhanging vegetation. Larvae live among streambank mosses. Photo through Creative Commons.

Figure 4. Phenological events (cyclic and seasonal natural phenomena, especially in relation to climate) of the life cycle of Osmylus fulvicephalus. From Elliott et al. 1996.

Osmylus fulvicephalus (Figure 3) is controversial in that its larvae live in wet mosses, but drown in 8-28 days of submersion (Elliott et al. 1996). Nevertheless, they do enter the water in search of food. It seems safe to say, however, that their relationship with mosses is damp, but not aquatic. The larva feeds among these mosses. When movement is detected, it jabs at it with the long proboscis, then injects it with a salivary secretion that paralyzes it. A chironomid larva is paralyzed within 10 seconds. The O. fulvicephalus then sucks out the interior of the prey. The larvae stop eating during mid autumn and burrow down to the moss rhizoids to hibernate for the winter. Fortunately, in this state they can survive occasional submersion in water, thus surviving spates (sudden flood in a river, especially one caused by heavy rains or melting snow). In spring they spin a silken cocoon, sometimes incorporating bits of moss in the cocoon. Just before pupation the long jaws break off (Figure 5). The pupa becomes immobile during pupation. It grows a pair of mandibles that it uses to cut its way out of the cocoon.

Figure 5. Osmylus fulvicephalus larva showing large jaws. Photo by Walter Pfliegler, with permission.

Figure 6. Kempynus sp larva, member of the small family Osmylidae that inhabits mosses near streams. Photo by Stephen Moore, Landcare Research NZ, with permission.

Figure 7. Pterygophyllum quadrifarium, a moss habitat for Kempynus sp. at stream borders and in springbrooks in New Zealand. Photo by Bill and Nancy Malcolm, with permission.

Figure 8. Cratoneuropsis relaxa, a moss habitat for Kempynus sp. at stream borders and in springbrooks in New Zealand. Photo by Tom Thekathyil, with permission.

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Chapter 11-8: Aquatic Insects: Holometabola – Neuroptera and Megaloptera

Chrysopidae There are a number of reports of the larvae of the green lacewing Leucochrysa pavida (Figure 9-Figure 12) using bits of lichen as camouflage (Tauber et al. 2009; Moskowitz & Golden 2012). In fact, Wilson and Methven (1997) found that the larvae at their Illinois, USA, site were somewhat specific in the species of lichens they chose. But Slocum and Lawrey (1976) found that this insect was not totally specific. In addition to the lichens, it also includes pieces of bark, angiosperm pollen, fungal spores, insect debris, and (of course) bryophyte gametophytes. Slocum and Lawrey demonstrated that the lichens, at least, are still alive and that they have photosynthetic rates equal or greater than those same lichen species still growing on a bark substrate. Furthermore, these lichen propagules are still viable when the cocoons are attached to the bark, giving the lichens the opportunity and establish in this new location. Unfortunately, there are no similar studies on the bryophytes in this camouflage arrangement, but it at least provides the possibility for a means of dispersal. Figure 11. Leucochrysa pavida larva showing ventral side. Photo by Jim McCormac, with permission.

Figure 9. Leucochrysa pavida larva with lichen back pack, showing its camouflage against tree bark lichens. Photo by Jim McCormac, with permission.

Figure 12. Leucochrysa pavida larva showing head and large mandibles of this carnivore. Photo by Jim McCormac, with permission.

MEGALOPTERA Alderflies Figure 10. Leucochrysa pavida larva with lichen back pack, showing the legs and mandibles of the larva. Photo by Jim McCormac, with permission.

–

Dobsonflies

and

Megaloptera means large wing; one adult is known with a wingspan of 21 cm, the largest of any aquatic insect in the world (Megaloptera 2014). The order is relatively small, and is close to the Neuroptera. Its members have

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aquatic larvae, but they pupate on land in damp soil or under logs. The pupae are fully mobile and can defend themselves against predators with their large mandibles. Female adults lay 1000's of eggs on overhanging vegetation where larvae can drop into the water (Figure 13). The adults often live only a few hours and usually don't eat.

Figure 15. Sialis lutaria larva, the aquatic stage that migrates into the water, sometimes from streamside bryophytes. Photo by André Karwath, through Creative Commons.

Figure 13. Sialis fuliginosa eggs. Lindsey, with permission.

Photo by James K.

Sialidae – Alderflies This is a small family that can be up to 25 mm long (Alderfly 2014). They occur sparsely worldwide with a concentration of known species in Europe (Sialidae 2015). I have only found reference to one genus of bryophyte dwellers, Sialis (Figure 13-Figure 17) (Lithner et al. 1995). I likewise found this genus occasionally among bryophytes in Appalachian Mountain, USA, streams (Glime 1968). It has aquatic larvae, but adults are terrestrial and lay eggs near water (Alderfly 2014). Fully grown larvae of Sialis pupate in soil, mosses, under stones, and other locations, usually near water. In Canada, after about one month the adults appear. Sialis nigripes prefers mosses for egg laying (Elliott et al. 1996). Sialis lutaria (Figure 15-Figure 17) was used in a study comparing heavy metal accumulation in mosses (Fontinalis spp.; Figure 18), insects, and fish (Lithner et al. 1995).

Figure 16. Sialis lutaria adult. Photo ©entomart, through Creative Commons.

Figure 17. Sialis lutaria adults mating. Photo by James K. Lindsey, with permission.

Figure 14. Sialis adult, a genus that sometimes pupates and lays eggs among streamside bryophytes. Photo by Patrick Coin, through Wikimedia Commons.

On the South African Cape, pupae of Sialidae along streams or waterfalls live in Sphagnum (Figure 19) and other mosses (Barnard 1931). These pupae require a wet, but not submersed, habitat, so the mosses must be soaking wet.

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Figure 20. Nigronia serricornis larva showing powerful jaws. The aquatic larva often crawls into mosses to pupate. Photo by Jason Neuswanger, with permission. Figure 18. Fontinalis antipyretica, home to numerous kinds of insects and useful for comparing heavy metal accumulation. Photo by Malcolm Storey, through Creative Commons.

Figure 21. Nigronia serricornis adult. Pupae of this insect often reside in mosses. Photo by Phil Myers, through Creative Commons.

Figure 19. Sphagnum fimbriatum, a genus that lives in Africa and is a potential home for pupae of Sialidae. Photo by Blanka Shaw, with permission.

Corydalidae- Dobsonflies and Fishflies This family occurs mostly in the Northern Hemisphere and in South America, including both temperate and tropics (Corydalidae 2014). Their body size is usually greater than 25 mm and ranges up to 80 mm (Penny et al. 1997; Bartlett 2004). The larvae are aquatic, are called hellgrammites, and are predators. Nigronia, an aquatic member of the Corydalidae, is not typically a moss inhabitant, although I did occasionally find larvae of this genus among Appalachian Mountain stream bryophytes (Glime 1968). But like many other aquatic insects, Nigronia serricornis (Figure 20-Figure 21) pupates among mosses as well as under stones and logs (Needham et al. 1901). Likewise, Chauliodes pectinocornis (Figure 22) and C. rastricornis (Figure 24Figure 24) pupate in these habitats. Pupation lasts about 2 weeks in these Corydalidae.

Figure 22. Chauliodes pectinicornis adult, a species that lives in the water as larvae and pupates among mosses. Photo by Stephen Cresswell, with permission.

Figure 23. Chauliodes rastricornis larva, a species that may move to mosses to pupate. Photo by Tom Murray, through Creative Commons.

Chapter 11-8: Aquatic Insects: Holometabola – Neuroptera and Megaloptera

Figure 24. Chauliodes rastricornis adult, a species that lives in the water as larvae and pupates among mosses. Photo by Stephen Cresswell, with permission.

Summary The Holometabola have a complete life cycle with egg, larva, pupa, and adult. The Neuroptera are represented among aquatic bryophytes by only one family, the Osmylidae. The larvae of Osmylus may live among bryophytes in streams or on streambanks and obtain food there. Some species lay their eggs on mosses that overhang streams. Larvae bore into mosses in or out of the water. Kempynus species often live among mosses in springbrooks. The Megaloptera, like the Neuroptera, have few aquatic bryophyte dwellers. Sialis (Sialidae) larvae occasionally occur among stream bryophytes; the pupae are often among terrestrial mosses. Some species lay eggs among mosses. Wet Sphagnum along streams or near waterfalls serves as a home for some Sialidae. Some members of Nigronia and Chauliodes, both in the Corydalidae, pupate among mosses.

Acknowledgments I appreciate the availability of images in Creative Commons and the family information available through BugGuide, Wikipedia, and EOL. Eileen Dumire reviewed the chapter from the perspective of a lay person and checked for grammatical errors.

Literature Cited Alderfly. 2015. Accessed 19 January 2015 at . Barnard, K. H. 1931. The Cape alder-flies: (Neuroptera, Megaloptera.). Trans. Royal Soc. S. Afr. 19: 169-184. Bartlett, Troy. 2004. Corydalidae. BugGuide. Accessed 19 January 2015 at .

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Corydalidae. 2015. Wikipedia. Accessed 19 January 2015 at . Cowie, B. and Winterbourn, M. J. 1979. Biota of a subalpine springbrook in the Southern Alps. N. Z. J. Marine Freshwat. Res. 13: 295-301. Elliott, J. M., Kimmins, D. E., and Worthington, C. J. 1996. British Freshwater Megaloptera and Neuroptera: A Key with Ecological Notes. Freshwater Biological Association, Cumbria, 68 pp. Flint, O. S. Jr. 1977. Neuroptera. In: Hurlbert, S. H. (ed.). Biota Acuatica De Sudamerica Austral., San Diego State University, pp. 187-188. Glime, J. M. 1968. Aquatic Insect Communities Among Appalachian Stream Bryophytes. Ph.D. Dissertation, Michigan State University, East Lansing, MI, 180 pp. Lithner, G., Holm, K., and Borg, H. 1995. Bioconcentration factors for metals in humic waters at different pH in the Roennskaer area (N. Sweden). In: Grennfelt, P., Rodhe, H., Thoerneloef, E., and Wisniewski, J. (eds.). Acid Reign '95? Proceedings from the 5th International Conference on Acidic Deposition: Science and Policy, held in Goteborg, Sweden, 26-30 June 1995. Water Air Soil Pollut. 85: 785-790. Megaloptera. 2014. Wikipedia. Last updated 26 July 2014. Accessed 31 August 2014 at . Moskowitz, D. and Golden, D. 2012. First Records of the green lacewing Leucochrysa pavida (Hagen) (Neuroptera: Chrysopidae) in New Jersey. Entomol. News 122(1): 55-58. NatureSpot. 2015. Accessed 19 January 2015 at . Needham, J. G., Betten, C., MacGillivray, A. D., Coquillett, D. W., and Ashmead, W. H. 1901. Aquatic Insects in the Adirondacks. N. Y. State Mus. Bull. 47: 1-612. Osmylidae. 2014. Australian Freshwater Invertebrates. Accessed 31 August 2014 at . Penny, N. D., Adams, P. A., and Stange, L. A. 1997. Species catalog of the Neuroptera, Megaloptera, and Raphidioptera of America North of Mexico. Proc. Calif. Acad. Sci. 50: 39114. Sialidae. 2015. Encyclopedia on Line. Accessed 19 January 2015 at . Slocum, R. D. and Lawrey, J. D. 1976. Viability of the epizoic lichen flora carried and dispersed by green lacewing (Nodita pavida) larvae. Can. J. Bot. 54: 1827-1831. Tauber, M. J., Tauber, C. A., and Albuquerque, G. S. 2009. Neuroptera (Lacewings, Antlions), pp. 695-707. In: Resh, V. H. and Cardé, R. (eds.). Encyclopedia of Insects, 2nd Edition. Academic Press, San Diego, 1132 pp. Ward, P. H. 1965. A contribution to the knowledge of the biology of Osmylus fulvicephalus (Scopoli 1763) (Neuroptera, Osmylidae). Entomol. Gaz. 16: 175-182. Wilson, P. J. and Methven, A. S. 1997. Lichen use by larval Leucochrysa pavida (Neuroptera: Chrysopidae). Bryologist 100: 448-453.

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Glime, J. M. 2017. Aquatic Insects: Holometabola – Coleoptera, Suborder Adephaga. Chapt. 11-9. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. Ebook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 19 July 2020 and available at .

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CHAPTER 11-9 AQUATIC INSECTS: HOLOMETABOLA – COLEOPTERA, SUBORDER ADEPHAGA TABLE OF CONTENTS COLEOPTERA BACKGROUND ........................................................................................................................... 11-9-2 Suborder Adephaga ........................................................................................................................................... 11-9-4 Carabidae – Ground Beetles....................................................................................................................... 11-9-4 Gyrinidae – Whirligig Beetles ................................................................................................................... 11-9-5 Haliplidae – Crawling Water Beetles ......................................................................................................... 11-9-5 Hygrobiidae – Squeak Beetles ................................................................................................................... 11-9-6 Dytiscidae – Predaceous Diving Beetles and Noteridae – Burrowing Water Beetles ................................ 11-9-6 Moors, Bogs, and Fens........................................................................................................................ 11-9-8 Summary ......................................................................................................................................................... 11-9-13 Acknowledgments ........................................................................................................................................... 11-9-13 Literature Cited ............................................................................................................................................... 11-9-13

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CHAPTER 11-9 AQUATIC INSECTS: HOLOMETABOLA – COLEOPTERA, SUBORDER ADEPHAGA

Figure 1. Lancetes angusticollis adults on moss, South Georgia in the Antarctic. Lancetes angusticollis has a two-year life cycle, with overwintering possible in three life stages – aquatic larvae, terrestrial pupae (not proven), and aquatic adults. Note the air supply at the tip of the abdomen. This external air supply makes it necessary for these beetles to cling to vegetation, when they are not swimming, to avoid floating to the surface, hence their use of mosses. Photo by Roger S. Key, with permission.

COLEOPTERA BACKGROUND The Coleoptera seem to have a somewhat closer relationship to terrestrial life than other aquatic bryophyte dwellers. First of all, they get their air from the atmosphere or underwater plants where they grab an air bubble (Figure 2). They can accumulate air as bubbles under the elytra (hardened forewings; wing covers), through the plastron (breast plate breathing apparatus; Figure 3) (Oliveira de Sousa et al. 2012), or an anal bubble. The plastron is a ventral structure that acts as a physical gill by using various combinations of hairs, scales, and undulations projecting from the cuticle. This apparatus holds a thin layer of air along the outer surface of the body (Figure 3). In all three

of these mechanisms, the nitrogen in the air bubble diffuses into the water slowly while the replacement oxygen diffuses into it 2-3 times as fast (Rich Merritt, pers. comm. 28 January 2015). Thus, as the insect uses up the oxygen from the bubble, the water replaces it by oxygen diffusion for a reasonable period of time. The CO2 from respiration enters the bubble and rapidly diffuses into the water, having little effect on bubble size. Many beetles attach an anal gas bubble (Figure 1, Figure 18-Figure 19) that uses this diffusion mechanism. They may have hairs that help hold the bubble in place. (See Elmidae in Coleoptera, Suborder Polyphaga, for details of the plastron functioning in that family.)

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Percival and Whitehead (1930) noted that the mosses in streams in the UK were very important to both larvae and adults of the small Coleoptera. In 1949, Badcock indicated that beetles were more common among mosses than associated with stones, especially loose stones. Ogbogu (2000) found Coleoptera among the insects associated with Fontinalis (Figure 5) in an intermittent reservoir spillway in Ile-Ife, Nigeria. Many of the Coleoptera in rivers of northwest Spain prefer moss as a substrate, as indicated by both species richness and abundance (Fernández-Diaz 2003; Sarr et al. 2013). They attributed this to the abundance of food available for the herbivores (Passos et al. 2003; Sarr et al. 2013). This applied particularly to the Elmidae and Hydraenidae. Figure 2. Berosus luridus adult on moss where air bubbles from photosynthesis can be used to replenish the air supply. Photo by Tim Faasen, with permission.

Figure 5. Fontinalis antipyretica on rocks of a stream bed. Photo by Betsy St. Pierre, with permission.

Figure 3. Chaetarthria siminulum adult with plastron. When the plastron is full of air, the beetle must cling to vegetation in order to descend into the water column. Photo by Gerard Visser , with permission.

Nearly all aquatic Coleoptera go to land to pupate (Leech & Chandler 1956; Pennak 1978; Erman 1984), then return to the water as adults. Others clamber about on the surface of the plants. Some of these are associated with floating plants, including Ricciocarpos natans (Figure 4) (Scotland 1934). To get below the surface requires muscle action to break the surface tension (Leng 1913).

Among the most common of these bryophyte dwellers are the Elmidae (Figure 6), small beetles only a few mm in length (Percival & Whitehead 1930; Glime 1994). But many studies miss the small Coleoptera that live among the bryophytes, necessitating special collecting techniques for such habitats as submerged roots, wood, and mosses (Zaťovičová et al. 2004). Zaťovičová and coworkers found 13-61% more species when they used qualitative sampling that included these habitats.

Figure 6. Elmidae adult, one of the most common of beetle families among bryophytes. Photo by Stephen Moore, Landcare Research, NZ, with permission. Figure 4. Ricciocarpos natans, a floating liverwort. Photo by Jan-Peter Frahm, with permission.

Whereas mosses in streams and lakes are not especially important for beetles, bogs and fens have greater

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species numbers. Some live in the acidic pools, some burrow into the moss mats, and some run about the surface. The Dytiscidae (Figure 18-Figure 55) are particularly important in the pools. These bog dwellers, although often not adapted to a submerged aquatic habitat, will be included here. Jones (1950) did extensive gut analysis of insects from the River Rheidol and found that none of the Coleoptera had mosses (Fontinalis antipyretica, Figure 5) in their guts, although Plecoptera and Trichoptera did. Rather, these Coleoptera were all carnivores.

Suborder Adephaga

Figure 8. Pterostichus rhaeticus adult, a blanket-bog dweller. Photo by Niels Sloth, with permission.

This suborder is comprised of a group of highly specialized beetles. Carabidae – Ground Beetles The Carabidae forms a large family (>40,000 species) (Ground Beetle 2015), ranging 0.7-66 mm long (Bartlett 2004a). Despite this large number of species, they are mostly either shiny black or metallic and have ridged elytra (Ground Beetle 2015). Their distribution is worldwide, but records from Africa and Asia are scant. Typical homes are under tree bark, under logs, and among rocks or sand by the edge of ponds and rivers. Many expel an especially noxious and painful liquid for their defense. They are predators, often rapidly chasing their prey, usually at night (Bartlett 2004a). These are not aquatic beetles, but they do live in bogs (Boyce 2011). In Dartmoor, UK, Agonum ericeti (Figure 7) prefers mires that have both Sphagnum (Figure 7) hummocks and warm, bare peat. Here they run around on the bog surface and are one of the most "important" species in the bog. They occur only where there are abundant bog mosses.

Figure 9. Pterostichus diligens adult, an inhabitant of mosses and leaves in blanket bogs. Photo by Niels Sloth, with permission.

Figure 10. Acupalpus dubius on leafy liverworts and mosses. Photo ©Roy Anderson , with permission.

Figure 7. Agonum ericeti adult, a mire dweller, on Sphagnum. Photo by Niels Sloth, with permission.

Pterostichus rhaeticus (Figure 8) prefers to live among Sphagnum (Figure 7) of a blanket bog (Boyce 2011). Pterostichus diligens (Figure 9) likewise lives in blanket bogs, but lives in litter as well as among mosses. Acupalpus dubius is sometimes restricted to the moss Drepanocladus aduncus (Kopecky 2001).

Figure 11. Drepanocladus aduncus, home for Acupalpus dubius. Photo by Bob Klips, with permission.

Chapter 11-9: Aquatic Insects: Holometabola – Coleoptera, Suborder Adephaga

Gyrinidae – Whirligig Beetles This family is aptly named for its behavior of skating in whirling patterns on the water surface. The most unusual feature of this family is the eyes. They are divided so that two eyes are above the water and two are below, protecting the beetles from predators above and permitting them to see what is beneath them (Gyrinidae 2015). Their size ranges 3 to 18 mm long (Whirligig Beetles 2014). They eat insects that fall into the water, sensing the vibrations of their struggles by using their antennae. They are worldwide, with a heavy concentration in Europe. But even these insects sometimes use mosses. At least some members of the Gyrinidae (Figure 12-Figure 14) use mosses as hiding places during the day (Leng 1913). And in the Appalachian Mountain, USA, streams, the mosses may provide a refuge for Dineutus (Figure 12-Figure 14) during times of high flow (Glime 1968).

Figure 12. Dineutus discolor (whirligig beetles) on the water surface. Photo by Janice Glime.

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Haliplidae – Crawling Water Beetles The Haliplidae are clumsy swimmers, alternating the motion of their legs (Haliplidae 2014). Hence, they move about mostly by crawling. The adults are convex on the dorsal side and range 1.5-5.0 mm long. The hind legs have large coxal plates and are immobile. The primary function of these legs seems to be that of storing air, supplementing the air stored under the elytra. The larvae eat only algae, but the adults are omnivorous. They live among aquatic vegetation around the borders of small ponds, lakes, and quiet streams. Their worldwide distribution is similar to that of the Scirtidae, with the greatest diversity known in Europe (Haliplidae 2015). These are mostly not bryophyte dwellers, but the genus Haliplus (Figure 15) still benefits from the presence of Sphagnum (Figure 7). Haliplus variegatus (Figure 16) in Poland lives in canals that are created by beavers in floating Sphagnum mats (Buczyński et al. 2014).

Figure 15. Haliplus larva. Some members of this genus live in bogs and H. variegatus lives in beaver canals in floating Sphagnum mats. Photo by Dana R. Denson, Florida Association of Benthologists, with permission.

Figure 13. Dineutus assimilis adult showing split eyes. Photo by Joyce Gross, with permission.

Figure 14. Dineutus larva, a genus that sometimes occurs among bryophytes when it is resting. Photo by Bob Henricks, with permission.

Figure 16. Haliplus variegatus adults, inhabitants of beaver canals in floating Sphagnum mats of Poland. These color phases and the spots can help to camouflage the beetles among the mosses. Photo by Stefan Schmidt, through Creative Commons.

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In my own studies (Glime 1968) in the Appalachian Mountain, USA, streams, I found the genus Brychius (Figure 17). The generic name suggests a possible moss habitat, but I was unable to find additional information on the habitat.

are passive predators, waiting quietly until a prey organism passes nearby (Dytiscidae 2014). On the other hand, several members of the family are eaten by humans in China, Japan, and Mexico, as well as other places in the world. This worldwide family has a large range of sizes (1.2-40 mm long) (Bartlett 2004b). They are distributed throughout the world, but with the best known concentrations in North America, Europe, and Australia (Dytiscidae 2015). The larvae live in the water, but they climb to land and bury themselves in the mud for pupation, returning to the water as adults. The adult Dytiscidae, like other beetles, lack true gills. Instead, they carry a bubble of air with them as they descend down the water column. This bubble is either held against the body or stored under the elytra (outer hardened wings) (Figure 1). As oxygen is used up, nitrogen maintains the size of the bubble so that oxygen can diffuse into the bubble. When the bubble becomes too small, they must obtain another bubble from plant surfaces or the water surface by exposing the tip of the abdomen (Figure 19).

Figure 17. Brychius elevatus adult, a genus with moss inhabitants in Appalachian Mountain, USA, streams. Photo by Udo Schmidt, through Creative Commons.

Hygrobiidae – Squeak Beetles This small family has only one genus, Hygrobia, with six species, and is distributed in Europe, North Africa, China, and Australia (Hygrobia 2014). Hygrobia adults make a grating noise, earning them their name of squeak beetles (Pendleton & Pendleton 2014). Their size is moderate (8.5-10 mm). They are most common in stagnant water, where they walk or swim; they do not dive (Watson & Dallwitz 2003a). They obtain their oxygen from the air collected and stored under the elytra. Hygrobia hermanni (Figure 18) reaches large populations at pond margins where it lives among the submerged Sphagnum (Figure 39) (Denton 2013).

Figure 18. Hygrobia hermanni adult, an inhabitant of submerged Sphagnum. Note the anal air bubble. Photo by Trevor and Dilys Pendleton, with permission.

Dytiscidae – Predaceous Diving Beetles and Noteridae – Burrowing Water Beetles The Noteridae are often included with the Dytiscidae and I will do so here because it makes the discussion easier. The larvae of Dytiscidae are known as water tigers. They

Figure 19. Rhantus suturellus adult replenishing air supply at surface. Photo by Niels Sloth, with permission.

Based in my own studies on moss-dwelling aquatic insects in the Appalachian Mountains, USA, it seemed that the predaceous diving beetles (Dytiscidae) do not typically hang out among the bryophytes. But many of the species occur in mossy wet areas, especially associated with bogs and fens. Usinger (1974) describes three types of ovipositors in the Dytiscidae. Those with a long ovipositor are able to inject their eggs into moss mats growing in the water. And some species even ingest mosses occasionally (Jones 1949). Roger Key (pers. com. 31 October 2014) considers the primary role of bryophytes in the life of the predaceous aquatic beetles to be that of a structural component, a place for cover to escape predators. But these beetles are mostly predators themselves (Figure 20). In some cases the mosses are important as a place to hang or climb to avoid being carried to the surface by their air supply – the plastron apparatus or air layer under the elytra. For example, Lancetes in South Georgia may make use of mosses, among other anchored substrata, to get back under the surface or to stay there when it is not actively swimming. In places like South Georgia, mosses are the predominant, if not the only, vegetation at the margins of streams, hence providing these roles for aquatic beetles there.

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bogs (Figure 26) in Poland. In spring-fed boggy areas one can find Hydroporus longulus (Figure 27) among mosses and leaves (Denton 2013).

Figure 22. Oreodytes davisii adult, a bryophyte dweller in UK streams. Photo by Udo Schmidt, with permission. Figure 20. Dytiscus larva eating young fish. Photo by Roger S. Key, with permission.

Graphoderus zonatus (spangled diving beetle; Figure 21) occurs where Fontinalis (Figure 5) provides the major vegetation in a heathland mire in Hampshire, UK (Roger S. Key, pers. comm. 31 October 2014). This diving beetle is frequently found associated with the mosses and can be collected by shaking the mosses over a container. The bryophyte role, as suggested above, is one of cover. Oreodytes davisii (Figure 22) and O. sanmarkii (Figure 23) both live among aquatic bryophytes in a stream in Yorkshire, UK (Gilbert et al. 2005). Oreodytes rivalis may occasionally even ingest mosses such as Fontinalis antipyretica (Figure 5) (Jones 1949), perhaps in their attempts to capture one of the other invertebrates dwelling there.

Figure 23. Oreodytes sanmarkii adult, a stream bryophyte dweller in the UK. Photo by Christoph Benisch , with permission.

Figure 21. Graphoderus zonatus adult in a heathland mire in Hampshire, UK. Photo by Roger S. Key, with permission.

Foster (1992) found Hydroporus umbrosus (Figure 24) among mosses at the edge of a pond in Inner Hordaland, Norway. Usinger (1974) describes the small members of the genus Hydroporus as able to occupy mosscovered seepages no bigger than a hand. Buczyński et al. (2014) reported H. incognitus (Figure 25) from Sphagnum

Figure 24. Hydroporus umbrosus adult, a moss dweller at the edge of ponds in Norway. Photo by Niels Sloth, with permission.

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Graphoderus zonatus (Figure 28) in North Hampshire, UK, lives in a variety of habitats, particularly in Sphagnum-dominated (Figure 39) lake margins (Denton 2013).

Figure 28. Graphoderus zonatus adult with Sphagnum. Photo by Niels Sloth, with permission. Figure 25. Hydroporus incognitus adult, an inhabitant of Sphagnum bogs in Poland. Photo by Niels Sloth, with permission.

Figure 26. Sphagnum blanket bog, home to many kinds of beetles. Photo through Creative Commons.

Figure 27. Hydroporus longulus adult, a beetle one can find among mosses in spring-fed boggy areas. Photo by Tim Faasen, with permission.

Moors, Bogs, and Fens These three habitats are partially aquatic, providing wet or damp bryophytes and pools where there may be submerged bryophytes. Moors, a term used more commonly in Europe, are upland habitats including heathlands and fens and characterized by low vegetation and acidic soils (Moorland 2014). The term bog has a mixed history, with North Americans using a much broader definition than that of the northern Europeans. Until relatively recently, North Americans tended to include any wetland with Sphagnum as a bog. English language dictionaries go even further to define a bog as any muddy or spongy wetland. The more restrictive European definition is a habitat that is dominated by Sphagnum and receives only precipitation as a source of new nutrients. By contrast, a fen may have Sphagnum or other dominant bryophytes, but it receives nutrients through surface or ground water in addition to precipitation. Most of the habitats that North Americans have called bogs (including most current definitions and websites on the internet) are actually poor fens, i.e., wetland habitats with low nutrients, ground or surface water, and Sphagnum species similar to those of true bogs. Fens and bogs provide habitats for a number of Dytiscidae and provide the most common associations with bryophytes. The genus Agabus is among these common inhabitants (Nelson 1996). Agabus affinis (Figure 29) can be considered a characteristic species, a tyrphobiont (species living only in peat-bogs and mires) in high moors (Hebauer 1974), often accompanied by A. unguicularis (Figure 30), in the moss lawns of lowland fens and bogs of Ireland (Nelson 1996) and flooded Sphagnum (Figure 39) (Denton 2013). In Scotland A. unguicularis occurs in peaty water with mosses or other dense vegetation (Knight 2014). Agabus melanocornis is less common and occurs in mossy drains, fens, and bogs (Nelson 1996). Agabus melanarius (Figure 31) is easily overlooked in North Hampshire, UK, where it lives in shallow water with mosses.

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(2011) also reported Hydroporus tristis in small, peaty pools that had Sphagnum (Figure 39). Boyce also found Hydroporus gyllenhalii (Figure 40) among Sphagnum in bogs and in small peat pools that likewise had at least some Sphagnum in both undisturbed and eroded blanket mires. Hydroporus obscurus (Figure 42-Figure 43) was more restricted, living only in relatively pristine blanket bogs where it lived in small Sphagnum-dominated peat pools.

Figure 29. Agabus affinis adult with Sphagnum. Photo by Tim Faasen, with permission.

Figure 32. Dytiscus lapponicus larva, a species associated with Sphagnum cuspidatum. Photo by James K. Lindsey, with permission.

Figure 30. Agabus unguicularis adult, a common inhabitant of bogs and fens, carrying an anal air bubble. Photo by Niels Sloth, with permission.

Figure 31. Agabus melanarius adult, a species from shallow water among mosses. Photo by James K. Lindsey, with permission.

In contrast to other bryophyte habitats, bogs are a mix of terrestrial and aquatic microhabitats that provide homes for a number of Dytiscidae. Brink and Terlutter (1983) found Dytiscus lapponicus (Figure 32-Figure 34), Hydroporus tristis (Figure 35), H. erythrocephalus (Figure 36), and Acilius canaliculatus (Figure 37), as well as Noteridae (burrowing water beetles, sometimes included in the Dytiscidae) – Noterus crassicornis (Figure 38), to be acid tyrphophiles (characteristic of bogs but not confined to them) associated with Sphagnum cuspidatum (Figure 39). Acilius is one of the genera with a long ovipositor that permits egg-laying among mosses and other substrata (Usinger 1956). These eggs are laid in the water and sometimes out of water. From Dartmoor, UK, Boyce

Figure 33. Dytiscus lapponicus adult with mosses and aquatic plants. Photo by Niels Sloth, with permission.

Figure 34. Dytiscus lapponicus adult with mosses and aquatic plants. Photo by Niels Sloth, with permission.

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Figure 35. Hydroporus tristis adult amid aquatic mosses. Photo by Tim Faasen, with permission.

Figure 39. Sphagnum cuspidatum, home for some Dytiscidae and Noteridae. Photo by Jan-Peter Frahm, with permission.

Figure 36. Hydroporus erythrocephalus adult with leaf and Sphagnum. Photo by Tim Faasen, with permission. Figure 40. Hydroporus gyllenhalii adult, a species that lives among Sphagnum in bogs and bog pools. Photo by Niels Sloth, with permission.

In his studies in Central Europe, Hebauer (1974) similarly found Hydroporus pubescens (Figure 41) to be a tyrphobiont, as well as such tyrphobionts as Hydroporus obscurus (Figure 42-Figure 43) and H. melanocephalus in the high moors (Hebauer 1994). The smallest member of Irish Hydroporus is H. scalesianus (Figure 44) (Nelson 1996). In the Appalachian Mountain, USA, streams, this genus lives among stream mosses (Glime 1968), whereas in Ireland it lives exclusively among mossy carpets of undisturbed fens, mires, and lake basins. Figure 37. Acilius canaliculatus adult, a species associated with Sphagnum cuspidatum (Figure 39). Photo by Niels Sloth, with permission.

Figure 38. Noterus crassicornis adult on leaf litter in stream. Photo by Niels Sloth, with permission.

Figure 41. Hydroporus pubescens adult among Sphagnum. Photo by Tim Faasen, with permission.

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October 2014). But Ilybius is not restricted to bogs and moors, appearing among mosses in Appalachian Mountain, USA, streams (Glime 1968).

Figure 42. Hydroporus obscurus adult on Sphagnum. Photo by Tim Faasen, with permission.

Figure 45. Rhantus larva. Photo by Dana R. Denson, Florida Association of Benthologists, with permission.

Figure 43. Hydroporus obscurus adult climbing on a moss. Photo by Niels Sloth, with permission.

Figure 44. Hydroporus scalesianus adult, the smallest Hydroporus, on Sphagnum, from the high moors of Europe. Photo by Tim Faasen, with permission.

Other tyrphobionts in the high moors included Rhantus suturellus (Figure 19, Figure 45) (Hebauer 1974), a species also found in Poland in peaty pools (Boyce 2011). In Ireland, Graptodytes granularis (Figure 46) lives in mossy carpets of undisturbed fens, mires, and lake basins, but requires permanently wet mosses (Nelson 1996). Ilybius crassus and I. aenescens (Figure 47-Figure 48) are tyrphobionts in European high moors (Hebauer 2994). Ilybius aenescens also occurs in flooded Sphagnum (Figure 39) of heathlands of North Hampshire, UK, but it is rare (Denton 2013). Boyce (2011) found that Ilybius montanus usually occur in shallow bog pools where there are dense growths of Sphagnum. Ilybius fuliginosus (Figure 49) is quite ubiquitous and thus might be found hiding among the mosses (Tim Faasen, pers. comm. 20

Figure 46. Graptodytes granularis adult, dwelling in the high moors of Europe. Photo by Tim Faasen, with permission.

Figure 47. Ilybius aenescens adult among mosses. Photo by Tim Faasen, with permission.

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Figure 48. Ilybius aenescens adult, a bog dweller. Photo by Niels Sloth, with permission.

Figure 51. Hydaticus seminiger adult, a mossy fen dweller. Photo by Tim Faasen, with permission.

Floating moss carpets are often associated with bogs and fens. Bidessus grossepunctatus (Figure 52) is one of the inhabitants of these moss carpets in small lakes, ponds, fen pools, and mires (Nilsson & Holmen 1995).

Figure 49. Ilybius fuliginosus adult, a ubiquitous species that hides among vegetation, shown here on mosses. Photo by Tim Faasen, with permission.

Laccornis oblongus (Figure 50) is a flightless beetle that lives in Irish fens that lack open water (Nelson 1996). It occurs among wet moss carpets, especially those associated with clumps of sedges. Hydaticus seminger (Figure 51) is a dweller of typical mossy fens. This species is not frequent in North Hampshire, UK, but it does occur among flooded Sphagnum and in detritus pools (Denton 2013).

Figure 50. Laccornis oblongus adult, a flightless beetle known from moss carpets in Irish fens. Photo by Niels Sloth, with permission.

Figure 52. Bidessus grossepunctatus adult, an inhabitant of floating moss carpets, on Sphagnum. Photo by Tim Faasen, with permission.

Special techniques can facilitate collecting bog and fen species. Since bryophytes in these habitats are typically underlain by water, these semi-terrestrial beetles can be collected by depressing the mosses, creating a depression until they are covered by water (Nilsson & Holmen 1995; Knight 2014). The beetles can then be swept from the water with a tea strainer. Knight (2014) considers this technique especially useful for sampling Hydraenidae and small Hydrophilidae. In the Japanese rice fields, many invertebrates find refuge. Some of these fields even have peat mosses. Such communities include Cybister japonicus (Figure 53-Figure 54) (Ohba 2009), a species eaten by humans in Japan (Dytiscidae 2014). These carnivores feed on insects such as Odonata in early instars, but starting in the third instar they feed on small vertebrates such as amphibia as well. In the last larval stage, they burrow into the peat moss and enter the pupation period.

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Figure 56. Drepanocladus aduncus, home of Liodessus cantralli in North America. Photo from Dale A. Zimmerman Herbarium, Western New Mexico University. Figure 53. Cybister japonicus adult, a species that hides among peat mosses in Japanese rice fields. Photo through Creative Commons.

Summary

Figure 54. Cybister japonicus larva, a species that hides among peat mosses in Japanese rice fields. Photo through Creative Commons.

Liodessus cantralli (Figure 55) lives in small pools in North America, but also lives in moss mats of fens (less often in bogs) (Larson & Roughley 1990). They are particularly associated with Drepanocladus s.l. (Figure 56) in depressions in the moss mats.

Coleoptera can live in the water as larvae and as adults, but the pupae are generally on land. The aquatic adults gain oxygen by using a plastron, accumulating air under the forewings, or from an anal bubble. Some live on the surface and may crawl over plants such as Ricciocarpos natans. Smaller beetles live among mosses in streams. But the greatest number of aquatic bryophyte associations for beetles occurs in bogs and fens. The order Coleoptera (beetles) has two sub orders: Adephaga and Polyphaga. In the Adephaga the families Carabidae, Gyrinidae, Haliplidae, Hygrobiidae, and Dytiscidae. The Dytiscidae are especially common and diverse in bog pools and this is the only family of Adephaga frequently associated with bryophytes.

Acknowledgments Tim Faasen not only gave me permission to use his wide collection of insect images, he also helped me to understand the ecology of some of the species and provided me with additional images I needed. Thank you to Roger S. Key not only for his permission to use his images but for sharing his experiences with me regarding beetle use of bryophytes. Ronald Willson verified my beetle identifications for the mid-Appalachian Mountain study. And thank you to Amy Marcarelli, Wayne Minshall, and especially Rich Merritt for helping me with my query about the anal air bubble in aquatic beetles. Eileen Dumire proofread the chapter and suggested changes to provide more clarity.

Literature Cited

Figure 55. Liodessus adult; L. cantralli lives in moss mats of fens. Photo © Stephen Luk through BugGuide noncommercial use, with permission.

Badcock, R. M. 1949. Studies in stream life in tributaries of the Welsh Dee. J. Anim. Ecol. 18: 193-208. Bartlett, Troy. 2004a. Family Carabidae – Ground Beetles. BugGuide. Accessed 15 January 2015 at .

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Bartlett, Troy. 2004b. Family Dytiscidae – Predaceous Diving Beetles. BugGuide. Accessed 15 January 2015 at . Boyce, David. 2011. Invertebrate survey of blanket bog on Dartmoor, 2010. Report accessed 20 October 2014 at . Brink, M. and Terlutter, H. 1983. Beitrag zur Habitatbindung der aquatilen Coleopterenfauna. Abhandlungen aus dem Westfälischen Museum für Naturkunde 45: 50-61. Buczyński, P., Przewoźny, M., Pakulnicka, J., Buczyński, E., Dawidowicz, Ł, and Wagner, G. 2014. Materials to the knowledge of beetles (Coleoptera) of aquatic habitats in the Suwalski Landscape Park. Ann. Univ. Maieae CurieSkłodowska Lublin – Polonia Sec. 2 69: 7-27. Denton, J. 2013. The Water Beetles of North Hampshire. (VC12). Albion Ecology, Four Marks. Accessed 20 October 2014 at . Dytiscidae. 2014. Wikipedia. Accessed 15 January 2015 at . Dytiscidae. 2015. Encyclopedia of Life. Accessed 15 January 2015 at . Erman, N. A. 1984. The use of riparian systems by aquatic insects. In: Warner, R. E. and Hendrix, K. (eds.). California Riparian Systems: Ecology, Conservation, and Productive Management, pp. 177-182. Fernández-Diaz, M. 2003. Estudio faunistico y ecoógico de los coleópteros acuáticos (Adelphaga y Polyphaga) en la cuenca del rio Avia (Ourense, NO España): Distribución espacial y temporal. Tesis de Licenciatura, Universidad de Vigo, 146 pp. Foster, G. N. 1992. Some aquatic Coleoptera from Inner Hordaland, Norway. Fauna Norw. Ser. B 39: 63-67. Gilbert, O., Goldie, H., Hodgson, D., Marker, M., Pentecost, A., Proctor, M., and Richardson, D. 2005. The ecology of Cowside Beck, a tributary of the River Skirfare in the Malham area of Yorkshire. Field Studies Council, Settle, North Yorkshire, UK. Glime, J. M. 1968. Aquatic Insect Communities Among Appalachian Stream Bryophytes. Ph.D. Dissertation, Michigan State University, East Lansing, MI, 180 pp. Glime, J. M. 1994. Bryophytes as homes for stream insects. Hikobia 11: 483-497. Ground Beetle. 2015. Wikipedia. Accessed 15 January 2015 at . Gyrinidae. 2015. Encyclopedia of Life. Accessed 15 January 2015 at . Haliplidae. 2014. Wikipedia. Accessed 15 January 2015 at . Haliplidae. 2015. Encyclopedia of Life. Accessed 15 January 2015 at . Hebauer, F. von. 1974. Über die Ökologische Nomenklatur wasserbewohnender Käferarten. Nachr. Bl. bayer. Entomol. 23(5): 87-92. Hebauer, F. 1994. Entwurf einer Entomosoziologie aquatischer Coleoptera in Mitteleuropa (Insecta, Coleoptera, Hydradephaga, Hydrophiloidea, Dryopoidea). Lauterbornia 19: 43-57. Hygrobia. 2014. Wikipedia. Accessed 15 January 2015 at .

Jones, J. R. E. 1949. A further ecological study of calcareous streams in the 'Black Mountain' district of South Wales. J. Anim. Ecol. 18: 142-159. Jones, J. R. E. 1950. A further ecological study of the river Rheidol: The food of the common insects of the mainstream. J. Anim. Ecol.19: 159-174. Knight, L. R. F. D. 2014. CSM Monitoring of Designated Aquatic Invertebrate Features at Woodhall Loch, Buckstruther Moss, Firth of Forth, Lochs of Harray & Stenness and Rannoch Moor SSSIs. Scottish Natural Heritage Commissioned Report No. 677, 72 pp. Kopecky, T. 2001. Zajímavy vztah mezi strevlíckem a mechem. Ziva, casopis pro biologickou práci 2/2001 str. 82. Larson, D. J. and Roughley, R. E. 1990. A review of the species of Liodessus Guignot of North America north of Mexico with the description of a new species (Coleoptera: Dytiscidae). J. N. Y. Entomol. Soc. 98: 233-245. Leech, H. B. and Chandler, H. P. 1963. Aquatic Coleoptera. In: Usinger, R. L. (ed.). Aquatic Insects of California. University of California Press, Berkeley, Calif., pp. 293-371. Leng, C. W. 1913. Aquatic Coleoptera. J. N. Y. Entomol. Soc. 21: 32-42. Moorland. 2014. Wikipedia. Accessed 22 February 2015 at . Nelson, B. 1996 Species Inventory for Northern Ireland: Aquatic Coleoptera. Ulster Museum, Belfast, 36 pp. Nilsson, A. N. and Holmen, M. 1995. The aquatic Adephaga (Coleoptera) of the Fennoscandia and Denmark. II. Dytiscidae. E. J. Brill, Leiden. Ogbogu, S. S. 2000. Submerged beds of Fontinalis sp. (Bryophyta) as a microhabitat for caddisfly larvae (Trichoptera) in an intermittent reservoir spillway, Ile-Ife, Nigeria. Trop. Freshwat. Biol. 9: 11-16. Ohba, S. Y. 2009. Ontogenetic dietary shift in the larvae of Cybister japonicus (Coleoptera: Dytiscidae) in Japanese rice fields. Environ. Entomol. 38: 856-860. Oliveira de Sousa, W., Rosado-Neto, G. H., and Marques, M. I. 2012. Functionality of the plastron in adults of Neochetina eichhorniae Warner (Coleoptera, Curculionidae): Aspects of the integument coating and submersion laboratory experiments. Rev. Brasil. Entomol. 56: 347-353. Passos, M. I. S., Nessimian, J. L., and Dorville, L. F. M. 2003. Life strategies in an Elmidae (Insecta: Coleoptera: Elmidae) community from first order stream in the Atlantic Forest, southeastern Brazil. Acta Limnol. Brasil. 15(2): 29-36. Pendleton, Trevor and Pendleton, Dilys. 2014. Hygrobia hermanni (Fabricius, 1775). Accessed 15 January 2015 at . Pennak, R. W. 1978. Freshwater Invertebrates of the United States. Second Edition. John Wiley & Sons, New York, 803 pp. Percival, E. and Whitehead, H. 1930. Biological survey of the river Wharf. II. Report on the invertebrate fauna. J. Ecol. 18: 286-295. Sarr, A. B., Benetti, C. J., Fernández-Díaz, M., and Garrido, J. 2013. The microhabitat preferences of water beetles in four rivers in Ourense Province, Northwest Spain. Limnetica 31: 1-10. Scotland, M. B. 1934. The animals of the Lemna association Ecology 15: 290-294. Usinger, R. L. 1956. Aquatic Insects of California: With Keys to North American genera and Species. University of California Press, Berkeley, CA.

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Usinger, R. L. 1974. Aquatic Insects of California: With Keys to North American genera and Species. University of California Press, Berkeley, CA. Watson, L. and Dallwitz, M. J. 2003a onwards. British Insects: The families of Coleoptera. Last updated 25 July 2012. Accessed 15 January 2015 at . Whirligig Beetles. 2014. Wikipedia. Accessed 15 January 2015 at .

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White, D. A. 1967. Trophic dynamics of a wild brook trout stream. Unpublished Ph.D. thesis. Univ. Wisconsin, Madison. Zaťovičová, Z., Čiampor, F. Jr., and Kodada, J. 2004. Aquatic Coleoptera (Insecta) of streams in the Nízke Beskydy Region (Slovakia): Faunistics, ecology and comparison of sampling methods. Biologia, Bratislava 59(15): 181-189.

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Glime, J. M. 2017. Aquatic Insects: Holometabola – Coleoptera, Suborder Polyphaga. Chapt. 11-10. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. Ebook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 19 July 2020 and available at .

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CHAPTER 11-10 AQUATIC INSECTS: HOLOMETABOLA – COLEOPTERA, SUBORDER POLYPHAGA TABLE OF CONTENTS Suborder Polyphaga ........................................................................................................................................ 11-10-2 Helophoridae ............................................................................................................................................ 11-10-2 Hydrochidae ............................................................................................................................................. 11-10-3 Hydrophilidae – Water Scavenger Beetles............................................................................................... 11-10-3 Hydraenidae – Minute Moss Beetles ....................................................................................................... 11-10-9 Ptiliidae – Featherwing Beetles .............................................................................................................. 11-10-11 Silphidae – Large Carion Beetles ........................................................................................................... 11-10-12 Staphylinidae – Rove Beetles................................................................................................................. 11-10-13 Scirtidae (=Helodidae) – Marsh Beetles ................................................................................................ 11-10-15 Elmidae – Riffle Beetles ........................................................................................................................ 11-10-16 Dryopidae – Long-toed Water Beetles ................................................................................................... 11-10-24 Chelonariidae – Turtle Beetles ............................................................................................................... 11-10-25 Lampyridae – Lightning Bugs ............................................................................................................... 11-10-25 Latridiidae – Minute Brown Scavenger Beetles..................................................................................... 11-10-25 Curculionidae – Weevils ........................................................................................................................ 11-10-25 Lagriidae ................................................................................................................................................ 11-10-26 Summary ....................................................................................................................................................... 11-10-26 Acknowledgments ......................................................................................................................................... 11-10-26 Literature Cited ............................................................................................................................................. 11-10-26

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CHAPTER 11-10 AQUATIC INSECTS: HOLOMETABOLA – COLEOPTERA, SUBORDER POLYPHAGA

Figure 1. Ilybius erichsoni adult on Sphagnum. Photo by Niels Sloth, with permission.

Suborder Polyphaga This suborder includes more than 90% of the Coleoptera species. As its name suggests, it eats a tremendous variety of foods.

Helophorus strigifrons (Figure 4) lives in bogs in North Hampshire, UK, among moss and litter (Denton 2013).

Helophoridae This is a family of North America and Europe and has only one genus, Helophorus (Helophoridae 2014). They are relatively small (2-9 mm) (Helophoridae 2014) and live primarily in wetlands (Helophoridae 2015). Most adults live in shallow standing water where they are saprophagous (Fikáček 2009) (organism that feeds on decaying organic matter). Larvae, on the other hand, live in terrestrial, but moist, habitats near water and are predators on small invertebrates. Helophorus grandis (Figure 2) occurs among the aquatic mosses in a stream in Yorkshire, UK (Gilbert et al. 2005). In Canada, Helophorus orientalis (Figure 3) occurs in wet mosses beside small streams (Majka 2008).

Figure 2. Helophorus grandis, an inhabitant of stream mosses in the UK. Photo by Tim Faasen, with permission.

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Figure 5. Hydrochus ignicollis adult, a rare inhabitant of mossy calcareous fens in Ireland. Photo by Tim Faasen, with permission.

Figure 3. Helophorus orientalis adult, a species that lives among wet mosses along streams in Ontario, Canada. Photo by Tom Murray, through Creative Commons.

Figure 6. Marl lake in Jasper National Park, Canada. Photo by Janice Glime.

Figure 4. Helophorus strigifrons adult, a bog dweller in North Hampshire, UK, among moss and litter. Photo by Zoologische Staatssammlung Muenchen, through Creative Commons.

Hydrochidae Although this family is worldwide, it has only one genus, and most of the records are from Europe (Hydrochidae 2015a). Adults and larvae live in both quiet and flowing water where they are herbivores – shredders (Hydrochidae 2015b). The adults range 4-60 mm long. Some of these are associated with bryophytes. Hydrochus ignicollis (Figure 5), a very rare species in Ireland, appeared in collections only twice between 1988 and 1996 (Nelson 1996). Both finds were from mossy calcareous fens adjacent to marl lakes (calcium carbonate or lime-rich lakes. These are alkaline lakes with unconsolidated calcium carbonate or lime-rich mud or mudstone which contains variable amounts of clays and silt (Figure 6-Figure 7).

Figure 7. Marl at margin of marl lake in Jasper National Park, Canada. Photo by Janice Glime.

Hydrophilidae – Water Scavenger Beetles This is a worldwide, mostly aquatic family, typically in open water (Cotinus 2005). The larvae often emerge from the water to pupate, usually hanging from moss at the edge of the water (Water Beetles 2014). The final larval skin is found beneath the pupa. The adults (1-40 mm) are mostly scavengers, but some are predators; larvae are often predators (Cotinus 2005).

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Some Hydrophilidae join the Dytiscidae as common beetles swimming in bog waters. Enochrus (Figure 8Figure 9) is a common genus there (Denton 2013). Enochrus affinis (Figure 10) is often abundant in Sphagnum-dominated (Figure 51) areas of acidic heathland pools (Figure 11) of North Hampshire, UK Enochrus coarctatus (Figure 12) is a mire dweller, preferring older detritus pools but also living in Sphagnum-filled large bog pools. Enochrus ochropterus (Figure 13) does not occur in areas of pure Sphagnum where the Enochrus is exclusively E. affinis. However, it does occur in richer areas with E. coarctatus. The importance of the Sphagnum in its habitats may be due to its role in acidification. Enochrus fuscipennis (Figure 14) lives in the Sphagnum-choked shallow pools of undisturbed blanket bogs in Dartmoor, UK (Boyce 2011). Enochrus hamiltoni (Figure 15), on the other hand, lives in wet mosses next to small streams on Prince Edward Island, Canada (Majka 2008). In the Appalachian Mountain streams, eastern USA, the genus Enochrus can occasionally be found among mosses, as well as the genus Tropisternus (Figure 16-Figure 17) (Glime 1968).

Figure 10. Enochrus affinis adult, an abundant species in Sphagnum-dominated heathland pools in North Hampshire, UK. Photo by Christoph Benisch , with permission.

Figure 8. Enochrus larva, common among bog bryophytes. Photo by Dana R. Denson, Florida Association of Benthologists, with permission. Figure 11. Heathland with a pool. Photo by Jim Champion, through Creative Commons.

Figure 9. Enochrus larval head. Photo by Dana R. Denson, Florida Association of Benthologists, with permission.

Figure 12. Enochrus coarctatus adult, an inhabitant of mire pools, often among Sphagnum. Photo by Udo Schmidt, with permission.

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Figure 13. Enochrus ochropterus adult, a species of rich mires, often associated with Sphagnum. Photo by Niels Sloth, with permission.

Figure 17. Tropisternus natator adult, an occasional moss inhabitant in Appalachian Mountain, USA, streams. Photo by Donald S. Chandler, with permission.

Figure 14. Enochrus fuscipennis adult, a species that lives in Sphagnum-filled shallow pools in blanket bogs. Photo by James K. Lindsey, with permission.

The genus Laccobius (Figure 18-Figure 19) associates with mosses in both stream and mire habitats. Laccobius reflexipennis (see Figure 18) live in wet mosses next to small streams on Prince Edward Island, Canada (Majka 2008). Laccobius atratus in Ireland and Great Britain occurs in Sphagnum (Figure 51) bogs and other peatlands (Friday 1987; Nelson 1996; Denton 2013). Laccobius ytenensis adults live among mosses around the tiny pools that occur in the seepage lines of UK bogs (Denton 2013).

Figure 18. Laccobius sp. adult, a genus with several species that live in water or bog mosses. Photo by Gerard Visser , with permission. Figure 15. Enochrus hamiltoni adult, a dweller of wet mosses next to small streams on Prince Edward Island, Canada. Photo by Tom Murray, through Creative Commons.

Figure 16. Tropisternus sp. larva, an occasional moss inhabitant in Appalachian Mountain, USA, streams. Photo by Tom Murray, through Creative Commons.

Figure 19. Laccobius adult with open wings showing the membranous wings under the hardened elytra. Photo by Michael Schmidt, through Creative Commons.

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Chaetarthria siminulum (Figure 20) can be present in "huge" numbers among mosses at the edges of ponds (Denton 2013). It also lives among mosses in fens and in fen litter.

Figure 22. Hydrobius fuscipes adult, a species of Sphagnum bogs. Photo by Tim Faasen, with permission.

Figure 20. Chaetarthria siminulum adult with plastron. When the plastron is full of air, the beetle must cling to vegetation in order to descend into the water column. Photo by Gerard Visser, with permission.

Hebauer (1994) found Crenitis punctatostriata (Figure 21) in the high moors, living as a tyrphobiont. Hydrobius fuscipes (Figure 22-Figure 23) on Prince Edward Island (Majka 2008) occurs in Sphagnum (Figure 51) bogs and other peatlands.

Figure 21. Crenitis punctatostriata adult, a beetle that lives in bogs of the high moors. Photo by Udo Schmidt, with permission.

Figure 23. Hydrobius larval head showing large mandibles. Photo by Dana R. Denson, Florida Association of Benthologists, with permission.

Friends are wonderful, and I recently received this story and all the images from Andrea Ares. She found an "amazing place" covered with the leafy liverwort Jungermannia vulcanicola (Figure 24-Figure 25) in Chatubomigoke Park, Gunma Prefecture, Japan. Soon she also discovered a small (6-7 mm) black beetle wending its way upon and within the "big, robust carpet" of the liverwort in this acid stream. This beetle was identified by Itouga san as Hydrobius pauper (Figure 26-Figure 28), the only member of the genus in Japan. There was not just one, but the bases of the liverworts were "full" of them.

Figure 24. Cushions of Jungermannia vulcanicola (chartreuse-colored cushions) in Chatubomigoke Park in Japan. Photo courtesy of Angela Ares.

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Figure 28. Hydrobius pauper adult. Photo by Itago san.

Berosus luridus (Figure 29, Figure 30) is tyrphophilic, living among Sphagnum (Figure 51), but can also be found in other places (Tim Faasen, pers. comm.). I have found no other records of it living among Sphagnum, but it is rare in the Netherlands and may be rare elsewhere. Perhaps the Sphagnum provides a relict habitat, a safe site where conditions are still tolerable. Figure 25. Habitat of Jungermannia vulcanicola (chartreuse-colored cushions) in Chatubomigoke Park in Japan. Photo courtesy of Angela Ares.

Figure 26. Cushion of Jungermannia vulcanicola with its inhabitants, Hydrobius pauper. Photo courtesy of Angela Ares.

Figure 27. Disturbed cushion of Jungermannia vulcanicola showing bases of plants with its inhabitants, Hydrobius pauper. Photo courtesy of Angela Ares.

Figure 29. Berosus luridus adult on moss, a rare beetle in the Netherlands, but present in bogs among Sphagnum there. Note the air bubbles on the moss; these can be used to replenish the air supply. Photo by Tim Faasen, with permission.

Figure 30. Berosus larva, a moss dweller in bogs of New Zealand. Photo by Stephen Moore, Landcare Research, NZ, with permission.

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In tropical Africa, the genus Anacaena is probably more common than is recognized. Komarek (2004) described nine new species. Among these, four were from mosses. Anacaena capensis occurs among the mosses and leaf litter of mountain rivulets in South Africa. Anacaena glabriventris lives among mosses in small streams; A. reducta likewise lives among mosses in small streams, but with steep channels. Anacaena tenella lives among hygropetric mosses (mosses growing on vertical rock faces where a thin film of water flows) in mountain streams. Anacaena limbata (Figure 31) lives in wet mosses next to small streams on Prince Edward Island, Canada (Majka 2008). Figure 33. Helochares punctatus adult on moss. Photo by Niels Sloth, with permission.

Figure 31. Anacaena limbata adult, an inhabitant of wet mosses adjacent to streams. Photo by Tim Faasen, with permission.

Anacaena globulus (Figure 32) lives among Sphagnum (Figure 32) in bogs in Europe and can be collected by squeezing the moss (Buczyński et al. 2014). However, Faasen (personal communication) does not find them typically in Sphagnum bogs in the Netherlands, but considers them widespread, occasionally occurring in bogs. Also in Dartmoor, UK, Helochares punctatus (Figure 33) is an obligate mire species, living among saturated Sphagnum, particularly S. cuspidatum (Figure 34), of pools and acid flushes.

Figure 32. Anacaena globulus adult on Sphagnum, one of its many habitats. Photo by Tim Faasen, with permission.

Figure 34. Sphagnum cuspidatum, home for Helochares punctatus. Photo by David T. Holyoak, with permission.

Nelson (1996) found several additional species of Hydrophilidae in Irish mossy fens. These included Cercyon convexiusculus (Figure 35-Figure 36) in mossy fens. In North Hampshire, UK, Denton (2013) found this species to be abundant in detritus and rotting leaf litter, but also among mosses that bordered richly vegetated sites. Cercyon marinus similarly occupied mosses or decaying organic matter at the water's edge in Ireland (Nelson 1996). Cercyon ustulatus (Figure 37) occurs in mossy areas of ponds and also occurs among mosses growing on sewage filter beds (Denton 2013).

Figure 35. Cercyon convexiusculus adult, an inhabitant of mossy fens. Photo by Tim Faasen, with permission.

Chapter 11-10: Aquatic Insects: Holometabola – Coleoptera, Suborder Polyphaga

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Hydraenidae – Minute Moss Beetles

Figure 36. Cercyon convexiusculus adult, an inhabitant of mossy fens. Photo by Christoph Benisch , with permission.

Adults of Hydraenidae (Figure 39), known as minute moss beetles (1-3 mm length), are aquatic, but the larvae drown if completely submersed (Watson & Dallwitz 2012). Even adults are poor swimmers (EOL 2014); most eat plants, but a few are carnivorous or saprophagous (feeding on decaying organic matter) (Hydraenidae 2014). They are sparsely distributed worldwide with a concentration in Europe (EOL 2014). Sarr et al. (2013) found that Hydraena was correlated with a moss substrate in Northwest Spain. Berthélemy (1966) found this family commonly among mosses in the Pyrénées, including Hydraena gracilis (Figure 40), H. minutissima, and H. pygmaea (Figure 41), with the latter two being considered muscicoles (thriving among mosses). He also considered Hydraena pulchella (Figure 42) and Hadrenya to be muscicoles. Nelson (1996) reported Hydraena gracilis as a common and widespread species in Britain where it lives on mossy rocks in fast-flowing streams and rivers.

Figure 37. Cercyon ustulatus adult, an inhabitant of mossy areas of ponds and filter beds. Photo by Tom Murray, through Creative Commons.

Paracymus scutellaris (Figure 38) occurs among peat mosses in Ireland (Nelson 1996).

Figure 38. Paracymus scutellaris adult, a peat moss dweller in Ireland. Photo by Udo Schmidt, with permission.

Figure 39. Hydraenidae adult, an aquatic minute moss beetle that commonly lives among mosses in the Pyrénéenes. Photo by Stephen Moore, Landcare Research, NZ, with permission.

Figure 40. Hydraena gracilis adult, a common aquatic moss inhabitant in the Pyrénées. Photo by Tim Faasen, with permission.

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Chapter 11-10: Aquatic Insects: Holometabola – Coleoptera, Suborder Polyphaga

Hebauer (1994) found similar species representation from this family in middle Europe. Among the stream mosses he found Hydraena minutissima, H. pygmaea (Figure 41), and H. pulchella (Figure 42). Several more used mosses or algae as a substrate: Ochthebius granulatus (Figure 44), O. metallescens (Figure 45), O. exsculptus (Figure 46), O. melanescens, O. colveranus, and O. halbherri. Eggs of Ochthebius are either naked or somewhat covered by loosely applied silk provided by the mother; the eggs hatch in 7-10 days. In rivers in Northwest Spain, Sarr et al. (2013) found that Ochthebius heydeni was likewise correlated with a moss substrate.

Figure 41. Hydraena pygmaea adult, a muscicole in the Pyrénées. Photo by Tim Faasen, with permission.

Figure 44. Ochthebius granulatus adult, a stream moss dweller in middle Europe. Photo by Magnus Manske.

Figure 42. Hydraena pulchella adult, a tiny beetle that lives among stream mosses in Europe. Image through Creative Commons.

Hydraena nigrita is a tiny beetle that lives among mosses at the edges of streams, but it will climb out if the moss is placed under water (Anderson 2014). It is considered vulnerable because of siltation and loss of habitat (Foster et al. 2009). Hydraena rufipes (Figure 43) lives among mosses (Nelson 1996; Knight 2014) and fine shingle (mass of small rounded pebbles) along rivers (Nelson 1996).

Figure 43. Hydraena rufipes adult, a species that lives among mosses along rivers. Photo from Zoologische Staatssammlung Muenchen, through Creative Commons.

Figure 45. Ochthebius metallescens adult, a beetle that uses mosses and algae as substrates. Photo by Tim Faasen, with permission.

Figure 46. Ochthebius exsculptus adult, a European stream moss dweller. Photo by Udo Schmidt, with permission.

Chapter 11-10: Aquatic Insects: Holometabola – Coleoptera, Suborder Polyphaga

Limnebius nitidus (Figure 47) is among the smallest of the water beetles and in addition to wet mud, it makes mosses in swamps and at the edges of pools and streams its home (Nelson 1996). Adults are a mere mm long, so these scavengers of dead plants and animals are easily overlooked (Hilsenhoff 1975). Eggs of this genus are either naked or somewhat covered with loosely applied silk and hatch in 7-10 days (Usinger 1956). In my studies in the Appalachian Mountain streams of the eastern US, this genus likewise occurred among submerged mosses (Glime 1968).

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Ptiliidae – Featherwing Beetles This is a large, worldwide family of minute (0.3-2 mm long) beetles (Ptiliidae 2015). The egg size is half the length of the body and only one is developed at a time, permitting the female to store a large energy supply in the egg. Their wide-ranging habitats include moist leaf litter, under bark of dead trees, along sand and gravel banks of rivers and streams, beneath seaweed on beaches, in mammal nests, on dung, rotting cacti, ant and termite colonies, and other habitats containing rotting or damp organic material. And some seem to live their entire lives in bogs. The small size of several Ptiliidae beetles – Tychobythinus bythinioides (Staphylinidae or Ptiliidae; Figure 65), Ptiliopycna moerens (Figure 49), Acrotrichis (Figure 50) – and other small beetles in bogs seems to correlate with a high incidence of parthenogenesis (reproduction from an unfertilized egg) in relict (habitat that survived from an earlier period) bogs (Dybas 1978), most likely having poor dispersal as an additional selection factor.

Figure 47. Limnebius nitidus adult, one of the smallest of all water beetles and a moss dweller in swamps. Photo through United States public domain.

Hygrotus decoratus (Figure 48) lives in shallow, mossy fens in North Hampshire, UK, where mosses may provide safe sites for larvae and adults (Denton 2013). Hygrotus novemlineatus was reared with Chironomidae larvae as a food source (Nilsson 1983). Mosses were provided in the culture chamber. After a few days, the beetles laid eggs, attaching them to branches of mosses. But is this a normal substrate for egg-laying in nature? The habitat seems suitable, providing lots of Chironomidae larvae as food. This genus should be sought among bryophytes in other fens.

Figure 48. Hygrotus decoratus adult, a species of shallow mossy fens, at surface getting air. Photo by Niels Sloth, with permission.

Figure 49. Ptiliopycna moerens adult, a parthenogenetic inhabitant of relict bogs. Photo © Stephen Luk for noncommercial use, with permission.

Figure 50. Acrotrichis sp. adult, a parthenogenetic inhabitant of relict bogs. Photo by Joyce Gross, with permission.

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Chapter 11-10: Aquatic Insects: Holometabola – Coleoptera, Suborder Polyphaga

Ptiliopycna moerens is minute, less than 1.0 mm long, and lives in the northeastern United States and adjacent Canada (Dybas 1978). It lives in Sphagnum in bogs and swamp forests, confined within the limits of Wisconsinian glaciation. Males are seemingly restricted to the northern part of the range. More southern locations have parthenogenetic females, a common character of small beetles in relict bogs. (See the chapter on Terrestrial Insects – Coleoptera for further discussion of beetles in bogs.) Silphidae – Large Carrion Beetles This family is predominantly in the Northern Hemisphere, although scattered records exist in the Southern Hemisphere (Silphidae 2015a). Ranging in size from 7-45 mm, the family is rare in the tropics where ants might out-compete them (Silphidae 2015b). As the common name implies, the family feeds on decaying organic matter. Because of this feeding behavior, forensic scientists use their stage of development to determine how long a body has been dead. Despite the need to find new carcasses as their carcass home ages, the Silphidae use walking as their primary means of locomotion (Silphidae 2015b). Most of their activity occurs at night. The Silphidae have a variety of defenses (Silphidae 2015b). These include color warnings from aposematism (use of bright colors to advertise danger or unpalatability) to Batesian mimicry (mimicking coloration or behavior of poisonous or unpalatable species), chemical defenses, and parental care. And many of them use camouflage, having dark colors with a mix of gold, black, and brown to blend with their environment. Some carrion beetles (Silphidae) occur in bogs. Beninger and Peck (1992) described the resource use by Nicrophorus species (carrion beetles, Silphidae) in a Sphagnum (Figure 51) bog near Ottawa, Canada, and found that resource use differed little from resource use in forested habitats. However, Nicrophorus vespilloides (Figure 52) used only small carrion (Figure 53) in the bog for reproduction, whereas the closely related N. defodiens (Figure 54) went to the nearby forest for reproduction. Likewise, N. sayi (Figure 55), N. orbicolis (Figure 56), and N. tomentosus (Figure 57), also bog inhabitants, were rarely associated with the small carrion of the bog, but rather reproduced mostly in the forest.

Figure 51. Sphagnum blanket bog. Photo through Creative Commons.

Figure 52. Nicrophorus vespilloides adult, a common carrion beetle that occurs in bogs. Photo by Tim Faasen, with permission.

Figure 53. Nicrophorus vespilloides with small carrion, a preferred substrate for its reproduction in bogs. Photo by Niels Sloth, with permission.

Figure 54. Nicrophorus defodiens adult, a bog dweller that goes to the forest to reproduce. Photo by Derek Sikes, through Creative Commons.

Chapter 11-10: Aquatic Insects: Holometabola – Coleoptera, Suborder Polyphaga

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Staphylinidae – Rove Beetles

Figure 55. Nicrophorus sayi adult, a bog dweller that goes to the forest to reproduce. Photo by Tom Murray, through Creative Commons.

These beetles are distinctive in having short wings that cover less than half the abdomen (Bartlett 2004). This large family has about 58,000 species, ranging in size from 1 to 35 mm, but mostly 2-8 mm (Rove Beetle 2014). Distribution is worldwide, but records are lacking in vast areas of Asia and Africa. They live in every imaginable type of habitat and likewise eat everything – except living plants! There is now one exception to that – a recent discovery of a herbivore. Like the Carabidae, the Staphylinidae are not aquatic, but likewise inhabit bogs (Boyce 2011). In Dartmoor, UK, Gymnusa brevicollis (Figure 58) is stenotopic (able to tolerate only a restricted range of habitats or ecological conditions). Its preferred habitat is saturated Sphagnum (Figure 51) in extremely wet acid mires where they can be found at the edge of bog pools.

Figure 58. Gymnusa brevicollis adult, a beetle that lives among saturated Sphagnum at the edge of bog pools of wet acid mires. Photo from Zoologische Staatssammlung Muenchen, through Creative Commons. Figure 56. Nicrophorus orbicolis adult, a bog dweller that goes to the forest to reproduce. Photo by Tom Murray, through Creative Commons.

Figure 57. Nicrophorus tomentosus adult, a bog dweller that goes to the forest to reproduce. Photo by Tom Murray, through Creative Commons.

Myllaena kraatzi (Figure 59), a nationally (UK) rare species, is restricted to very high quality acid mires with abundant bog mosses (Boyce 2011). It is collected by shaking the Sphagnum (Figure 51) and litter, suggesting close ties with these two substrates. Oxypoda procerula (Figure 60) is likewise sampled by shaking the litter and Sphagnum, indicating that it is directly a moss dweller.

Figure 59. Myllaena vulpina adult. Myllaena kraatzi is a rare species of high quality acid mires in the UK. Photo by Reginald Webster, Jan Klimaszewski, Georges Pelletier, and Karine Savard through Creative Commons.

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Chapter 11-10: Aquatic Insects: Holometabola – Coleoptera, Suborder Polyphaga

Figure 60. Oxypoda procerula adult, a Sphagnum and litter dweller. Photo by Udo Schmidt, through Creative Commons.

Philonothus nigrita (Figure 61) is a characteristic species in Sphagnum-dominated (Figure 51) acid mires (Boyce 2011). It can be found by treading on the moss cushions, causing it to float out of the saturated Sphagnum. Stenus brevipennis (see Figure 62) lives among Sphagnum in blanket bogs. Stenus kiesenwetteri (Figure 63) is rare in the UK, occurring in very wet Sphagnum (Butler 1886).

Figure 63. Stenus kiesenwetteri adult, a rare beetle inhabiting very wet Sphagnum. Photo by Udo Schmidt, through Creative Commons.

Unlike the other Staphylinidae discussed here, Dianous coerulescens (Figure 64) lives where water trickles over mosses and liverworts (Butler 1886).

Figure 61. Philonothus nigrita adult, a species that characterizes Sphagnum-dominated acid mires. Photo by Marko Mutanen, through Creative Commons.

Figure 64. Dianous coerulescens adult on leafy liverwort. Photo by Malcolm Storey, through Creative Commons.

Figure 62. Stenus biguttatus adult. Stenus brevipennis lives among Sphagnum of blanket bogs. Photo through Creative Commons.

The Pselaphinae beetles are represented along the postglacial fringe in the central and eastern United States where they inhabit Sphagnum (Figure 51) bogs (Reichle 1966). More than 20 species of pselaphids characterize these bogs. They are relict species with specific habitat requirements and poor dispersal ability. Some have very specific temperature range requirements: Tychobythinus bythinioides (=Bythinopsis tychoides; Figure 65), 21.5±0.81, 25.9-15.3°C; Decarthron defectum, 28.5±0.55, 31.4-24.0; Pselaphus ulkei, 19.5±0.86, 24.7-13.0; Reichenbachia borealis (a short-winged mold beetle; Figure 66), 21.±0.99, 26.2-14.4; Rybaxis clavata (Figure 67), 28.3±0.41, 29.9-25.1 (Reichle 1967). The moss microhabitats provide them with both the required nearsaturation humidities and the multiple temperature ranges they require. Changes in temperature stratification regimes result in different species occurring at different seral stages in the bogs.

Chapter 11-10: Aquatic Insects: Holometabola – Coleoptera, Suborder Polyphaga

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Scirtidae (=Helodidae) – Marsh Beetles

Figure 65. Tychobythinus bythinioides adult, a minute beetle that takes advantage of the temperature and moisture stratification in a Sphagnum bed to meet its needs. Photo from Biodiversity Institute of Ontario, through Creative Commons.

This is a worldwide family, but is most diverse in the temperate region (Murray 2005). The larvae live in both stagnant and flowing water where abundant decomposing plant material is present. Adults live on vegetation and on rotting vegetation. The Scirtidae are soft-bodied relative to other beetles and are slightly flattened to nearly subglobular (almost globe-shaped) (TOL 2011). Their sizes range 1-15 mm long. Some females secrete substances that may be pheromones used to stimulate males into courtship (Ruta 2008). This is typically a beetle of open water, but in a subalpine springbrook in the southern Alps of New Zealand, Scirtidae (Figure 68) are most abundant in the moss Acrophyllum quadrifarium (=Pterygophyllum quadrifarium; Figure 69) at the edge of the inner spray zone where the mosses are saturated (Cowie & Winterbourn 1979).

Figure 66. Reichenbachia borealis adult, a minute beetle that takes advantage of the temperature stratification in a Sphagnum bed to meet its temperature needs. Photo by Tom Murray, through Creative Commons.

Figure 68. Helodidae adult, a beetle that is abundant among Acrophyllum quadrifarium in the subalpine springbrooks of the southern Alps of New Zealand. Photo from Pybio at , with permission.

Figure 67. Rybaxis clavata adult, a minute beetle that takes advantage of the temperature stratification in a Sphagnum bed to meet its temperature needs. Photo by Tom Murray, through Creative Commons.

Figure 69. Achrophyllum quadrifarium, a bryophyte habitat for Helodidae in streams in the Southern Alps of Australia. Photo by Jan-Peter Frahm, with permission.

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Chapter 11-10: Aquatic Insects: Holometabola – Coleoptera, Suborder Polyphaga

Cyphon (Scirtidae; Figure 70-Figure 72) has been collected from wet mosses at the edge of a cold spring (Usinger 1974). Cyphon hilaris (Figure 71) in Dartmoor, UK, prefers bog pools that have Sphagnum (Figure 51) (Boyce 2011). In North Hampshire, UK, C. hilaris occurs infrequently in wetlands with peaty soils, acidic bogs, and fens (Denton 2013). Cyphon padi (Figure 72), also in North Hampshire, prefers peaty areas in wooded sites where the Sphagnum is flooded.

Figure 70. Cyphon pupa. Photo by Dana R. Denson, Florida Association of Benthologists, with permission.

Elmidae – Riffle Beetles These are small beetles (1-8 mm) (Gordon & Post 1965). The Elmidae have a distribution similar to that of the Silphidae, but there are more known locations, including southern Africa (Harrison 2009). As the common name describes, these beetles usually live in the riffles of cool, rapid streams (Arnett et al. 2002; Harpootlian 2005). They feed mostly on decaying plants and algae (Epler 2010). Only three species of Elmidae are considered to be frequent aquatic bryophyte dwellers: Promoresia tardella (Figure 73), Atractelmis wawona (Figure 74), and Cleptelmis addenda (Figure 75) (Brown 1972; Shepard & Barr 1991; Bowles et al. 2003; Elliott 2008a), all from North America where the family has many more species (80 species) than in Europe (46 species) (Elliott 2008a). But if one looks among the liverworts in the Pacific states of USA, a fourth genus, Bryelmis (Figure 108-Figure 110) is lurking (Bowles et al. 2003 – see below); further searching among submerged leafy liverworts may expand this Bryelmis distribution. Nevertheless, a number of species use bryophytes at some stage in their lives. Both larvae and adults of some Elmidae are able to feed on mosses (Usinger 1974). When disturbed, Elmidae may play dead for a number of hours before attempting to relocate (Usinger 1956). Cleptelmis (Figure 75) may wait for 12-15 hours before moving. Such patience!

Figure 71. Cyphon hilaris adult, a species that occurs among wet mosses at the edge of a spring Photo by Stefan Schmidt, through Creative Commons.

Figure 73. Promoresia tardella adult, one of the few frequent bryophyte dwellers in the Elmidae. Photo through Creative Commons.

Figure 72. Cyphon padi adult, a species of flooded Sphagnum in peaty forested areas. Photo by Miroslav Deml, through Creative Commons.

Figure 74. Atractelmis wawona adult, a frequent bryophyte inhabitant. Photo through Creative Commons.

Chapter 11-10: Aquatic Insects: Holometabola – Coleoptera, Suborder Polyphaga

Figure 75. Cleptelmis addenda adult, one of the few frequent Elmidae bryophyte dwellers. Photo by Crystal Maier, through Creative Commons.

Elmidae colonize mosses when insect-free mosses are introduced, but some of the elmids may be slow to colonize. This is no surprise since they creep and don't swim. For example, Maurer and Brusven (1983) found that the elmid Cleptelmis ornata (Figure 76) was the only insect that was slow to colonize insect-free test clumps of Fontinalis neomexicana (Figure 77) during a field experiment in Idaho, USA. Elliott (2008a) summed up some of the characters that define the bryophyte dwellers. Their larvae have a triangular cross section. Among this group he included Elmis (Figure 87-Figure 86), Esolus (Figure 84-Figure 85), and Oulimnius (Figure 88-Figure 89), none of which were considered by earlier researchers mentioned above to be the frequent bryophyte dwellers. All members of the family have aquatic larvae and most have aquatic adults. The pupae are terrestrial. This means that the newly emerged adults must re-enter the water – no small feat for such a small insect. They must break through the surface tension – easy for us, but nearly impossible for them unless they have something to cling to and provide leverage for them to break through (see Figure 78). Bryophytes, plants, and rocks can help here.

Figure 76. Cleptelmis ornata adult, a slow colonizer of Fontinalis neomexicana. Photo from BIO Photography Group, Biodiversity Institute of Ontario, through Creative Commons.

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Figure 77. Fontinalis neomexicana, a moss that is avoided as home for liverwort-dwelling Bryelmis. Photo by Belinda Lo, through Creative Commons.

Figure 78. Dryops auriculatus (Dryopidae) adult entering water by clinging to a plant. Note the rings in the water and depression of the surface by the beetle body. Photo by Tim Faasen, with permission.

The aquatic adults use the plastron for oxygen availability (Thorpe & Crisp 1949) – they are air breathers. The plastron apparatus is seen as a silvery layer (Figure 79) on the ventral side of the beetle. Some members include the antennae as part of the apparatus that holds the air bubble. They groom the plastron with brushes on the femur of the leg and also use these brushes to add air bubbles to the plastron apparatus by smearing bubbles over the plastron. Most do not need to return to the surface, using the mouthparts to capture oxygen bubbles emitted by plants. If the plastron air layer is thick, it has a silvery sheen and is called a macroplastron (Figure 116). When air diminishes from the macroplastron to the normal, smaller plastron, air exchange with the water is generally adequate to maintain the duller-looking air bubble and meet their needs. This low need for fresh air is likely possible because these beetles do not swim, requiring less oxygen for their clambering movements. In a tributary of the Danube, Elmis maugetii and Riolus subviolaceus (Figure 80) were abundant in high flow areas among coarse mosses, whereas Esolus parallelepipedus (Figure 81) and Limnius volckmari (Figure 82-Figure 83) were among algae in moderately flowing water (Dietrich & Waringer 1999). Esolus

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Chapter 11-10: Aquatic Insects: Holometabola – Coleoptera, Suborder Polyphaga

angustatus (Figure 84-Figure 85) and Oulimnius tuberculatus (Figure 88-Figure 89) were more common in moderate flow with abundant moss-covered pebbles.

Figure 82. Limnius volckmari larva, an elmid that seems to prefer algae to mosses as a substrate. Photo by Urmas Kruus, with permission.

Figure 79. Riolus subviolaceus adult with thin plastron showing as a silver line where the elytra meets the ventral plastron. Photo by Tim Faasen, with permission. Figure 83. Limnius volckmari adult, an elmid that seems to prefer algae to mosses as a substrate. Photo by Urmas Kruus, with permission.

Figure 80. Riolus subviolaceus adult, inhabitants of high flow areas among coarse mosses. Photo by Tim Faasen, with permission.

Figure 81. Esolus parallelepipedus adult, a species with a high drift rate. Photo from Zoologische Staatssammlung Muenchen, through Creative Commons.

Figure 84. Esolus angustatus larva, member of a genus that has the triangular cross section that characterizes many bryophyte dwellers. Photo by Tim Faasen, with permission.

Figure 85. Esolus angustatus adult, member of a genus that is common among bryophytes. Photo by Tim Faasen, with permission.

Chapter 11-10: Aquatic Insects: Holometabola – Coleoptera, Suborder Polyphaga

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In British streams and rivers, Elmis aenea (Figure 86Figure 87), a moss dweller in rapid streams and rivers (both above and below water), occurred among bryophytes as both adults and larvae, but larvae were more abundant among small stones or under larger ones (Elliott 2008a). In these rivers and streams, Oulimnius tuberculatus (Figure 88-Figure 89) preferred tracheophytes.

Figure 88. Oulimnius tuberculatus adult, a European moss dweller. Photo by J. C. Schou, with permission.

Figure 86. Elmis aenea larva, a species whose distribution is related to elevation. Photo by Niels Sloth, with permission.

Figure 89. Oulimnius tuberculatus larva, an aquatic moss dweller. Photo by J. C. Schou, with permission.

Figure 87. Elmis aenea adult, a moss dweller in rapid streams and rivers. Photo by Tim Faasen, with permission.

In a 39-month study, Elliott (2008b) examined the effect of density on drift rate. Most of the larvae and adults of Elmidae drift at night with very few drifting in daytime. Elliott found that the Elmidae in the study, including the bryophyte dwellers, did not drift on the basis of density. Drift losses accounted for only about 0.07% of total losses in the benthos. The exception to this was the high drift, during a heavy rainfall, of early stages of immature adults of Elmis aenea (Figure 87), Oulimnius tuberculatus (Figure 88-Figure 89), and Esolus parallelepipedus (Figure 81), all species known from bryophytes. For Elmis aenea, the highest drift density was in the earliest life stage soon after egg hatching; for O. tuberculatus it was the start of the larval overwintering period. Frost (1942) found that Oulimnius tuberculatus lives among mosses (and other habitats); moving to land for pupation most likely subjects this insect to the drift.

Nelson (1996) described Elmis aenea (Figure 86Figure 87) as a species from moss-covered rocks in rapid rivers and streams. Berthélemy (1966) found larvae (Figure 86) and adults (Figure 87) of E. aenea and E. maugetii were often abundant among mosses and liverworts in the Pyrénées. The moss-dwelling species were generally smaller than those among stones. Nelson found that the proportion of E. aenea vs E. rioloides (Figure 90) among mosses was related to elevation.

Figure 90. Elmis rioloides adult, a moss dweller whose distribution is affected by elevation. Photo through Creative Commons.

Gurtz and Wallace (1984) found larvae of the elmid Promoresia in only one sample in Big Hurricane Branch.

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Chapter 11-10: Aquatic Insects: Holometabola – Coleoptera, Suborder Polyphaga

They had learned from J. Haefner (personal communication) that these larvae in Sawmill Branch occurred almost exclusively among aquatic mosses (Haefner & Wallace 1981). I found Promoresia elegans (Figure 91-Figure 92) frequently among the bryophytes [Fontinalis dalecarlica (Figure 93-Figure 94), Hygroamblystegium fluviatile (Figure 95), Scapania undulata (Figure 112)] of Appalachian Mountain, USA, streams. This is a genus that exhibits the triangular cross section that Elliott (2008a) suggested to be characteristic of bryophyte dwellers.

Figure 94. Fontinalis dalecarlica showing the dangling streamers. Photo by Jan-Peter Frahm, with permission.

Figure 91. Promoresia elegans adult, a common stream moss inhabitant. Photo through Creative Commons.

Figure 95. Hygroamblystegium fluviatile, home to several species of Elmidae. Photo by Janice Glime.

Figure 92. Promoresia elegans, a larva that is common among bryophytes. Photo by Erin Hayes-Pontius, through Wikimedia Commons.

In addition to Elmis, Berthélemy (1966) found Riolus cupreus (Figure 96-Figure 97), Esolus parallelepipedus (Figure 81), and Oulimnius tuberculatus (Figure 88-Figure 89) among mosses in streams in the Pyrénées. Elmis and Oulimnius were strong muscicoles (living among or in association with mosses). Hebauer (1994) found Elmis obscura, E. rioloides (Figure 90), and Oulimnius tuberculatus among mosses in streams in Central Europe.

Figure 93. Riffles with Fontinalis dalecarlica, home for Promoresia elegans. Photo by Janice Glime.

Figure 96. Riolus cupreus larva, an inhabitant of Pyrénées stream mosses. Photo by Urmas Kruus, with permission.

Chapter 11-10: Aquatic Insects: Holometabola – Coleoptera, Suborder Polyphaga

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Figure 97. Riolus cupreus adult, an inhabitant of Pyrénées stream mosses. Photo by Urmas Kruus, with permission.

The Elmidae spend their larval life in the water, pupate on land, then after their initial dispersal flight they return to the water. The interesting note here is that once they return to the water, they lose their ability to fly (Ward 1992). This locks them into their habitat no matter what the water conditions. For those inhabiting stream mosses, this means that if the water level drops, they must remain in the habitat of the mosses, unable to disperse for any significant distance. But for them it seems to be no problem because they have a high drought tolerance (Larimore et al. 1959; Iverson et al. 1978). Steffan (1961) suggested that the mosses such as Fontinalis (Figure 94) were necessary for some Elmidae and Dryopidae to make the transition from water to land (and back to the water) during their amphibious life. Bryophytes would permit them to gain a firm hold while breaking through the surface tension in either direction. In Louisiana, USA, the endangered riffle beetle Heterelmis comalensis (Figure 98-Figure 99) lives on submerged roots and aquatic mosses (Barr & Chapin 1988). In this same habitat, Microcylloepus pusillus (Figure 100Figure 101) likewise uses these substrata. In the Appalachian Mountain streams, USA, I found a species of Microcylloepus among the submerged mosses (Glime 1968).

Figure 98. Heterelmis comalensis adult, a moss dweller, as well as living on submerged roots. Photo through Creative Commons.

Figure 99. Heterelmis comalensis larva, a moss inhabitant. Photo by Mike Quinn, through Creative Commons.

Figure 100. Microcylloepus pusillus larva, an inhabitant of submerged roots and mosses. Photo by Mike Quinn, through Creative Commons.

Figure 101. Microcylloepus pusillus adult, an inhabitant of submerged roots and mosses. Photo by Mike Quinn, through Creative Commons.

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Chapter 11-10: Aquatic Insects: Holometabola – Coleoptera, Suborder Polyphaga

My experience with Elmidae among the stream bryophytes in the Appalachian Mountains, USA, differs from that in many of the reports cited here (Glime 1968). I found six species, and among these only Microcylloepus (Figure 100-Figure 101) and Promoresia elegans (Figure 91-Figure 92) (both larvae and adults) have been reported in the other studies cited herein. The numbers of Promoresia elegans actually exceeded the numbers of Chironomidae among bryophytes in one stream in March; in winter I found only two adults. In addition I found two species of Optioservus (Figure 102-Figure 103) on Hygroamblystegium fluviatile (Figure 95); on Fontinalis dalecarlica (Figure 93-Figure 94), I found Stenelmis crenata (Figure 105-Figure 104) and one species of Dubiraphia (Figure 106-Figure 107).

Figure 105. Stenelmis crenata adult, a moss dweller in Appalachian Mountain, USA, streams. Photo by Tom Murray, through Creative Commons.

Figure 102. Optioservus fastiditus adult, member of a genus that lives among mosses in Appalachian Mountain, USA, streams. Photo by Sarah McManus, through Creative Commons.

Figure 106. Dubiraphia larva. Photo by Dana R. Denson, Florida Association of Benthologists, with permission.

Figure 103. Optioservus larva, member of a genus that lives among mosses in Appalachian Mountain, USA, streams. Photo by Joseph C. Fortier, through Creative Commons.

Figure 107. Dubiraphia vittata adult. Photo by Dana R. Denson, Florida Association of Benthologists, with permission.

Figure 104. Stenelmis larvae, an inhabitant of bryophytes in Appalachian Mountain, USA, streams. Photo by Erin HayesPontius, through Creative Commons.

It is no surprise that new species remain to be discovered among the bryophytes. But one such recent discovery in the western states of the USA was not just a new species, but a new genus, widespread, and with multiple species! And these were among aquatic bryophytes, particularly leafy liverworts (Barr 2011). These three species were Bryelmis idahoensis (Figure 108), B. rivularis (Figure 109), and B. siskiyou (Figure 110) from streams and springs in the states of Washington,

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Oregon, California, and Idaho. Once Barr alerted her colleagues in neighboring states of her find, they began searching this new habitat, the leafy liverwort Chiloscyphus polyanthos rivularis (Figure 111). After searching through 652 adult and over 200 larval specimens from museum and new collections, she distinguished three species, all previously unknown. And now all these people know the difference between a moss and a liverwort – the latter houses Bryelmis.

Figure 111. Chiloscyphus polyanthos, primary home to the recently discovered genus Bryelmis. Photo by Jan-Peter Frahm, with permission.

Barr had found Bryelmis idahoensis in association with aquatic bryophytes on rocks, but some also occurred on water-soaked wood. Bryelmis rivularis preferred Chiloscyphus polyanthus rivularis (Figure 111) and Scapania undulata (Figure 112) and tended to avoid both of the mosses Fontinalis neomexicana (Figure 77) and Platyhypnidium riparioides (Figure 113). By targetting aquatic liverworts she discovered another new species, B. siskiyou. Figure 108. Bryelmis idahoensis adult male, a species that seems to be restricted to leafy liverworts. Photo by Traci Grzymala, with permission.

Figure 109. Bryelmis rivularis adult male, a species that seems to be restricted to leafy liverworts. Photo by Traci Grzymala, with permission.

Figure 110. Bryelmis siskiyou adult male, a species that seems to be restricted to leafy liverworts. Photo by Traci Grzymala, with permission.

Figure 112. Scapania undulata, home for some members of Bryelmis. Photo by Hermann Schachner, through Creative Commons.

Figure 113. Platyhypnidium riparioides, a habitat rejected by Bryelmis, a leafy liverwort inhabitant. Photo by Hermann Schachner, through Creative Commons.

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In Mexico, Central America, and the West Indies, Lara avara (Figure 114-Figure 115) spends 4-6 years as larvae, going through seven instars (Spangler & SantiagoFragoso 1992). The larvae leave the stream water in spring and move to mosses at the stream bank in their last instar. In their last instar they burrow into small "cells" under mosses at water's edge (Spangler & Santiago-Fragoso 1992) or under mosses on the upper surface of emergent logs (Elliott 2008a). When the moss dries in early summer the larvae begin pupation (Spangler & Santiago-Fragoso 1992). This pupation lasts only two or more weeks.

Sphagnum (Figure 51) mats. In rivers of Northwest Spain, Dryops luridus preferred moss substrata (Sarr et al. 2013). In the Appalachian Mountain, USA, streams, I found a species of Helichus (Figure 118) (Glime 1968).

Figure 116. Dryops luridus adult with plastron surrounding entire body, a macroplastron. Photo by Tim Faasen, with permission.

Figure 114. Lara avara adult, a species that pupates among mosses. Photo through Creative Commons.

Figure 115. Lara avara larva, a species that crawls out of the water to pupate among mosses at the water's edge. Photo by Arlo Pelegrin, with permission.

Figure 117. Dryops anglicanus adult, an inhabitant of beaver-made canals in floating Sphagnum mats. Photo by Stefan Schmidt, through Creative Commons.

Dryopidae – Long-toed Water Beetles The Dryopidae are mostly Northern Hemisphere (Dryopidae 2015), but the scant records in the Southern Hemisphere may reflect limited collecting rather than absence of beetles. This is an interesting family in that the larvae are mostly terrestrial, living in decaying plant material, rotting wood, and soil, whereas the adults (3.5-5.5 mm long) return to running water to lay eggs (Watson & Dallwitz 2003). They are unable to swim and clamber about by clinging to plants. They eat plants as adults, but larvae may also prey on small animals. The Dryopidae occur on every continent except Antarctica and Australia, but they are most common in the tropics (Dryopidae 2015). They use hairs to create a plastron apparatus (see introductory information), enabling them to breathe under water. The Dryopidae (Figure 116) seem seldom to be reported among the bryophytes of aquatic habitats. Nevertheless, Percival and Whitehead (1930) found that the Helminae (Dryopidae) reached 1244 per dm2 in the mossy area of streams in the UK, whereas among stones with no mosses they reached only 10-15 per dm2. Buczyński et al. (2014) reported that in Poland Dryops anglicanus (Figure 117) lives in canals created by beavers in floating

Figure 118. Helichus lithophilus adult, member of a genus with bryophyte dwellers in Appalachian Mountain, USA, streams. Photo by Mike Quinn, through Creative Commons.

Chapter 11-10: Aquatic Insects: Holometabola – Coleoptera, Suborder Polyphaga

Chelonariidae – Turtle Beetles These are relatively small beetles (adults 2.5-10 mm long) and somewhat resemble turtles in that their heads are hidden and their legs can be tucked into depressions in the abdomen made for them (Harpootlian 2006). They are best known from eastern North America, western South America, and Central America, but there are some records from eastern Asia (Chelonariidae 2015). They reach their greatest diversity in the Neotropics. Sometimes it is hard to determine if the insects are aquatic or terrestrial. Perhaps it is just a wide niche with a wide water tolerance. In other cases, entrance into the aquatic world may be accidental. Such seems to be the case with Chelonarium (Figure 119), a genus that inhabits damp moss (Spangler 1980). From these damp mosses, they may occasionally get washed into the nearby stream by rain or high water (Brown 1972). The larvae, once considered aquatic, lack gills (Spangler 1980). Members of the genus are often associated with the roots of terrestrial epiphytes (plants that grow on other plants but are not parasitic) and often feed on ants and termites.

Figure 119. Chelonarium lecontei adult, a species once thought to have aquatic larvae. Note how the legs fit into the exoskeleton. Photo through Creative Commons.

Lampyridae – Lightning Bugs "When night closes in, fireflies flicker with an ethereal and haunting light" (WWF 2011). This is the family of fireflies (Figure 120) that delighted us as children. And one of them, Luciola ficta (see Figure 121), lives in the water as a larva and uses mosses (Ho et al. 2010)! The adults court, mate, and females oviposit on mosses (or under leaf litter, in root gaps, or in soil clefts), but on land. The young hatchlings must make their way to the water. This unique Asian beetle is in danger of extinction because its habitat is disappearing. However, the Chinese are attempting to save it by learning its development (Ho et al. 2006) and creating small pools for it (WWF 2011).

Figure 120. Lampyridae adult showing the portion that lights up. Photo by Andy Deans, through Creative Commons.

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Figure 121. Luciola lusitanica adult. Luciola ficta is a species with aquatic larvae and adults that oviposit on terrestrial mosses. Photo by Tim Faasen, with permission.

Latridiidae – Minute Brown Scavenger Beetles Minute it is, with sizes up to 3 mm (McClarin 2005). The family mostly eats fungi and slime molds, frequenting decaying vegetation (Latridiidae 2015). Records of this family are concentrated in Europe, with scattered records in North America, South America, Africa, and Australia. But this family is even present in the Antarctic region. In South Georgia (southern Atlantic Ocean) bryophytes often play an important role as habitats for insects. One such inhabitant is Aridius malouinensis (Figure 122) (Arnold & Convey 1998).

Figure 122. Aridius malouinensis adult, a moss dweller on the island of South Georgia. Photo by Roger S. Key, with permission.

Curculionidae – Weevils Despite the fact that Curculionidae (Figure 123) is the third largest animal family (Curculionidae 2014), its presence is missing among aquatic mosses. Its distribution is worldwide, although records are lacking in vast areas of Asia and Africa (Curculionidae 2015). Adults range 1-40 mm long and are plant feeders.

Figure 123. Cionus hortulanus adult, showing one of many thousands of bizarre forms present in this family. Photo by Lukas Jonaitis, through Creative Commons.

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This terrestrial family has limited associations with the water. The marine weevil Palirhoeus eatoni, on the Prince Edward Islands south of Africa, lives among tufts of algae as larvae (Doyen 1976). When it pupates it goes above the high water level among clumps of the shoreline moss Grimmia amblyophylla (Jeannel 1940, 1953). Lagriidae Our records of bryophyte dwellers from Africa seem to be rare, so it is pleasing to see a study on bryophagy in South Africa (Chown 1993). Chown found that members of the Lagriidae in the Afromontane forest fed on both green and brown parts of the moss Braunia secunda. This is a family with poorly known feeding habits, and the species discovered here was unnamed.

Figure 124. Lagria hirta adult, a beetle that eats the moss Braunia secunda. Photo by Udo Schmidt, with permission.

bog pools, where diversity is high, but some also occur among stream bryophytes. The Hydraenidae are tiny beetles that live primarily among bryophytes in streams and fast rivers. Some small members of the Ptiliidae are parthenogenetic and live in relict bogs. The Silphidae are carrion feeders and those in bogs breed on small carrion such as frogs. The Staphylinidae are not typical bryophyte dwellers, and are not aquatic, but they live in bogs. The Scirtidae find suitable habitat in the saturated mosses of the spray zone of the springbrooks in the Alps of New Zealand. The best adapted family of the beetle bryophyte dwellers is the Elmidae. They use a plastron to breathe and are small enough to clamber about among the bryophyte stems and leaves. The Dryopidae are similarly adapted and both families can be found among stream bryophytes. Some species of the Chelonariidae live among wet mosses of stream banks and seem to occasionally fall in. The species Luciola ficta is a firefly in the family Lampyridae. Its larvae live in the water and the adults deposit their eggs on mosses and other substrata near water. The Latridiidae are among the insects in South Georgia where one species lives among the bryophytes. The Curculionidae are weevils and few are associated with aquatic habitats. Some live on floating plants and one species leaves its water home to pupate among shoreline mosses.

Acknowledgments Tim Faasen not only gave me permission to use his wide collection of insect images, he also helped me to understand the ecology of some of the species and provided me with additional images I needed. Thank you to Roger S. Key not only for his permission to use his images but for sharing his experiences with me regarding beetle use of bryophytes. Ronald Willson verified my beetle identifications for the mid-Appalachian Mountain study. Eileen Dumire proofread the chapter and offered suggestions to improve clarity.

Literature Cited

Figure 125. Braunia secunda, home for some members of the Lagriidae. Photo by Efrain De Luna, with permission.

Summary The suborder polyphaga includes a number of families of beetles that live among bryophytes, especially the small members. These include Helophoridae that live in both bogs and streams among bryophytes. Hydrochidae live among bog mosses. Hydrophilidae are common in

Anderson, Roy. 2014. Hydraena nigrita – black moss beetle. Northern Ireland Priority Species. Accessed 29 August 2014 at

. Arnett, R. H. Jr., Thomas, M. C., Skelley, P. E., and Frank, J. H. (eds.). 2002. American Beetles, Volume II: Polyphaga: Scarabaeoidea through Curculionoidea. CRC Press LLC, Boca Raton, FL. Arnold, R. J. and Convey, P. 1998. The life history of the diving beetle, Lancetes angusticollis (Curtis) (Coleoptera: Dytiscidae), on sub-Antarctic South Georgia. Polar Biol. 20: 153-160. Barr, C. B. 2011. Bryelmis Barr (Coleoptera: Elmidae: Elminae), a new genus of riffle beetle with three new species from the Pacific Northwest, USA. Coleop. Bull. 65: 197212.

Chapter 11-10: Aquatic Insects: Holometabola – Coleoptera, Suborder Polyphaga

Barr, C. B. and Chapin, J. B. 1988. The aquatic Dryopoidea of Louisiana (Coleoptera: Psephenidae, Dryopidae, Elmidae). Tulane University, New Orleans, LA. Bartlett, Troy. 2004. Family Staphylinidae – Rove Beetles. Accessed 18 February 2015 at . Beninger, C. W. and Peck, S. B. 1992. Temporal and spatial patterns of resource use among Nicrophorus carrion beetles (Coleoptera: Silphidae) in a Sphagnum bog and adjacent forest near Ottawa, Canada. Can. Entomol. 124: 79-86. Berthélemy, C. 1966. Recherches écologiques et biogéographiques sur les Plécoptères et Coléoptères d'eau courante (Hydraena et Elminthidae) des Pyrénées. Ann. Limnol. 2: 227-458. Bowles, D. E., Barr, C. B., and Stanford, R. 2003. Habitat and phenology of the endangered riffle beetle Heterelmis comalensis and a coexisting species, Microcylloepus pusillus, (Coleoptera: Elmidae) at Comal Springs, Texas, USA. Arch. Hydrobiol. 156: 361-383. Boyce, David. 2011. Invertebrate survey of blanket bog on Dartmoor, 2010. Report accessed 20 October 2014 at . Brown, H. P. 1972. Aquatic dryopoid beetles (Coleoptera) of the United States. Biota of Freshwater Ecosystems Identification Manual No. 6. Water Poll. Conf. Res. Ser., Environmental Protection Agency, Washington, DC. Buczyński, P., Przewoźny, M., Pakulnicka, J., Buczyński, E., Dawidowicz, Ł, and Wagner, G. 2014. Materials to the knowledge of beetles (Coleoptera) of aquatic habitats in the Suwalski Landscape Park. Ann. Univ. Maieae CurieSkłodowska Lublin – Polonia Sec. 2 69: 7-27. Butler, E A. 1886. Pond Life: Insects. Sonnenschein, Lowrey & Company, London, 27 pp. Chelonariidae. 2015. Encyclopedia of Life. Accessed 15 January 2015 at . Chown, S. L. 1993. Bryophagy in Lagriidae (Coleoptera) from the Drakensberg, South Africa. Coleop. Bull. 47(2): 129130. Cotinus. 2005. Family Hydrophilidae – Water Scavenger Beetles. BugGuide. Accessed 15 January 2015 at . Cowie, B. and Winterbourn, M. J. 1979. Biota of a subalpine springbrook in the Southern Alps. N. Z. J. Marine Freshwat. Res. 13: 295-301. Curculionidae. 2014. Wikipedia. Accessed 15 January 2015 at . Curculionidae. 2015. Encyclopedia of Life. Accessed 15 January 2015 at . Denton, J. 2013. The Water Beetles of North Hampshire. (VC12). Albion Ecology, Four Marks. Accessed 20 October 2014 at . Dietrich, F. and Waringer, J. A. 1999. Distribution patterns and habitat characterization of Elmidae and Hydraenidae (Insecta: Coleoptera) in the Weidlingbach near Vienna, Austria. Internat. Rev. Hydrobiol. 84: 1-15. Doyen, J. T. 1976. Marine beetles (Coleoptera excluding Staphylinidae). Marine Insects. American Elsevier Publishing Company, New York, pp. 497-519. Dryopidae. 2015. Encyclopedia of Life. Accessed 18 February 2015 at .

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Dybas, H. S. 1978. The systematics and geographical and ecological distribution of Ptiliopycna, a Nearctic genus of parthenogenetic featherwing beetles (Coleoptera: Ptiliidae). Amer. Midl. Nat. 99: 83-100. Elliott, J. M. 2008a. The ecology of riffle beetles (Coleoptera: Elmidae). Freshwat. Rev. 1: 189-203. Elliott, J. M. 2008b. Ontogenetic changes in the drifting of four species of elmid beetles elucidate the complexity of driftbenthos relationships in a small stream in Northwest England. Freshwat. Biol. 53: 159-170. EOL. 2014. Hydraenidae. Minute Moss Beetle. Accessed 13 October 2014 at . Epler, J. H. 2010. Florida Department of Environmental Protection. Tallahassee, 414 pp. Fikáček, M. 2009. Order Coleoptera, family Helophoridae. Arthropod Fauna of the UAE 2: 142-144. Foster, G. N., Nelson, B., and O’Connor, Á. 2009. A regional red list for water beetles in Ireland. Report to National Parks and Wildlife, Dublin. Friday, L. E. 1987. New records of aquatic Coleoptera from Cos Cork and Kerry. Irish Nat. J. 22: 343-345. Frost, W. E. 1942. River Liffey survey IV. The fauna of submerged "mosses" in an acid and an alkaline water. Proc. Roy. Irish Acad. Ser. B13: 293-369. Gilbert, O., Goldie, H., Hodgson, D., Marker, M., Pentecost, A., Proctor, M., and Richardson, D. 2005. The ecology of Cowside Beck, a tributary of the River Skirfare in the Malham area of Yorkshire. Field Studies Council, Settle, North Yorkshire, UK. Glime, J. M. 1968. Aquatic Insect Communities Among Appalachian Stream Bryophytes. Ph.D. Dissertation, Michigan State University, East Lansing, MI, 180 pp. Gordon, R. D. and Post, R. L. 1965. North Dakota Water Beetles. North Dakota Insects – Publication No. 5. Department of Entomology, Agricultural Experiment Station, North Dakota State University. Gurtz, M. E. and Wallace, J. B. 1984. Substrate-mediated response of stream invertebrates to disturbance. Ecology 65: 1556-1569. Haefner, J. D. and Wallace, J. B. 1981. Production and potential seston utilization by Parapsyche cardis and Diplectrona modesta (Trichoptera: Hydropsychidae) in two streams draining contrasting southern Appalachian watersheds. Environ. Entomol. 10: 433-441. Harpootlian, Phillip. 2005. Elmidae. BugGuide. Accessed 14 January 2015 at . Harpootlian, Phillip. 2006. Chelonariidae. BugGuide. Accessed 15 January 2015 at . Harrison, J. D. G. 2009. Guides to the freshwater invertebrates of Southern Africa. Volume 10: Coleoptera. African Entomol. 17: 235-237. Hebauer, F. 1994. Entwurf einer Entomosoziologie aquatischer Coleoptera in Mitteleuropa (Insecta, Coleoptera, Hydradephaga, Hydrophiloidea, Dryopoidea). Lauterbornia 19: 43-57. Helophoridae. 2014. Accessed 15 January 2015 at . Helophoridae. 2015. Accessed 15 January 2015 at . Hilsenhoff, W. L. 1975. Aquatic Insects of Wisconsin. Generic Keys and Notes on Biology, Ecology and Distribution. Tech. Bull. No. 89, Department of Natural Resources, Madison, Wisconsin, pp. 1-53.

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Ho, J. Z., Chiang, P. H., and Yang, P. S. 2006. A new rearing method for an aquatic firefly Luciola ficta (Coleoptera: Lampyridae). Formosan Entomol. 26: 77-85. Ho, J. Z., Chiang, P. H., Wu, C. H., and Yang, P. S. 2010. Life cycle of the aquatic firefly Luciola ficta (Coleoptera: Lampyridae). J. Asia-Pacif. Entomol. 13: 189-196. Hydraenidae. 2014. Wikipedia. Accessed 15 January 2015 at . Hydrochidae. 2015a. Encyclopedia of Life. Accessed 15 January 2015 at . Hydrochidae. 2015b. Order Coleoptera - Family Hydrochidae. Digital Key to Aquatic Insects of North Dakota. Accessed 15 January 2015 at . Iverson, T. M., Wiberg-Larsen, P., Hansen, S. B., and Hansen, F. S. 1978. The effect of partial and total drought on the macroinvertebrate communities of three small Danish streams. Hydrobiologia 60: 235-242. Jeannel, R. 1940. Coléopteres. Croisière du Bougainville aux îles australes françaises. Mem. Mus. Natl Hist. Nat. (N.S) 14: 63201. Jeannel, R. 1953. Sur la faune entomologique de l'ile Marion. Rev. Franc. Entomol. 31: 319-417. Knight, L. R. F. D. 2014. CSM Monitoring of Designated Aquatic Invertebrate Features at Woodhall Loch, Buckstruther Moss, Firth of Forth, Lochs of Harray & Stenness and Rannoch Moor SSSIs. Scottish Natural Heritage Commissioned Report No. 677, 72 pp. Komarek, A. 2004. Taxonomic revision of Anacaena Thomson, 1859. I. Afrotropical species (Coleoptera: Hydrophilidae). Koleopt. Rund. 74: 303-349. Larimore, R. W., Childers, W. F., and Heckrotte C. 1959. Destruction and re-establishment of stream fish and invertebrates affected by drought. Trans. Amer. Fish. Soc. 88: 261-285. Latridiidae. 2015. Encyclopedia of Life. Accessed 15 January 2015 at . Majka, C. G. 2008. The aquatic Coleoptera of Prince Edward Island, Canada: New records and faunal composition. ZooKeys 2: 239-260. Maurer, M. A. and Brusven, M. A. 1983. Insect abundance and colonization rate in Fontinalis neo-mexicana (Bryophyta) in an Idaho batholith stream, USA. Hydrobiologia 98: 9-15. McClarin, Jim. 2005. Family Latridiidae – Minute Brown Scavenger Beetles. BugGuide. Accessed 15 January 2015 at . Murray, Tom. 2005. Family Scirtidae – Marsh Beetles. BugGuide. Accessed 15 January 2015 at . Nelson, B. 1996 Species Inventory for Northern Ireland: Aquatic Coleoptera. Ulster Museum, Belfast, 36 pp. Nilsson, A. N. 1983. The larva of the predaceous water beetle Coelambus novemlineatus (Coleoptera: Dytiscidae). Aquat. Ins. 5: 45-50. Percival, E. and Whitehead, H. 1930. Biological survey of the river Wharf. II. Report on the invertebrate fauna. J. Ecol. 18: 286-295. Ptiliidae. 2015. Encyclopedia of Life. Accessed 18 January 2015 at .

Reichle, D. E. 1966. Some pselaphid beetles with boreal affinities and their distribution along the postglacial fringe. Syst. Zool. 15: 330-344. Reichle, D. E. 1967. The temperature and humidity relations of some bog pselaphid beetles. Ecology 48: 208-215. Rove Beetle. 2014. Wikipedia. Accessed 15 January 2015 at . Ruta, R. 2008. Contribution to the knowledge of Seychellois Scirtidae (Coleoptera: Scirtoidea). Zootaxa 1913: 49-68. Sarr, A. B., Benetti, C. J., Fernández-Díaz, M., and Garrido, J. 2013. The microhabitat preferences of water beetles in four rivers in Ourense province, Northwest Spain. Limnetica 31: 1-10. Shepard, W. D. and Barr, C. B. 1991. Description of the larva of Atractelmis (Coleoptera: Elmidae) and new information on the morphology, distribution, and habitat of Atractelmis wawona Chandler. Pan-Pacif. Entomol. 67: 195-199. Silphidae. 2015a. Encyclopedia of Life. Accessed 13 January 2015 at . Silphidae. 2015b. Wikipedia. Accessed 13 January 2015 at . Spangler, P. J. 1980. Chelonariid larvae, aquatic or not? (Coleoptera: Chelonariidae). Coleop. Bull. 34: 105-114. Spangler, P. J. and Santiago-Fragoso, S. 1992. The aquatic beetle subfamily Larainae (Coleoptera: Elmidae) in Mexico, Central America, and the West Indies (No. 528). Smithsonian Institution Press. Steffan, A. W. 1961. Vergleichend-mikromorphologische Genital-Untersuchungen zur Klärung der phylogenetischen Verwandtschaftsverhaltnisse der mitteleuropäischen Dryopoidea Coleoptera). Zool. Jahrb. Syst. 88: 255-354. Thorpe, W. H. and Crisp, D. J. 1949. Studies on plastron respiration. IV. Plastron respiration in the Coleoptera. J. Exper. Biol. 26: 219-260. TOL. Tree of Life Web Project. 2011. Scirtidae. Marsh beetles. Version 15 February 2011. Accessed 15 January 2015 at . Usinger, R. L. 1956. Aquatic Insects of California: With Keys to North American genera and Species. University of California Press, Berkeley, CA. Usinger, R. L. 1974. Aquatic Insects of California: With Keys to North American genera and Species. University of California Press, Berkeley, CA. Ward, J. V. 1992. Aquatic Insect Ecology. 1. Biology and Habitat. John Wiley & Sons, Inc., N. Y., 438 pp. Water Beetles. 2014. Bumblebee.org. Accessed 21 October 2014 at

. Watson, L. and Dallwitz, M. J. 2003 onwards. British Insects: The families of Coleoptera. Last updated 25 July 2012. Accessed 15 January 2015 at . Watson, L. and Dallwitz, M. J. 2012. British insects: Water beetles. Last updated 18 September 2012. Accessed 29 August 2014 at . WWF. 2011. Hope for the Fireflies in Anlong Village. Accessed 14 October 2014 at .

Glime, J. M. 2017. Aquatic Insects: Holometabola – Trichoptera, Suborder Annulipalpia. Chapt. 11-11. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. Ebook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 19 July 2020 and available at .

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CHAPTER 11-11 AQUATIC INSECTS: HOLOMETABOLA – TRICHOPTERA, SUBORDER ANNULIPALPIA TABLE OF CONTENTS LEPIDOPTERA ..................................................................................................................................................... 11-11-2 TRICHOPTERA .................................................................................................................................................... 11-11-2 Drift ................................................................................................................................................................. 11-11-4 Food ................................................................................................................................................................ 11-11-4 Case Building .................................................................................................................................................. 11-11-5 SUBORDER ANNULIPALPIA ..................................................................................................................... 11-11-6 Hydropsychoidea ..................................................................................................................................... 11-11-6 Ecnomidae ........................................................................................................................................ 11-11-6 Hydropsychidae – Net-spinning Caddisflies ..................................................................................... 11-11-6 Pupal Sites ................................................................................................................................. 11-11-7 Crowding and Niche Separation ................................................................................................ 11-11-8 Food ......................................................................................................................................... 11-11-11 Role of Water Velocity ............................................................................................................ 11-11-12 Role Below Impoundments ..................................................................................................... 11-11-14 Polycentropodidae – Tube Maker Caddisflies ................................................................................ 11-11-14 Psychomyiidae – Net Tube Caddisflies .......................................................................................... 11-11-15 Philopotamoidea .................................................................................................................................... 11-11-15 Philopotamidae – Finger-net Caddisflies ........................................................................................ 11-11-15 Summary ....................................................................................................................................................... 11-11-17 Acknowledgments ......................................................................................................................................... 11-11-17 Literature Cited ............................................................................................................................................. 11-11-17

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CHAPTER 11-11 AQUATIC INSECTS: HOLOMETABOLA – TRICHOPTERA, SUBORDER ANNULIPALPIA

Figure 1. Fontinalis antipyretica in a small stream. This moss is often home to many kinds of insects, including even larger Trichoptera. Photo by Betsy St. Pierre, with permission.

LEPIDOPTERA – Moths and Butterflies This predominantly terrestrial order has a number of aquatic members whose larvae live on tracheophytes. These include such families as the Pyralidae (Figure 2) and Noctuidae. Larvae of some aquatic species possess gills (Bouchard et al. 2004). The aquatic Pyralidae are the only Lepidpotera with aquatic pupae. I have not been able to find any records of this order on bryophytes. However, on one occasion I found a caterpillar of the Nymphalidae in a bed of Fontinalis in the Red Cedar River, East Lansing, MI. Unfortunately, I was there for a different purpose and don't have any further details.

TRICHOPTERA – Caddisflies The Trichoptera are distinguished as adults by the hairs on their wings (Figure 3) and the resting position that looks like a pup tent (Figure 4). Their distribution is worldwide and size varies greatly. Most build cases that serve as retreats for both larvae and pupae (immature stages, often immobile) between larvae and adults).

Figure 2. Petrophila larva (ventral view), a common aquatic moth that lives among aquatic plants. Photo by Bob Henricks, with permission.

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and sticks) for attachment; Tricorythodes (Ephemeroptera: Leptohyphidae) burrows among the stems and rhizoids; and the caddisfly Chimarra (Philopotamidae; Figure 6) lives in the gravel and sand at the base of the mosses, all in the riffles of one Wyoming river (Armitage 1961).

Figure 3. Brachycentrus appalachia adult wings showing hairs. Photo by Jason Neuswanger, with permission. Figure 5. Brachycentrus occidentalis larvae. Arlen Thomason, with permission.

Photo by

Figure 4. Limnephilus frijole adult showing wings folded like a pup tent. Photo by Bob Newell, with permission.

Caddisflies are common inhabitants among mosses (Oswood 1979; Glime 1994; Ogbogu 2000; Ogbogu & Akinya 2001). Berg and Petersen (in Macan 1963) found a mean of 260 Trichoptera in just 1 sq meter of Fontinalis (Figure 1) in Lake Gribso. And Frost (1942) found 492,200 individuals per gram of mosses in Ireland. Several families of caddisfly larvae have members that use bryophytes in the construction of their homes (Glime 1978). In North America, caddisfly larvae are closely associated with mosses such as Fontinalis (Figure 1) (Ogbogu 2001a). As the density of these mosses increases, so does the density of the caddisfly larvae. Ogbogu suggested that use of the mosses as part of their life cycle strategy permits these larvae to survive in the unstable habitats of streams. Krno (1990) found that some Trichoptera were able to climb out of the water to move about among the wet emergent mosses. However, the fauna there was not as rich as that among submerged mosses. Galdean (1994) found that some caddisflies were common on the mosses lining the walls of the Somequl Cald Gorges. These mosses were clean, lacking detritus (organic matter produced by the decomposition of organisms), and formed a felt on the walls. Some insect assemblages even partition the moss into several habitats. The caddisfly Brachycentrus (Brachycentridae; Figure 5) uses mosses (as well as rocks

Figure 6. Chimarra tsudai larva, member of a genus that lives in gravel and sand at the bases of mosses in riffles. Photo by Takao Nozaki, with permission.

In the case of Helicopsyche sperata (Helicopsychidae; Figure 7), the aquatic surroundings are achieved by living on mossy rocks out of the stream but in the sun in locations kept wet by constantly dropping water (McLachlan 1880).

Figure 7. Helicopsyche sp. larva and case, a genus that lives on wet mosses in the splash of streams. Photo by Stephen Moore, Landcare Research, with permission, NZ.

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Chapter 11-11: Aquatic Insects: Holometabola – Trichoptera, Suborder Annulipalpia

Drift Unlike most of the drifting aquatic insect species, many species of Trichoptera are day-active and do most of their drifting during the day (Waters 1972). This makes this group more vulnerable to predation by fish (White 1967), and this would particularly apply to the caseless caddisflies that are the most common caddisflies among bryophytes. However, Brusven (1970) found that among the caseless net-spinning caddisflies, Arctopsyche (Figure 8) drifted mostly at night and Hydropsyche (Figure 9) was rare in the drift. It is reasonable to assume that the bryophyte habitat may help to keep these caddisflies anchored as they move about, hence offering a safe refuge.

Rhyacophila dorsalis (Figure 15) had bryophyte fragments in only one out of nine larvae. An image on Garden World Images by Dave Bevan (Bevan 2014) suggests that some Stenophylax species eat mosses. (The image looks like either protonemata or a filamentous alga.)

Figure 10. Glyphotaelius pellucidus larva in its case, a genus known to eat bryophytes. Photo by Niels Sloth, with permission.

Figure 8. Arctopsyche ladogensis (Hydropsychidae) larva, a night drifter. Photo by Donald S. Chandler, with permission.

Figure 11. Limnephilus rhombicus larva showing two very different cases for the same species. This species eats bryophytes. Photo by Niels Sloth, with permission.

Figure 9. Hydropsyche pellucidula larva (Hydropsychidae), a rare drifter that can be found among bryophytes. Photo by Niels Sloth, with permission.

Food Slack (1936) compared the food of twelve species of caddisflies. Among these, all but three had bryophyte leaf fragments in the gut. Those with more than half the larvae having bryophyte fragments were Limnephilidae: Glyphotaelius sp. (Figure 10), Limnephilus rhombicus – an opportunist in using a variety of materials to build its case (Figure 11), Stenophylax sp. (Figure 12), and Halesus sp. (Figure 13) and Sericostomatidae: Sericostoma personatum (Figure 14). Among common bryophyte dwellers, Hydropsyche sp. (Figure 9) had none and

Figure 12. Stenophylax permistus adult, a genus known to eat bryophytes. Photo by Wouter Bosgra, through Creative Commons.

Chapter 11-11: Aquatic Insects: Holometabola – Trichoptera, Suborder Annulipalpia

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acetylenic acids as biomarkers of Fontinalis antipyretica (Figure 1) to demonstrate consumption of this moss by Trichoptera in the Yenisei River. Case Building

Figure 13. Halesus radiatus larva, a genus which has bryophyte consumers. Photo by Malcolm Storey, through Creative Commons.

Figure 14. Sericostoma personatum larva, a genus known to eat mosses. Photo by J. C. Schou, with permission.

Case building provides most species of Trichoptera with a mobile home that protects them from predation. Some of these case-builders use bryophytes in their construction, including the New Zealand genus Zelolessica (Helicophidae; Figure 16) that sometimes uses bryophytes exclusively (Suren 1988). Frost (1942) found that a rather dominant caddisfly in her acid site on the River Liffey, Ireland, made cases from fragments of Fontinalis (Figure 1), but the larvae were too small for identification.

Figure 16. Zelolessica, a caddisfly that sometimes uses bryophytes in case construction. Photo by Stephen Moore, Landcare Research, NZ, with permission.

Elliot and Spribille found that in a northwest Montana fen caddisfly larvae use living Scorpidium scorpioides (Figure 17) to build cases. The larvae harvest small tips of branches (ca. 2 cm) of the S. scorpioides from plants that grow submerged in shallow water and attach them to their cases. Elliot and Spribille suggested that the moss provides a "buoyant platform" from which the caddisfly can emerge, prey on the invertebrate fauna, and then fly off without being trapped by the surface tension.

Figure 15. Rhyacophila dorsalis larva, a common bryophyte dweller that had no moss in the gut of 8 out of 9 individuals. Photo by Walter Pfliegler, with permission.

Trichoptera is a large order, surpassing Ephemeroptera, Odonata, and Plecoptera in the number of genera (Wiggins & Mackay 1978). Most of the filterfeeders are in eastern North America in the deciduous forest biome. In addition to filter feeders, they are represented by grazers, especially upstream in the mountains where waters are cool. Shredders, especially in the Limnephilidae, can be found in lakes, ponds, streams, and even terrestrial habitats. Shredder-collectors are more common upstream and grazer-collectors are more common downstream. Some are predators. Cairns (2005) reported that some caddisfly larvae consumed stream mosses. Kalachova et al. (2011) used

Figure 17. Scorpidium scorpioides, a moss used for building caddisfly cases. Photo by Malcolm Storey , through Creative Commons.

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Chapter 11-11: Aquatic Insects: Holometabola – Trichoptera, Suborder Annulipalpia

SUBORDER ANNULIPALPIA Hydropsychoidea Ecnomidae This is a relatively small family with worldwide distribution (Holzenthal et al. 2007). Although records of this family are worldwide, their main distribution is Gondwanan (Ecnomidae 2014). The larvae are of moderate size (5-10 mm) and live in retreats that they construct of silk in slow-water streams or lakes. They are predators, but some eat algae and detritus. From Ceylon, Schmid (1958) reported Ecnomus ceylanicus (see Figure 18) and a new species, Ecnomus vaharika, from large, mossy rocks in the torrent.

Figure 18. Ecnomus tenellus adult, member of a genus in which some species live in mossy torrents in Ceylon. Photo by Dick Belgers, through Creative Commons.

Hydropsychidae – Net-spinning Caddisflies This worldwide family occupies a wide range of rivers and streams, always requiring flowing water to obtain its food (Hydropsychidae 2014). For example, in Ceylon Schmid (1958) reported Pseudoleptonema ceylanicum (see Figure 19) from a small, mossy creek in the jungle.

Figure 19. Pseudoleptonema supalak adult. In Ceylon, larvae of P. ceylanicum live in a mossy creek. Photo from Biodiversity Institute of Ontario, through Creative Commons.

The larvae can be relatively large, ranging 5-25 mm (Hydropsychidae 2015). The larvae of this family build retreats from plant and mineral fragments. These retreats open into the nets used to catch their food, including algae, detritus, and small animals. When another caddisfly attempts to occupy the retreat, the current occupant uses its hind legs, rubbing them under the head, to produce stridulations that warn the intruder to vacate (Jansson & Vuoristo 1979). Larvae of Hydropsyche angustipennis, H. siltalai, H. nevae, and H. pellucidula will enter any suitable retreat when forced to leave their own, and it need not be their own species or unoccupied. When it is already occupied, a vigorous fight will ensue. Larger defenders lost more fights as the size of the intruder increased. Stridulation increased the likelihood of a defender winning the fight. Several researchers have supported the importance of mosses in the habitats of net-spinning caddisflies (Sprules 1947; Tanaka 1968). Oswood (1979) found that in a lake outlet stream in Montana, USA, larvae of Hydropsychidae had greater densities on moss-covered substrata (up to >1400 0.2 m-2) than elsewhere. In a gorge of the Some River, Galdean (1994) considered the mosses on the walls of the gorge to create the conditions needed for the Hydropsychidae to develop. The boulders were cleaned by the river velocity on the concave bank, permitting the mosses, hence the Hydropsychidae, to develop there. Parapsyche cardis preferred substrata in the order of mossy rock face > cobble riffle > pebble riffle > sandy reach (Gurtz & Wallace 1986). This relationship held true for all instars (larval stages) in both studied streams. Thus, mossy rock faces accounted for 94.8% of the total production of Parapsyche (Figure 20) in Hugh White Creek (with 36.5% rocky channel) and 87.3% in Big Hurricane Branch (with 16.8% rocky channel) in the southern Appalachian Mountains, USA. Haefner and Wallace (1981a, b) likewise found that the distribution of P. cardis was highly correlated with the distribution of moss in Sawmill Branch. In several Maryland, USA, streams, Parapsyche apicalis occurred among bryophytes, mostly Fontinalis dalecarlica, and at the time were new records for Maryland, but it was not one of the more common Hydropsychidae represented among the midAppalachian bryophytes (Glime 1968).

Figure 20. Parapsyche apicalis larva, a species I collected among bryophytes in several Maryland streams. Parapsyche carda distribution is correlated with moss cover. Photo by Donald S. Chandler, with permission.

Chapter 11-11: Aquatic Insects: Holometabola – Trichoptera, Suborder Annulipalpia

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Wulfhorst (1994) examined the relative abundance of the caddisfly larva Diplectrona (Figure 29) in mosses and in interstitial spaces (spaces between individual sand grains in soil or aquatic sediments) in the hyporheic zone (region beneath and alongside a stream bed, where mixing of shallow groundwater and surface water occurs) of two streams in the Harz Mountains of West Germany. She found that Diplectrona was more abundant among the mosses at most collection stations, but that they were also abundant in the interstitial spaces of the hyporheic zone at 10 and 30 cm depths (Figure 21).

Figure 22. Hydropsyche orientalis, a species that provides shelter used by the mayfly Serratella setigera. Photo by Takao Nozaki, with permission.

Figure 21. Mean abundance ± 95% CI of Diplectrona spp. in moss clumps in two streams in the Harz Mountains, West Germany. Numbers of samples are shown at the bottom. Redrawn from Wulfhorst 1994.

The high density of Hydropsychidae among stream mosses is supported by their ability to colonize that habitat rapidly. Smith-Cuffney (1987) found that artificial mosses reached their capacity of these net-spinning colonizers in only 7 days; Georgian and Thorp (1992) found that 6-9 days provided enough time for them to reach their constant colonization density among the artificial mosses. Mosses provide a particularly easy place to colonize relative to other stream habitats because their rough surface makes it easy to gain a hold that rescues them from the speeding water. The Hydropsychidae can be considered ecosystem engineers (Nakano et al. 2005). In Japan, Hydropsyche orientalis (Figure 22, Figure 23) make their larval retreats on the upper surfaces of stones. These retreats provide a safe site for naiads of the mayfly Serratella setigera, providing them with the slower flow that they prefer. It is likely that in the absence of these caddisflies and their nets that mosses could play a similar role in creating a suitable refuge. And in some cases it appears that the hydropsychids use the mosses in place of some, but not all, nets (Figure 24). Ogbogu (2000) found Hydropsychidae associated with Fontinalis (Figure 1) in Nigeria and reported that the density of larvae increased when the moss grew. Both Cheumatopsyche (Figure 45) and Amphipsyche formed close associations and Ogbogu (2001a, b) suggested that the moss served as a refugium (area in which population of organisms can survive through period of unfavorable conditions, even glaciation) during vulnerable life cycle stages.

Figure 23. Hydropsyche orientalis net where Ephemerella setigera takes refuge. Photo by Takao Nozaki, with permission.

Figure 24. Hydropsychidae nets among mosses. Photo by Janice Glime.

Pupal Sites Frost, in her 1942 study of the River Liffey, Ireland, found that few Trichoptera pupae were present among the mosses. She considered this an expected absence because the caddisfly larvae usually seek another type of environment instead of mosses for pupation (period of development of pupa). For example, Ceratopsyche morosa (Figure 25) lives among moss and algae in young larval stages (Stern & Stern 1969), but just prior to pupation it moves to stones.

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Chapter 11-11: Aquatic Insects: Holometabola – Trichoptera, Suborder Annulipalpia

Temperature can signal that it is time to pupate. At least some Hydropsyche species cannot live below 8°C (Kaiser 1965). Instead, they build loose cases and go into the pupa state in autumn. Sleight (1913) found Hydropsyche pupae (Figure 26-Figure 28) among mosses in strong currents in the eastern USA. At maturity, these pupae moved to the surface where the pupal case would split and adults would emerge. The larval hooks made it possible for these caddis larvae to climb over the vegetation to find a suitable place for the pupa.

could account for the differences in productivity. Mosses provide a suitable substrate for attaching the nets (Figure 30) and retreats of these caddisflies while providing a range of current velocities. The nets themselves do not, however, appear to contribute directly to their food; none were found in the gut analysis (Haefner & Wallace 1981a). The larvae are also relatively common among Hygroamblystegium fluviatile (Figure 31), Platyhypnidium riparioides (Figure 32), and Fontinalis dalecarlica (Figure 33) in Appalachian Mountain streams (Glime 1968).

Figure 25. Ceratopsyche morosa larva, a moss dweller that leaves the mosses to pupate among stones. Photo by Bob Henricks, with permission.

Figure 27. Hydropsyche pupae removed from their pebble cases. Photo by Mark Melton, with permission.

Figure 26. Hydropsyche pupae, a genus that pupates among the protective mosses in strong currents. Photo by Mark Melton, with permission.

Crowding and Niche Separation It appears that mosses might separate the niches of cohabiting net spinners. Late instar Diplectrona modesta (Figure 29) has a somewhat uniform occupancy among substrata in Big Hurricane Branch (Gurtz & Wallace 1986). The first three instars are most abundant on the (mossy) rock face and the fourth and fifth are more evenly distributed. But in Hugh White Creek, the rocks have a lower density of moss, and D. modesta is less common than in Big Hurricane Branch, where the moss is thicker. In fact, in Hugh White Creek, D. modesta is most abundant in the cobble riffle and least abundant in the rock face samples, while first instars are most common on sand. Gurtz and Wallace suggested that the lower density of moss in the Hugh White Creek may not provide enough microhabitats and that differences in available substrata

Figure 28. Hydropsyche pupa, common among mosses in strong currents. Photo by Jason Neuswanger, with permission.

Figure 29. Diplectrona modesta larva, a species that is more common among mosses in early instars but is more evenly distributed between mosses and other substrata in later instars. Photo by Bob Henricks, with permission.

Chapter 11-11: Aquatic Insects: Holometabola – Trichoptera, Suborder Annulipalpia

Figure 30. Cheumatopsyche larval net. These are often attached to bryophytes and are able to trap detritus and algae. Photo by Justin Montem, through Creative Commons.

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When Cheumatopsyche sp. (Figure 34) reaches high densities it becomes more aggressive (Glass & Bovbjerg 1969). This aggressiveness dictates a pattern of dispersion (pattern of distribution of individuals within a habitat) that is a function of density. Hildrew and Edington (1979) found that larvae are able to make ultrasonic sounds to discourage intruders when they approach. Fortunately, for overlapping generations of the same species larval sizes differ at a given point in time, permitting them to use different net sizes (Figure 35-Figure 36) and avoid competition for food.

Figure 31. Hygroamblystegium fluviatile, a home for smaller insects. Photo by Michael Lüth, with permission.

Figure 34. Cheumatopsyche larva, a caddisfly that becomes less aggressive when it has shelter. Photo by Bob Henricks, with permission.

Figure 32. Platyhypnidium riparioides, a home for smaller insects, sometimes serving as food and case-building materials. Photo by David Holyoak, with permission.

Figure 33. Fontinalis dalecarlica, home to some larvae of Cheumatopsyche. Photo by J. C. Schou, with permission.

Figure 35. Hydropsyche net showing mesh size that can differ in size with species. Photo by Michael Wiesner , with permission.

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Chapter 11-11: Aquatic Insects: Holometabola – Trichoptera, Suborder Annulipalpia

Chimarra aterrima (Figure 39), a potential competitor, occupied the spaces under large stones. The two hydropsychid species share the same sites, eat the same foods, and have similar life cycles. In contrast to Chimarra aterrima, these net-spinning caddisflies have mechanisms in their gut for crushing diatoms, important constituents of the diet and one that separates their niche from that of C. aterrima.

Figure 37. Cheumatopsyche oxa larva, an occupant of mossy areas on boulders. Photo by Trevor Bringloe, Biodiversity Institute of Ontario, through Creative Commons.

Figure 38. Ceratopsyche sparna larva, a species that prefers mossy areas to those under stones. Photo by Bob Henricks, with permission. Figure 36. Nets of the net-spinning caddisfly, Cheumatopsyche, on Fontinalis. The number of larvae usually greatly exceeds the number of nets on the Fontinalis, suggesting that they may be using the mosses as nets to gather detritus and diatoms. Photos by Janice Glime.

Williams and Hynes (1973) suggested that mossy habitats provide the greatest number of protected sites. Furthermore, the rapid flow typical of locations where mosses grow will bring more food per unit of time. Cheumatopsyche (Figure 37) larvae are common among the mosses Hygroamblystegium fluviatile (Figure 31), Platyhypnidium riparioides (Figure 32), and Fontinalis dalecarlica (Figure 33) in the mid-Appalachian Mountain streams (Glime 1968). And Cheumatopsyche (Figure 34) larvae seem to be less aggressive when shelter is readily available (Glass & Bovbjerg 1969). Williams and Hynes (1973) found that the hydropsychids Cheumatopsyche oxa (Figure 37) and Ceratopsyche sparna (Figure 38) occupied the mossy areas of boulders, whereas the philopotamid

Figure 39. Chimarra aterrima larva, a species that occupies spaces under rocks in preference to that of mosses. Photo by Stroud Water Research Center, Stroud Water Research Center, through Creative Commons.

Chapter 11-11: Aquatic Insects: Holometabola – Trichoptera, Suborder Annulipalpia

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Hydropsyche pellucidula (Figure 40-Figure 41) occurs among submerged mosses in the River Rajcianka (Krno 1990). Elsewhere, when Hydropsyche pellucidula and H. siltalai (Figure 42) occur together, the moss cover is important in permitting these two caddisflies to partition the rocks into two functional feeding (net-spinning) niches and co-exist throughout their larval lives (Hildrew & Edington 1979). In late winter and early spring, there is rapid growth of moss (particularly Fontinalis antipyretica, Figure 43) on boulders and bedrock in rapids. Hydropsyche siltalai (but not H. pellucidula) migrates onto the moss in spring. Although large numbers of H. siltalai occupied the moss, not a single H. pellucidula could be found there. Plastic artificial grass, similar to moss mats, proved to be a suitable surface for net-spinning. Figure 43. Fontinalis antipyretica. Haynold, through Wikimedia Commons.

Photo by Bernd

Hydropsyche siltalai (Figure 42) filters its food with a fine-meshed net (mean 100x70 µm) while H. pellucidula (Figure 40-Figure 41) is larger and uses nets with a mean mesh of 370x240 µm (Hildrew & Edington 1979). Migration of H. siltalai onto mosses (Fontinalis antipyretica; Figure 43) in spring further separates their niches. Englund (1993) observed that whereas small IV instar larvae were able to construct nets on the mosses, the physical structure seemed unsuitable for the larger V instar larvae to do so. Figure 40. Hydropsyche pellucidula larva, a species that occurs among mosses in the River Rajcianka of Slovakia. Photo by Niels Sloth, with permission.

Figure 41. Hydropsyche pellucidula larva showing the large jaws. Photo by Niels Sloth, with permission.

Figure 42. Hydropsyche siltalai larva, a species that migrates to mosses to avoid competition from H. pellucidula. Photo by Urmas Kruus, with permission.

Food Although Frost (1942) reported several studies in which Hydropsyche instabilis ate primarily Chironomidae, and Slack (1936) found that it ate diatoms, it also ingests mosses. In Great Britain (Percival & Whitehead 1929) and in calcareous streams in South Wales, Hydropsyche instabilis (Figure 44) ingested Fontinalis antipyretica (Figure 43) (Percival & Whitehead 1929; Jones 1949). Frost (1942) found that Hydropsyche instabilis (Figure 44) lived primarily among mosses in an acid stream, but in the alkaline stream it was Cheumatopsyche lepida (Figure 45) that was dominant among the mosses, in this case where there was more silt. Jones (1950) did extensive gut analysis of insects from the River Rheidol; among the Trichoptera, only Hydropsyche instabilis of the six species examined had fragments of Fontinalis antipyretica (Figure 43) in the gut (7 out of 27). Fragments of this moss were present in nine of the 23 analyses with identifiable gut contents (Jones 1949). Algae and detritus were the most common foods.

Figure 44. Hydropsyche instabilis adult, a species whose larvae sometimes eat mosses. Photo from Biodiversity Institute of Ontario, through Creative Commons.

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Chapter 11-11: Aquatic Insects: Holometabola – Trichoptera, Suborder Annulipalpia

Figure 45. Cheumatopsyche lepida larva, a dominant caddisfly among mosses with lots of silt in an alkaline stream. Photo through Creative Commons.

On the other hand, occurrence of net-spinning caddisflies among mosses may offer the advantage of a greater number of prey organisms. Although these insects trap their food on finely constructed nets, they are also carnivores. Haefner (1980) found a significantly higher (2x) density of prey organisms (Baetis spp., Ephemerella spp., Nemoura spp., Hydroptila sp., and Chironomidae) in rock face samples, where mosses were typically dense. These organisms are common among stream mosses – Hydroptila less so (Glime 1994), thus the abundance of prey invertebrates may account for the greater productivity of Parapsyche cardis (see Figure 20) there. Although Diplectrona modesta (Figure 29) had little correlation with mossy rocks in one of two Appalachian Mountain streams, and few such rocks existed in the other (Haefner & Wallace 1981a,b), this and other studies (Gurtz & Wallace 1986) suggest that the mosses provide a variety of niches that benefit both the potential prey organisms and the net-spinning caddisflies. In a study to determine the source of foods for aquatic invertebrates, Torres-Ruiz et al. (2007) used the distinctive fatty acids for green algae, diatoms, and bryophytes, each of which also differed from fatty acids of terrestrial food sources. They determined that Hydropsyche spp. (Figure 40-Figure 42) consumed primarily autochthonous (originating from within the stream system) food sources, not the terrestrial allochthonous (originating from elsewhere) food such as leaf litter. In Appalachian Mountain streams the Hydropsychidae, including species of Hydropsyche, seemed to use the mosses instead of constructing nets to capture their food (Glime 1968). There always seemed to be many more larvae than nets. Gut pH is often important in determining the digestible food sources. Hydropsyche betteni (Figure 46-Figure 47) had a gut pH close to neutral but somewhat alkaline (Barlocher & Porter 1986). Hence, this species was unable to hydrolyze (break down a compound by chemical reaction with water) proteins of maple leaves that were not yet conditioned by decomposer organisms. They could, however, digest starch and laminarin (storage product in many seaweeds). Unlike those in the cranefly Tipula, the fungal carbohydrases (enzymes that break down carbohydrates) ingested with decomposing leaves remained active in the guts of this species.

Figure 46. Hydropsyche betteni larva, with a gut pH that is alkaline. Photo by Bob Henricks, with permission.

Figure 47. Hydropsyche betteni larva showing ventral gills. Photo by Donald S. Chandler, with permission.

Role of Water Velocity The larvae of the Hydropsychidae are able to partition the niches of the most immature from those of the nearly mature (Osborne & Herricks 1987; Muotka 1990). Osborne and Herricks (1987) found that Hydropsyche (Figure 40-Figure 42) species in their study separated the larger larvae into communities at higher velocities, whereas the smaller, less mature larvae sought areas of diminished flow. The same size distribution occurs between species. These larvae seek out depressions where they can gather passing detritus but where sedimentation is minimal. Turbulence seems to play a role in determining distribution, perhaps contributing to food availability and preventing

Chapter 11-11: Aquatic Insects: Holometabola – Trichoptera, Suborder Annulipalpia

sedimentation. Larger larvae are apparently able to occupy greater velocities; this is coupled with the construction of a larger mesh size, hence dividing the feeding niche from that of smaller larvae. The net-spinning caddisflies prefer a habitat with a stable substrate and high water velocity. Georgian and Thorp (1992) showed that 96% of the Hydropsychidae larvae selected artificial moss substrates that had high velocity water flowing over them. They estimated that a prey item would be consumed within 5.5 m of travel in the drift. It appears that one advantage afforded these moss dwellers is that they can take advantage of high-flow rates while themselves finding a flow-rate suitable for their own safety. Current speed also influences net-spinning activity, with a greater percentage of larvae spinning nets at 20 cm sec-1 (73%) than at 10 cm sec-1 (10%) (Edington 1965). Edington found that hydropsychid larvae formed tunnels into the moss mats with nets at the moss surface. When the nets were removed (and when they were not) and the flow was artificially reduced, the larvae moved to a different area. When something restricts the flow, the larvae move to a new location and construct new nets (Edington 1965, 1968). Muotka (1990) considered that it was the flow pattern, rather than the flow velocity itself, that determined the pattern of occupancy by filter-feeding caddisfly larvae. He based this on the ability of multiple sizes of caddisflies, including Hydropsyche (Figure 40-Figure 42) to coexist at the same flow rates. Nevertheless, he concluded that species were often ecologically closer to other species than to other instars of their own species. In their study, many of the sites were covered with bryophytes [mosses Fontinalis antipyretica (Figure 43), Cratoneuron commutatum (Figure 48), leafy liverwort Jungermannia exsertifolia (Figure 49)] and the uneven surface of this substrate would create multiple flow patterns. It is noteworthy that in the stream that lacked bryophytes only one filter-feeding caddisfly was present – Hydropsyche saxonica (Figure 50) – whereas seven species occurred in the two streams with heavy bryophyte cover.

Figure 48. Cratoneuron commutatum, a moss that alters flow patterns, as it is doing here. Photo through Creative Commons.

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Figure 49. Jungermannia exsertifolia ssp cordifolia, contributor to flow patterns that allow niche partitioning for Hydropsychidae. Photo by Michael Lüth, with permission.

Figure 50. Hydropsyche saxonica larva, the only filterfeeding caddisfly in a stream with no mosses. Photo by Niels Sloth, with permission.

Food capture is important in the location of nets, and water velocity helps to determine the food available. Mosses on the rocks actually prevent some insects from living there. The caddisfly Leucotrichia (Hydroptilidae; Figure 51) is unable to live on a substrate dominated by heavy moss growth and instead the net spinner Hydropsyche (Figure 40-Figure 42) occupies those locations (McAuliffe 1983). The larvae arrange their nets very evenly downstream but are often crowded across the substrate, preventing the water from being filtered by a net above them.

Figure 51. Leucotrichia pictipes larva, a genus that cannot live on a substrate with heavy moss cover. Photo by Stroud Water Research Center, through Creative Commons.

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Chapter 11-11: Aquatic Insects: Holometabola – Trichoptera, Suborder Annulipalpia

As I already noted in the Appalachian Mountain streams, some caddisflies actually use the mosses to help them gather food. Hildrew and Edington (1979) found that favorable situations for net-spinning caddis larvae (Hydropsychidae), such as moss covered rocks, often seem to be occupied to capacity. I have observed the same relationship, but it appeared that the caddisflies in some cases took advantage of the collecting ability of the moss and did not make nets. This would be useful for those species that eat primarily small invertebrates living among the bryophytes (Ross & Wallace 1983), but it could also take advantage of the bryophytes as filter traps. Role Below Impoundments Mosses are important habitats at impoundments. In Valley Creek in Minnesota, USA, Hydropsychidae caddisflies use mosses and filamentous algae as sites for attachment and building materials for retreats, with the mosses providing an environment that protects the larvae from the abrasive sand deposited by the impoundment (Mackay & Waters 1986). Ogbogu (2000; Ogbogu & Akinya 2001) likewise found that Fontinalis (Figure 1) was important to the Hydropsychidae in an impoundment at Ile-Ife, Nigeria. They occupied the spillway, among the Fontinalis, in large numbers when sampled in August (1233 m-2), September (900 m-2), and November (1178 m-2). The moss provided refuge from the rapid water of the spillway, protection from predators, and food (epiphytic diatoms and other algae) trapped among the mosses. Polycentropodidae – Tube Maker Caddisflies Members of this worldwide family are relatively small to moderate in size, with the forewing reaching 6-13 mm (Hickin 1967). Larvae live in both quiet and flowing waters and trap their food in a tube (Murray 2006). Polycentropus (Figure 52) is not a caddisfly one thinks of as a moss dweller because of its long, tubular net. But in both Ballysmuttan and Straffan, UK, it does occur among mosses, as well as other locations (Frost 1942). Percival and Whitehead (1929) found that Polycentropus flavomaculatus (Figure 52) was most abundant in thick mosses compared to other types of substrate. In midAppalachian Mountain streams, larvae of this genus are occasional inhabitants of bryophytes (Glime 1968).

Figure 52. Polycentropus flavomaculatus larva, a species that is more abundant in thick mosses than elsewhere. Photo by Dragiša Savić, with permission.

In one location in the Pyrénées Décamps (1967) found that Plectrocnemia scruposa (see Figure 53) comprised 4.5% of the Trichoptera fauna among mosses. Edington (1965) found that Plectrocnemia conspersa (see Figure 53) spun more nets at a flow rate of 10 cm sec-1 (80% of the larvae) than at 20 cm sec-1 (4%), a relationship just the opposite of that of Hydropsyche instabilis. Furthermore, in both species, those few making nets at the less favorable flow rate had a tendency to construct abberrant nets.

Figure 53. Plectrocnemia geniculata larva, member of a genus in which some larvae live among mosses Photo from Biodiversity Institute of Ontario, through Creative Commons.

From Ceylon, Schmid (1958) reported Nyctiophylax devanampriya (Figure 54), Pseudoneureclipsis watagoda (Figure 55), and P. thuparama from large, mossy rocks in the torrent.

Figure 54. Nyctiophylax sp larva; N. devanampriya occurs among mosses in torrents in Ceylon. Photo by Dana R. Denson Florida Association of Benthologists, with permission.

Chapter 11-11: Aquatic Insects: Holometabola – Trichoptera, Suborder Annulipalpia

Figure 55. Pseudoneureclipsis adult, a genus whose naiads can live on mossy rocks in torrents. Photo by Biodiversity Institute of Ontario, through Creative Commons.

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Figure 57. Psychomyia flavida larva. Psychomyia pusilla eats mosses. Photo from Stroud Water Research Center through Creative Commons, with permission.

But this family relies primarily on food trapped in its funnel-shaped net. Ross and Wallace (1983) demonstrated that 80% of the food for this family in a southern Appalachian Mountain, USA, stream was fine detritus. Another 15% was diatoms. So why do we find them among bryophytes at all? Psychomyiidae – Net Tube Caddisflies The Psychomyiidae are widespread, but are concentrated in the Oriental Region and absent in the Neotropical Region (Kjer 2010a). The adults are of moderate size (5-8 mm long forewings) (Watson & Dallwitz 2003). This family traps its food in a silken tube (Figure 56), with the diet consisting of algae, leaves, and animal matter (Neuswanger 2015). Grazing may occur both on the tubes and nearby, therefore consisting mostly of diatoms and other algae (Holzenthal et al. 2007; Kjer 2010a). Females dive to the bottom of the stream to lay their eggs (Neuswanger 2015).

Figure 58. Tinodes waeneri larva, a species that consumes mosses. Photo by Niels Sloth, with permission.

Figure 59. Tinodes waeneri larval tube. Photo by Niels Sloth, with permission.

Philopotamoidea Philopotamidae – Finger-net Caddisflies Figure 56. Psychomyiidae net. Photo by Janice Glime.

Mosses occurred in the guts of Psychomyia pusilla (see Figure 57) and Tinodes waeneri (Figure 58-Figure 59) in UK streams (Percival & Whitehead 1929), attesting to their residence among bryophytes.

The larvae of this worldwide family build nets that can require more than 1 km of silk (Wallace & Malas 1976); these are used to trap small particles for food (McLeod 2005). To use them, the larvae are restricted to fastflowing water of rivers and streams. The adult body is 5-9 mm long.

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Chapter 11-11: Aquatic Insects: Holometabola – Trichoptera, Suborder Annulipalpia

The net-building behavior would seem to preclude mosses as a substrate, but exceptions occur. Philopotamus montanus is not typically a bryophyte inhabitant and captures its food with a tube net. But this net can trap bits of mosses travelling downstream, and of the 15 guts with identifiable contents, two had Fontinalis antipyretica (Figure 43) (Jones 1949). Chimarra (Figure 39; Figure 60-Figure 65) lives among mosses but prefers the gravel and sand at their bases (Armitage 1961). Williams and Hynes (1973) suggested that the affinity of C. aterrima (Figure 39) for mosscovered rocks may have been more related to the large size of those rocks rather than the presence of the moss. For example, in a wooded Ontario, Canada, stream, Wormaldia moesta (Figure 66) preferred bare stones, whereas Rhyacophila minor (Rhyacophilidae) preferred mosscovered stones in the same area (Singh et al. 1984). Wormaldia moesta grazed on diatoms when its primary food supply, detritus/seston (living organisms and nonliving matter swimming or floating in a water body), became scarce. In my own studies of the fauna of bryophytes in the Appalachian Mountain streams, C. aterrima was occasionally present, but in small numbers, among Fontinalis dalecarlica (Figure 33) in larger streams (Glime 1968). It was absent in the other bryophytes.

Figure 62. Chimarra pupa showing on underside of sand case. Photo by Mark Melton, with permission.

Figure 63 Chimarra pupa removed from sand case, showing shed sclerotized parts from larva inside the pupal covering. Photo by Mark Melton, with permission.

Figure 64. Chimarra pupa removed from case. Photo by Mark Melton, with permission. Figure 60. Chimarra tsudai tubes with thallose liverworts at the funnel opening. Photo by Takao Nozaki, with permission.

Figure 61. Chimarra pupal case. Photo by Mark Melton, with permission.

Figure 65. permission.

Chimarra tsudai adult. Takao Nozaki, with

Chapter 11-11: Aquatic Insects: Holometabola – Trichoptera, Suborder Annulipalpia

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Summary

Figure 66. Wormaldia moesta larva, a species that prefers bare stones even when mosses are present. Photo by Donald S. Chandler, with permission.

Another occasional visitor to bryophytes in Appalachian Mountain, USA, streams was Dolophilodes distinctus (Figure 67) (Glime 1968). In this case, it occurred among all four of the primary bryophytes in the study: Hygroamblystegium fluviatile (Figure 31), Platyhypnidium riparioides (Figure 32), Fontinalis dalecarlica (Figure 33), and Scapania undulata (Figure 68), preferring the mats and turfs over Fontinalis streamers.

Figure 67. Dolophilodes distinctus larva, an occasional visitor to Appalachian Mountain stream bryophytes. Photo by Donald S. Chandler, with permission.

Lepidoptera apparently do not use aquatic bryophytes. Trichoptera, on the other hand, are among the common inhabitants. Those that enter the drift may use bryophytes as a means to get out of the drift. Some larvae use the bryophytes for food and many use them as a safe site for capturing food, using both filtering strategies and predation of smaller inhabitants. The mosses themselves may serve as filter traps for caddisfly food, including drifting algae, bacteria, decomposing organic matter, and detritus. For some caddisflies the bryophytes themselves serve as food and may be a seasonal staple when other foods are unavailable. Some build their cases from bryophytes and liver among the bryophytes to capture food. Larvae of most Trichoptera are aquatic, and many may also use the bryophytes as a site for pupation and emergence. The most common families among bryophytes are The Hydropsychidae and Rhyacophilidae. These are both caseless caddisflies, and the bryophytes may provide some of the protection otherwise afforded by cases. Hydropsychidae take advantage of the bryophytes to partition their niches and avoid competition for food. In some cases this is the result of changing diets at later instar stages. Others use differences in flow within the bryophyte mat. They seem to be able to use the bryophytes to trap food, and the bryophytes create locations with a variety of flow regimes. Still other caddisflies are selective about which species of bryophytes they use, with a few selecting leafy liverworts only and others avoiding them. The importance of the bryophytes as food remains a mystery. It is possible they are ingested along with adhering periphyton and detritus without being digested.

Acknowledgments As a graduate student I relied heavily on the expertise of Oliver Flint, Glenn Wiggins, Tom Waters, and Ken Cummins for both encouragement and identification help in my novice years. Thank you so much to Donna Bennett for making a special trip to photograph live Micrasema wataga eating and showing the mosses sprouting on the case, just for this chapter! She also made additional trips to the field to determine the identity of the moss. David Tempelman helped me to obtain some of the images and permission for use and provided me with references on Ptilocolepus. Eileen Dumire proofread for me and made suggestions to improve clarity.

Literature Cited

Figure 68. Scapania undulata, a leafy liverwort that can modify flow patterns and house insects. Photo by Michael Lüth, with permission.

Armitage, K. B. 1961. Distribution of riffle insects of Firehole River, Wyoming. Hydrobiologia 17: 152-174. Barlocher, F. and Porter, C. W. 1986. Digestive enzymes and feeding strategies of three stream invertebrates. J. N. Amer. Benthol. Soc. 5: 58-66.

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Bevan, Dave. 2014. Garden World Images. Accessed 8 November 2014 at . Bouchard, R. W., Ferrington, L. C., and Karius, M. L. 2004. Lepidoptera. Chapter 11. In: Guide to Aquatic Invertebrates of the Upper Midwest: Identification Manual for Students, Citizen Monitors, and Aquatic Resource Professionals. University of Minnesota, Water Resources Research Center. Brusven, M. A. 1970. Drift periodicity and upstream dispersion of stream insects. J. Entomol. Soc. Brit. Columb. 67: 48-59. Cairns, A. 2005. Consumption of stream mosses by caddis fly larvae. 10. (Abstract) In: VIIIth Australasian Bryophyte Workshop. James Cook University. Paluma, North Queensland, Australia. Décamps, H. 1967. Ecologie des trichoptères de la Vallée d'Aure (Hautes Pyrénées). Ann. Limnol. 3: 399-577. Ecnomidae. 2014. Wikipedia. Accessed 21 January 2015 at . Edington, J. M. 1965. The effect of water flow on populations of net-spinning Trichoptera. Mitt. Intern. Ver. Theoret. Angew. Limnol. 13: 40-48. Edington, J. M. 1968. Habitat preferences in net-spinning caddis larvae with special reference to the influence of water velocity. J. Anim. Ecol. 37: 675-692. Englund, G. 1993. Effects of density and food availability on habitat selection in a net-spinning caddis larva, Hydropsyche siltalai. Oikos 68: 473-480. Frost, W. E. 1942. River Liffey survey IV. The fauna of submerged "mosses" in an acid and an alkaline water. Proc. Roy. Irish Acad. Ser. B13: 293-369. Galdean, N. 1994. Biological division of the Someș River into zones according to mayflies fauna (Insecta: Ephemeroptera). Trav. Mus. Hist. Nat. Grigorc Antipa 34: 435-454. Georgian, T. and Thorp, J. H. 1992. Effects of microhabitat selection on feeding rates of net-spinning caddisfly larvae. Ecology 73: 229-240. Glass, L. W. and Bovbjerg, R. V. 1969. Density and dispersion in laboratory populations of caddisfly larvae (Cheumatopsyche; Hydropsychidae). Ecology 50: 10821084. Glime, J. M. 1968. Aquatic Insect Communities Among Appalachian Stream Bryophytes. Unpublished Ph.D. Dissertation, Michigan State University, East Lansing, MI, 180 pp. Glime, J. M. 1978. Insect utilization of bryophytes. Bryologist 81: 186-187. Glime, J. M. 1994. Bryophytes as homes for stream insects. Hikobia 11: 483-497. Gurtz, M. E. and Wallace, J. B. 1986. Substratum-production relationships in net-spinning caddisflies (Trichoptera) in disturbed and undisturbed hardwood catchments. J. N. Amer. Benthol. Soc. 5: 230-236. Haefner, J. D. 1980. The effects of old field succession on stream insects and production of two net-spinning caddisflies. M. S. Thesis, Univ. Georgia, Athens, 61 pp. Haefner, J. D. and Wallace, J. B. 1981a. Production and potential seston utilization by Parapsyche cardis and Diplectrona modesta (Trichoptera: Hydropsychidae) in two streams draining contrasting southern Appalachian watersheds. Environ. Entomol. 10: 433-441. Haefner, J. D. and Wallace, J. B. 1981b. Shifts in aquatic insect populations in a first-order southern Appalachian stream

following a decade of old field succession. Can. J. Fish. Aquat. Sci. 38: 353-359. Hickin, N. E. 1967. Caddis larvae: Larvae of the British Trichoptera. Fairleigh Dickinson University Press, Madison, Hildrew, A. G. and Edington, J. M. 1979. Factors facilitating the coexistence of hydropsychid caddis larvae (Trichoptera) in the same river system. J. Anim. Ecol. 48: 557-576. Holzenthal, R. W., Blahnik, R. J., Prather, A. L., and Kjer, K. M. 2007. Order Trichoptera Kirby, 1813 (Insecta), Caddisflies. Zootaxa 1668: 639-698. Hydropsychidae. 2014. Wikipedia. Accessed 21 January 2015 at . Hydropsychidae. 2015. Australian Freshwater Invertebrates. Accessed 21 January 2015 at . Jansson, A. and Vuoristo, T. 1979. Significance of stridulation in larval Hydropsychidae (Trichoptera). Behaviour 71: 197186. Jones, J. R. E. 1949. A further ecological study of calcareous streams in the 'Black Mountain' district of South Wales. J. Anim. Ecol. 18: 142-159. Jones, J. R. E. 1950. A further ecological study of the river Rheidol: The food of the common insects of the mainstream. J. Anim. Ecol.19: 159-174. Kaiser, P. 1965. Über Netzbau und Strömungssinn bei der Larven der Gattung Hydropsyche Pict. Internat. Rev. Ges. Hydrobiol. Hydrogr. 50: 169-224. Kalachova, G. S., Gladyshev, M. I., Sushchik, N. N., and Makhutova, O. N. 2011. Water moss as a food item of the zoobenthos in the Yenisei River. Central Eur. J. Biol. 6: 236-245. Kjer, Karl. 2010a. Psychomyiidae. The Tree of Life Web Project. Accessed 21 January 2015 at . Krno, I. 1990. Longitudinal changes in the structure of macrozoobenthos and its microdistribution in natural and moderately eutrophicated waters of the River Rajcianka (Strázovské vrchy). Acta Fac. Rer. Natur. Univ. Comen. Zool 33: 31-48. Macan, T. T. 1966. Freshwater Ecology. Wiley & Sons Inc., N. Y., 338 pp. Mackay, R. J. and Waters, T. F. 1986. Effects of small impoundments on hydropsychid caddisfly production in Valley Creek, Minnesota. Ecology 67: 1680-1686. McAuliffe, J. R. 1983. Competition, colonization patterns, and disturbance in stream benthic communities. In: Barnes, J. R. and Minshall, G. W. (eds.). Stream Ecology: Application and Testing of General Ecological Theory. Plenum Press, N. Y., pp. 137-156. McLachlan, R. 1880. A monographic revision of the synopsis of Trichoptera of the European Fauna. Published in 9 parts 1874-1880. McLeod, Robin. 2005. Family Philopotamidae – Fingernet Caddisflies. BugGuide. Accessed 21 January 2015. . Muotka, T. 1990. Coexistence in a guild of filter feeding caddis larvae: Do different instars act as different species? Oecologia 85: 281-292. Murray, Tom. 2006. Family Polycentropodidae – Tube Maker Caddisflies. Accessed 21 January 2015 at . Nakano, D., Yamamoto, M., and Okino, T. 2005. Ecosystem engineering by larvae of net-spinning stream caddisflies

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creates a habitat on the upper surface of stones for mayfly nymphs with a low resistance to flows. Freshwat. Biol. 50: 1492-1498. Neuswanger, Jason. 2015. Caddisfly Family Psychomyiidae. Accessed 21 January 2015 at . Ogbogu, S. S. 2000. Submerged beds of Fontinalis sp. (Bryophyta) as a microhabitat for caddisfly larvae (Trichoptera) in an intermittent reservoir spillway, Ile-Ife, Nigeria. Trop. Freshwat. Biol. 9: 11-16. Ogbogu, S. S. 2001a. Dynamics of caddisfly (Trichoptera) larvae associated with Fontinalis (Bryophyta) in an intermittent reservoir outflow. North American Benthological Society. Accessed on 16 April 2008 at . Ogbogu, S. S. 2001b. Observations on the seasonal dynamics of caddisfly larvae (Trichoptera) in an intermittent reservoir outflow at Ile-Ife, Nigeria. J. Aquat. Sci. 16(2): 139-143. Ogbogu, S. S. and Akinya, T. O. 2001. Distribution and abundance of insect orders in relation to habitat types in Opa Stream-Reservoir System, Nigeria . J. Aquat. Sci. 16: 7-12. Osborne, L. L. and Herricks, E. E. 1987. Microhabitat characteristics of Hydropsyche (Trichoptera: Hydropsychidae) and the importance of body size. J. N. Amer. Benthol. Soc. 6: 115-124. Oswood, M. W. 1979. Abundance patterns of filter-feeding Caddisflies (Trichoptera: Hydropsychidae) and seston in a Montana (U.S.A.) lake outlet. Hydrobiologia 63: 177-183. Percival, E., and Whitehead, H. 1929. A quantitative study of the fauna of some types of stream-bed. J. Ecol. 17: 282-314. Ross, D. H. and Wallace, J. B. 1983. Longitudinal patterns of production, food consumption, and seston utilization by netspinning caddisflies (Trichoptera) in a southern Appalachian stream (USA). Holarct. Ecol. 6: 270-284. Schmid, F. 1958. Trichopteres de Ceylan. Arch. Hydrobiol. 54: 1-173. Singh, M. P., Smith, S. M., and Harrison, A. D. 1984. Life cycles, microdistribution, and food of two species of caddisflies (Trichoptera) in a wooded stream in southern Ontario. Can. J. Zool. 62: 2582-2588. Slack, H. D. 1936. The food of caddis fly (Trichoptera) larvae. J. Anim. Ecol. 5: 105-115.

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Sleight, C. E. 1913. Relations of Trichoptera to their environment. J. N. Y. Entomol. Soc. 21: 4-8. Smith-Cuffney, F. L. 1987. Ecological interactions in the moss habitat of streams draining a clearcut and a reference watershed. Unpublished Ph.D. thesis, University of Georgia, Athens, 174 pp. Sprules, W. M. 1947. An ecological investigation of stream insects in Algonquin Park, Ontario. Univ. Toronto Stud. Biol. 56: 1-81. Stern, M. S. and Stern, D. H. 1969. A limnological study of a Tennessee cold springbrook. Amer. Midl. Nat 82: 62-82. Suren, A. M. 1988. Ecological role of bryophytes in high alpine streams of New Zealand. Internat. Ver. Theor. Angew. Limnol. 23: 1412-1416. Tanaka, H. 1968. The aggregation of net-spinning caddisworms in the drainage ditch of the fish-farm ponds. Bull. Freshwat. Fish. Res. Lab. Tokyo 18: 71-79. Torres-Ruiz, M., Wehr, J. D., and Perrone, A. A. 2007. Trophic relationships in a stream food web: Importance of fatty acids for macroinvertebrate consumers. J. N. Amer. Benthol. Soc. 26: 509-522. Wallace, J. B. and Malas, D. 1976. The fine structure of capture nets of larval Philopotamidae (Trichoptera), with special emphasis on Dolophilodes distinctus. Can. J. Zool. 54: 1788-1802. Waters, T. F. 1972. The drift of stream insects. Ann. Rev. Entomol. 17: 253-271. Watson, L. and Dallwitz, M. J. 2003 onwards. British insects: The families of Trichoptera. Version: 9 April 2007. Accessed 20 July 2007 at . White, D. A. 1967. Trophic dynamics of a wild brook trout stream. Unpublished Ph.D. thesis. Univ. Wisconsin, Madison. Wiggins, G. B. and Mackay, R. J. 1978. Some relationships between systematics and trophic ecology in Nearctic aquatic insects, with special reference to Trichoptera. Ecology 59: 1211-1220. Williams, N. E. and Hynes H. B. N. 1973. Microdistribution and feeding of the net-spinning caddisflies (Trichoptera) of a Canadian stream. Oikos 24: 73-84. Wulfhorst, J. 1994. Selected faunal elements of the hyporheos and in submerged moss clumps (bryorheal) along an acidification gradient in two brooks in the Harz Mountains, West Germany. Verh. Internat. Verein. Limnol. 25: 15751584.

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Glime, J. M. 2017. Aquatic Insects: Holometabolous Insects – Trichoptera, Suborders Integripalpia and Spicipalpia. Chapt. 11-12. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. Ebook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 19 July 2020 and available at .

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CHAPTER 11-12 AQUATIC INSECTS: HOLOMETABOLA – TRICHOPTERA, SUBORDERS INTEGRIPALPIA AND SPICIPALPIA TABLE OF CONTENTS Suborder Integripalpia ..................................................................................................................................... 11-12-2 Leptoceroidea ........................................................................................................................................... 11-12-2 Odontoceridae – Mortarjoint Casemakers ........................................................................................ 11-12-2 Limnephiloidea ........................................................................................................................................ 11-12-3 Goeridae ............................................................................................................................................ 11-12-3 Limnephilidae – Northern Caddisflies .............................................................................................. 11-12-4 Lepidostomatidae – Bizarre Caddisflies ......................................................................................... 11-12-11 Oeconesidae .................................................................................................................................... 11-12-11 Uenoidae ......................................................................................................................................... 11-12-12 Phryganeoidea ........................................................................................................................................ 11-12-13 Brachycentridae – Humpless Casemaker Caddisflies ..................................................................... 11-12-13 Brachycentrus .......................................................................................................................... 11-12-13 Micrasema ............................................................................................................................... 11-12-14 Adicrophleps hitchcockii.......................................................................................................... 11-12-16 Phryganeidae – Giant Casemakers .................................................................................................. 11-12-17 Sericostomatoidea .................................................................................................................................. 11-12-18 Beraeidae ........................................................................................................................................ 11-12-18 Conoesucidae .................................................................................................................................. 11-12-18 Helicophidae ................................................................................................................................... 11-12-18 Sericostomatidae – Bushtailed Caddisflies ..................................................................................... 11-12-19 Suborder Spicipalpia ..................................................................................................................................... 11-12-20 Glossosomatoidea .................................................................................................................................. 11-12-20 Glossosomatidae – Tortoise or Saddle-case Makers ....................................................................... 11-12-20 Hydroptiloidea ....................................................................................................................................... 11-12-20 Hydroptilidae – Microcaddisflies, Purse-case Caddisflies .............................................................. 11-12-20 Ptilocolepus ............................................................................................................................. 11-12-23 Palaeagapetus ......................................................................................................................... 11-12-23 Scelotrichia .............................................................................................................................. 11-12-24 Rhyacophiloidea .................................................................................................................................... 11-12-26 Rhyacophilidae – Free-living Caddisflies ....................................................................................... 11-12-26 Food ......................................................................................................................................... 11-12-27 Substrate Preference ................................................................................................................ 11-12-28 Summary ....................................................................................................................................................... 11-12-29 Acknowledgments ......................................................................................................................................... 11-12-30 Literature Cited ............................................................................................................................................. 11-12-30

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CHAPTER 11-12 AQUATIC INSECTS: HOLOMETABOLA – TRICHOPTERA, SUBORDERS INTEGRIPALPIA AND SPICIPALPIA

Figure 1. Adicrophleps hitchcocki (Brachycentridae), a larva that makes its case from mosses. Note the "furry" portion near the opening. Photo by D. N. Bennett, with permission.

SUBORDER INTEGRIPALPIA Leptoceroidea Odontoceridae – Mortarjoint Casemakers This worldwide family lives in springs and small to medium streams and rivers, typically with slow flow; some are associated with waterfalls (Holzenthal et al. 2010c). Also known as the strong case-maker caddis, the larvae make very strong cases from bits of rock with more than usual amounts of the silk glue (Henricks 2011). Although I never found Pseudogoera in my studies of stream insects among bryophytes in the mid Appalachians, P. singularis (Figure 2) is associated with mosses in waterfalls in the southern Appalachians, USA (Wallace & Ross 1971).

Figure 2. Pseudogoera singularis larva, a species that lives in mosses of waterfalls in the southern Appalachian Mountains. Photo by BIO Photography Group, through Creative Commons.

Chapter 11-12: Aquatic Insects: Holometabola – Trichoptera, Suborders Integripalpia and Spicipalpia

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In the mid-Appalachian Mountain streams, I found two species of Psilotreta (Figure 3) among Hygroamblystegium fluviatile (Glime 1968). This genus has forewings of 6-17 mm (Parker & Wiggins 1987), representing one of the larger of the bryophyte dwellers.

Limnephiloidea

Figure 3. Psilotreta larva, an inhabitant of Hygroamblystegium fluviatile in the Appalachian Mountains. Photo by Bob Henricks, with permission.

Figure 6. Goera pilosa adult, demonstrating the light brown wings typical of the family Goeridae. Photo from Biopix, through Creative Commons.

Figure 4. Hygroamblystegium fluviatile, home of Adicrophleps hitchcockii. Photo by Michael Lüth, with permission.

Figure 7. Pseudogoera singularis larva. Photo by BIO Photography Group, through Creative Commons.

Figure 5. Hygroamblystegium fluviatile leaf showing strong costa that seems to be used in making the cases of Adicrophleps hitchcockii. Photo by Michael Lüth, with permission.

Figure 8. Goera calcarata larva showing large rock fragments on sides of case. Photo by Bob Henricks, with permission.

Goeridae This family occurs on all continents except Australia and South America (Holzenthal et al. 2007). Adults have a forewing length of 6-9 mm and are typically light brown (Figure 6) (Houghton 2012). The larvae (Figure 7) live in cool, flowing water and graze on periphyton. Their larval cases consist entirely of rock fragments, sometimes with larger rocks on each side of the case (Figure 8).

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Chapter 11-12: Aquatic Insects: Holometabola – Trichoptera, Suborders Integripalpia and Spicipalpia

Goerita is a small genus with only three species and is restricted to the Appalachian Mountains and Allegheny Plateau in eastern North America (Parker 1998). The larvae are bryobionts, in this case living on rocks covered with mosses and liverworts where the rocks can be dry or covered by a film of water. The larvae do not eat the bryophytes, but instead feed on detritus and diatoms growing there. Goerita semata lives on the undersides of rocks (Flint 1960), but in western North Carolina, Huryn and Wallace (1985) found the larvae among liverworts and mosses on vertical rock faces; fewer than 2% were found on other substrata. Goerita betteni lives in a similar habitat (Wiggins 1973). Huryn and Wallace (1985) suggested that the bryophytes may offer the larvae some protection from desiccation. Pupae typically occur on these same rocks with mosses and a thin film of water. Ultimately, females lay their eggs away from water on bare rock, mosses, and liverworts. Food of the larvae consists primarily of fine amorphous detritus (65%), and diatoms (32%), but diatom composition increases to an average of 64% in spring. Bryophyte clumps are typically good sources of both. Although the mechanisms of desiccation resistance are unknown in larvae of this species, it is likely that they are adapted behaviorally by living among the bryophytes. In the River Rajcianka in Slovakia, Lithax niger (Figure 9) is a bryophyte dweller, living under water, but not in the wet emergent bryophytes (Krno 1990). This is a mountain species, occurring in the Alps and Balkans.

Figure 10. Sphagnum cuspidatum, a pupation site for Limnephilus peltus and Architremma ulachensis. Photo by Bernd Haynold, through Creative Commons.

Limnephilidae – Northern Caddisflies The Limnephilidae encompasses a wide variety of case-making caddisflies in a wide range of habitats. Their ingenuity in making these homes could challenge some of our most creative artists. This is one of the largest caddisfly families, with recent segregate families diminishing its numbers. Although it occurs worldwide, its records are concentrated in Europe and North America (Limnephilidae 2015). In North America it is often the dominant group in higher elevation streams. But these are mostly large caddisflies (15-35 mm) (Houghton 2012), making navigation difficult among bryophytes. Fontinalis (Figure 11), on the other hand, is a large enough moss with a streamer habit that permits these larger larvae to navigate (Glime 1968, 1994). Their dependence on terrestrial litter makes the larvae vulnerable to deforestation (Houghton 2012).

Figure 9. Lithax niger adult, a species whose larvae live among mosses in the River Rajcianka. Photo by Paul Frandsen, through public domain.

The larvae of Archithremma ulachensis move to a layer of Sphagnum (Figure 10) on the bank of a spring to pupate (Levanidova & Vshivkova 1984). These pupae are morphologically reduced, lacking long setae (hairs) and projections used to clean the silk disks that close the case. They also lack swimming legs. The larvae live in streams that have low water temperatures (3-5°C) in summer. In a cool mountain stream of central Japan Tada and Satake (1994) found that Pseudostenophylax ondakensis (Figure 12) was significantly more abundant on mats of the moss Platyhypnidium riparioides (Figure 13) than in bare rock areas. Décamps (1967, 1968) found Rhadicoleptus spinifer (see Figure 14) to be abundant among mosses in the Pyrénées; at one station it comprised ~15% of the moss Trichoptera fauna (Décamps 1967).

Figure 11. Fontinalis antipyretica, home to many kinds of insects. Photo by Kristian Peters, with permission.

Chapter 11-12: Aquatic Insects: Holometabola – Trichoptera, Suborders Integripalpia and Spicipalpia

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able to shift their diet based on availability, causing misinterpretations based on the general feeding guild classification of these insects. Dangles (2002) considered Chaetopterygopsis maclachlani (Figure 15) to be a specialist on bryophytes; they furthermore build their cases from Fontinalis (Figure 62) (Malicky 1994). As adults they typically crawl, not fly, among the riparian (streambank) vegetation.

Figure 12. Pseudostenophylax ondakensis larva, a species that is significantly more abundant on the moss Platyhypnidium riparioides than on bare rock. Photo by Takao Nozaki, with permission.

Figure 15. Chaetopterygopsis maclachlani adult, a species whose larvae live among Fontinalis and eat mosses as 65% of their diet. Photo from Biodiversity Institute of Ontario, through Creative Commons.

Figure 13. Platyhypnidium riparioides, home to Pseudostenophylax ondakensis in Japan. Photo by J. C. Schou, with permission.

Chaetopterygosis machlachlani is widespread in the Pyrenees to Baikal, specializing in Fontinalis and other streambed mosses (Báilint et al. 2011). In the mid-Appalachian Mountain, USA, streams, the Limnephilidae are poorly represented among bryophytes (Glime 1968). Furthermore, those few that are present differ from any of the species I found in the literature as moss dwellers. Two species of Pycnopsyche [P. luculenta, P. cf. scabripennis (Figure 16)] were the most common, appearing in clumps of Fontinalis (Figure 62) (Glime 1968). This restriction is most likely due to the large size of the Limnephilidae larvae, especially when their bulky case is considered. They would have real difficulty moving about in Hygroamblystegium fluviatile (Figure 4-Figure 5) or Platyhypnidium riparioides (Figure 13).

Figure 14. Rhadicoleptus alpestris adult. Rhadicoleptus spinifer larvae are abundant among mosses in the Pyrénées. Photo by Niels Sloth, with permission.

The larvae of Chaetopterygopsis maclachlani (Figure 15) typically occur among clumps of Fontinalis (Figure 11) in the Vosges Mountains, eastern France, mostly in areas with slower or laminar flow (Lehrian et al. 2010). The mosses constitute ~65% of their diet, with the remainder being coarse leaf detritus (Dangles 2002). Dangles warned that some species, including this one, are

Figure 16. Pycnopsyche scabripennis larva, a Fontinalis dweller. Photo by Tom Murray, through Creative Commons.

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Chapter 11-12: Aquatic Insects: Holometabola – Trichoptera, Suborders Integripalpia and Spicipalpia

In an experimental study on Limnephilus rhombicus (Figure 17), Higler (1975) was able to keep the larvae alive on a diet of Fontinalis antipyretica (Figure 11) with dead birch and oak leaves. However, it appears that its natural diet is mostly living plants (Slack 1936), dead leaves (Slack 1936; Lepneva 1966) and sometimes Naididae (aquatic segmented worms). It is not typically a moss dweller, so the moss diet was most likely unnatural. But Slack (1936) did find that it ate Fontinalis in the field. On the other hand, when Potamophylax rotundipennis (Figure 18Figure 19) was provided choices of birch, oak, and beech leaves and Fontinalis antipyretica, it avoided the moss and beech leaves.

Figure 17. Limnephilus rhombicus larva, showing yet a third very different case, one using snail shells. Photo by Dragiša Savić, with permission.

Figure 19. Potamophylax adult. Photo through Creative Commons.

Figure 20. Two Limnephilus externus larvae with the second grabbing the rear of the first. The two cases appear to be made of bits of grass and this camouflage most likely fools their predators because it confused my non-biologist reviewer! Photo by Wendy Brown , with permission.

Figure 18. Potamophylax larva and case. Potamophylax rotundipennis rejects Fontinalis antipyretica as a food choice. Photo by Michael Wiesner , with permission.

Although most of the Limnephilidae make large cases with large components of twigs and leaf fragments, some use bryophytes. Limnephilus externus (Figure 20-Figure 21) larvae are known to use the moss Leptodictyum riparium (Figure 22) to construct their barrel-shaped cases (Pritchard & Berté 1987). In experiments, this species was able to use wheat flakes, but not alder leaves, to make its case. In the same experiment, Nemotaulius hostilis (Figure 23) used alder, willow, and burreed but did not use wheat flakes or mosses. These same two insects are shredders that consume tracheophyte detritus, but the proportion of mosses in the diet increases as the larvae become older.

Figure 21. Limnephilus externus larva. Photo by Wendy Brown , with permission.

Limnephilus peltus (Figure 24) doesn't spend much time among mosses as a larva, but when it is time to pupate, it burrows into mosses along fen streams where it spends its pupal life (Erman 1984). Unfortunately, if the stream dries out, the pupa is likely to die.

Chapter 11-12: Aquatic Insects: Holometabola – Trichoptera, Suborders Integripalpia and Spicipalpia

Figure 22. Leptodictyum riparium, home of larvae of Limnephilus externus. Photo by Jan-Peter Frahm, with permission.

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Figure 25. Drusus discolor adult, a species that lives among the moss Bryum in the Pyrénées. Photo from Biodiversity Institute of Ontario, through Creative Commons.

Figure 23. Nemotaulius hostilis larva showing case made of leaf litter. Photo by Donald S. Chandler, with permission. Figure 26. Bryum pseudotriquetrum, home to several species of Drusus in Europe. Photo by Hermann Schachner, through Creative Commons.

Figure 24. Limnephilus sp. larva, a genus that sometimes pupates in mosses of fens. Photo by Jason Neuswanger, with permission.

The habitat of larvae of the high altitude Drusus discolor (Figure 25) in the Pyrénées consisted of filamentous algae and the moss Bryum (Figure 26) (Décamps 1968). This caddisfly is one of the two most abundant caddisflies among mosses (Décamps 1967). In the River Rajcianka in Slovakia, Drusus annulatus (Figure 27) occurs not only among submerged bryophytes but also moving about among the wet bryophytes that emerge above the water level (Krno 1990).

Figure 27. Drusus annulatus adult, a species whose larvae can live above or below the water surface among bryophytes. Photo by James K. Lindsey, with permission.

Frenesia difficilis (Figure 28) lays its eggs out of the water, sometimes on mosses that overhang the water (Flint 1956). In this terrestrial location the eggs may freeze in winter. In the Massachusetts, USA, fish hatchery, Flint found no other relationship with mosses during the life cycle.

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Chapter 11-12: Aquatic Insects: Holometabola – Trichoptera, Suborders Integripalpia and Spicipalpia

Figure 28. Fresnia difficilis male, a species that sometimes lays its eggs on mosses that overhang the water. Photo by Tom Murray, through Creative Commons.

The Arctic caddisfly Sphagnophylax meiops lives in Arctic pools in the tundra in the Northwest Territories of Canada (Wiggins & Winchester 1984) where the larvae take advantage of the surface water in the pool (Winchester et al. 1993). When the water recedes the larvae move to the organic materials accumulated above the permafrost to feed, grow, and metamorphose into pupae and adults. This caddisfly is flightless and has long bristles on its short wings. Most Trichoptera spend their larval life in the water, but in the genus Enoicyla (Limnephilidae; Figure 29), the larvae are terrestrial and the adult female has only vestigial wings, limiting her travel and agility. Males, however, are capable fliers. Larvae may live far from water among the mosses around tree roots (Watson & Dallwitz 2003). Green (2012) noted at least 50 of these larvae climbing up logs, with several browsing a black slime mold. One can observe many larvae together on the surface of mosses and liverworts growing on a stream bank following rain. Enoicyla pusilla (Figure 29) uses fine sand grains and other vegetable matter to make cases where it lives among the mosses (Watson & Dallwitz 2003). The larvae of Enoicyla, despite being terrestrial, require 100% humidity (Green 2012). But when they become saturated, they climb upwards to dry, then drop back down when they need to get wet again (at 7% relative humidity). Their respiration is through the cuticle; they lack gills.

In his arguments to support that the Trichoptera (with hairs on wings) and Lepidoptera (with scales on wings) were closely related, Crampton (1920) used the common ability to use mosses in the caddisfly Enoicyla (Limnephilidae; Figure 29) and the larvae of moths in Micropterygidae. The caddisflies living in peatlands are typically generalist taxa with wide habitat requirements (Flannagan & Macdonald 1987). But a few are tyrphobionts (living only in peat bogs and mires). The larvae of Phanocelia canadensis (Figure 30-Figure 31) are elusive. The second report of the larvae by Colburn and Clapp in 2006 was from kettle hole wetlands in Massachusetts, USA. Colburn and Clapp attribute the limited reports of larvae of this species to its limited habitat requirements. It lives in Sphagnum (Figure 10) habitats with low pH and makes its case from Sphagnum (Figure 30) [The picture below (Figure 31) indicates other mosses are used as well.] Larvae remain closely associated with the moss during development. They become dormant in summer, remaining in unsealed cases that are firmly attached to the moss. In autumn they seal the ends of the case and develop into pupae. Even fossil records support their preference for Sphagnum (Figure 10) bogs. The larva was originally described from floating Sphagnum at the edge of acidic ponds in a spruce-Sphagnum bog in New Brunswick, Canada (Fairchild & Wiggins 1989). It appears that adult habitats are much broader, perhaps misleading its collectors (Colburn & Clapp 2006).

Figure 30. Phanocelia canadensis larva showing its case made with Sphagnum. Photo from Biodiversity Institute of Ontario, through Creative Commons.

Figure 29. Enoicyla pusilla larvae, a terrestrial species that requires 100% humidity – a condition often found among mosses. Photo by Ernest van Asseldonk, through Creative Commons.

Figure 31. Phanocelia canadensis larva showing case made with at least some non-Sphagnum mosses. Photo from Biodiversity Institute of Ontario, through Creative Commons.

Chapter 11-12: Aquatic Insects: Holometabola – Trichoptera, Suborders Integripalpia and Spicipalpia

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Leberfinger and Bohman (2010) gave larvae of Limnephilus bipunctatus (Figure 32) choices of food that included grasses, mosses, algae, and leaves. The larvae preferred leaves of the shrubby cinquefoil. Although they ate little of the mosses, grass was the least preferred food.

Figure 34. Philocasca alba adult, a species whose larvae feed on the moss Hygrohypnum luridum in a Rocky Mountain, USA, stream in spring and summer. Photo from Biodiversity Institute of Ontario, through Creative Commons.

Figure 32. Limnephilus bipunctatus larva in case, a species that includes mosses in its diet. Photo by James K. Lindsey, with permission.

Philocasca is not a genus one often reads about in moss habitats. Nevertheless, mosses appear to be suitable sites for pupation. In describing the new species Philocasca rivularis (see Figure 33) Wiggins and Anderson (1968) state that pupae attach to the undersides of moss clumps along stream banks. Mutch and Pritchard (1984) found that instar V larvae of P. alba (Figure 34) in a Rocky Mountain stream had mostly moss (Hygrohypnum luridum – Figure 35) in the gut in spring and summer, but had leaf fragments in the gut in autumn. Furthermore, when fed detritus supplemented with moss these larvae grew significantly better than when fed detritus alone, suggesting that the moss was an important nutrient source.

Figure 35. Hygrohypnum luridum, a species that typically occurs both in the water and above it. Photo by Dale Vitt, with permission.

Onocosmoecus unicolor (Figure 36-Figure 37) is a large shredder that includes mosses in its varied diet (National Park Service 2014).

Figure 36. Onocosmoecus unicolor larva, a moss consumer. Photo by Jason Neuswanger, with permission.

Figure 33. Philocasca thor adult. Philocasca rivularis pupates on undersides of moss clumps on streambanks. Photo from Biodiversity Institute of Ontario, through Creative Commons.

Figure 37. Onocosmoecus unicolor adult. Photo by Bob Newell, with permission.

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Chapter 11-12: Aquatic Insects: Holometabola – Trichoptera, Suborders Integripalpia and Spicipalpia

Chyranda centralis (Figure 38) is a caddisfly of small spring streams among leaf accumulations. Its food includes leaves, bark, and may even include mosses (National Park Service 2014).

Figure 40. Allogamus auricollis larvae. Photo by Wolfram Graf, with permission.

Figure 38 Chyranda larva of small spring streams; it may sometimes eat mosses. Photo from California Department of Fish and Wildlife, through public domain.

Mosses provide vertical zonation possibilities for the caddisflies. Krno (1990) addressed these vertical zones in the River Rajcianka in Slovakia. There, the limnephilids Allogamus auricollis (Figure 39-Figure 40) (a shredder), A. uncatus, and Drusus annulatus (Figure 41) occurred among the submerged mosses, but above water only Allogamus auricollis and Drusus annulatus occurred among emergent wet mosses. On the other hand Parachiona picicornis (Figure 42) was only found above water among the wet mosses.

Figure 41. Drusus annulatus adult, a species whose larvae live among submerged mosses and will venture above the water among wet mosses. Photo by James K. Lindsey, with permission.

Figure 42. Parachiona picicornis adult, a species whose larvae live among submerged mosses but will not venture above the water among wet mosses. Photo by James K. Lindsey, with permission.

Chaetopterygopsis maclachlani larvae in the Carpathians are "specialized" on the aquatic moss Fontinalis (Figure 62) in mountain streams (Bálint et al. 2011).

Figure 39. Allogamus auricollis larva, a species that traverses among mosses both below and above the water surface. The larva is seen here breaking the surface tension. Photo through Creative Commons.

Figure 43. Chaetopterygopsis machlachlani larva, a Fontinalis dweller. Photo by Michael Balke, through Creative Commons.

Chapter 11-12: Aquatic Insects: Holometabola – Trichoptera, Suborders Integripalpia and Spicipalpia

Lepidostomatidae – Bizarre Caddisflies This family is widespread in the Northern Hemisphere, extending southward to Panama, New Guinea, and the Afrotropical region (Holzenthal et al. 2010a). Hilsenhoff (1975), in reporting on Wisconsin, USA, Lepidostomatidae, considered the larvae of this family to inhabit a wide range of clean streams. The larvae live among rocks, debris, and mosses on rocks and eat mostly detritus (BugGuide 2005). In North America the larvae inhabit springs, streams, and large slow-moving rivers where they eat detritus. They build a log cabin style of case from stem and leaf pieces or sand grains. I did find Lepidostoma americana in clumps of Hygroamblystegium fluviatile (Figure 4-Figure 5) in the Appalachian Mountain streams (Glime 1968). Some older cases of Lepidostoma sp. contained fragments of the liverwort Scapania undulata (Figure 74) in them near the opening. Lepidostoma hirtum (Figure 44-Figure 45) is common among mosses at both Ballysmuttan and Straffan in the UK (Frost 1942). Its diet consists of algae, mosses, and tracheophytes (Rousseau et al. (1921). The moss not only provides a suitable location to find its food, but provides it protection from trout and other fish that are its predators.

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rock material, and feed on plant debris (Holzenthal et al. 2007).

Figure 46. Crunoecia irrorata larva, a moss consumer. Photo by Niels Sloth, with permission.

In New Zealand, both Oeconesus maori (see Figure 47) and Zelandopsyche ingens (Figure 48) occasionally ingest bryophytes (Suren 1988). Suren and Winterbourn (1991) determined that of the 14 taxa that had bryophyte fragments in their guts, only Zelandopsyche ingens and Oeconesus similis consumed them regularly.

Figure 44. Lepidostoma hirtum larva, an inhabitant of bryophytes that also eats them. Photo by Urmas Kruus, with permission.

Figure 47. Oeconesus larva, a bryophyte dweller and bryophyte consumer in New Zealand. Photo by Stephen Moore, Landcare Research, NZ, with permission.

Figure 45. Lepidostoma hirtum larva head. Photo by Urmas Kruus, with permission.

Crunoecia irrorata (Figure 46) prefers moss cushions and fallen leaves (Köcherfliegen 2015). In UK streams, this species had mosses in the gut (Percival & Whitehead 1929). Oeconesidae This is a small family from Tasmania (1 species) and New Zealand (Holzenthal et al. 2007), but of a relatively large size (adults 30-38 mm) (Oeconesidae 2013). Larvae live in small, forested streams, make cases from plant and

Figure 48. Zelandopsyche larva and case, a bryophyte dweller and regular bryophyte consumer. Photo by Stephen Moore, Landcare Research, NZ, with permission.

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Chapter 11-12: Aquatic Insects: Holometabola – Trichoptera, Suborders Integripalpia and Spicipalpia

Uenoidae This family lives mostly in cool, fast-flowing headwaters and is distributed in North America, southern Europe, and eastern Asia (Holzenthal et al. 2007). Their cases may be constructed either of coarse pebbles, as in Neophylax (Figure 53-Figure 55), or of fine sand, flattened, and shaped like the shell of a limpet, as in Thremma (Figure 49). Larvae eat diatoms and fine particulate matter that they scrape from rocks. These larvae are among the smaller caddisflies, being up to 15 mm (Wiggins 2004), although for moss dwellers they would be in the medium to large category.

Figure 51. Neothremma alicia larva with case, a moss dweller in small, headwater streams. Photo from Biodiversity Institute of Ontario, through Creative Commons.

Figure 52. Neothremma alicia larva outside its case. Photo from Biodiversity Institute of Ontario, through Creative Commons.

Figure 49. Thremma gallicum larva showing limpet type of case. Photo from Guillaume Doucet, with permission.

Thremma sp. (Figure 49) in the trout streams of Yellowstone National Park, USA, occurs among mosses and the alga Cladophora in strong rapids (Muttkowski & Smith 1929). Each of these caddisflies collected from the mosses had mosses in the gut, averaging 70% of the contents. The alga Epithemia (Figure 50), most likely living among the mosses, comprised the remaining 30%. Brown (2007) found significant numbers of Neothremma alicia (Figure 51-Figure 52) in small, mossy streams in the headwaters of the East River, Colorado, USA.

Figure 50. Epithemia, a diatom genus that is a common food source for the caddisfly Thremma. Photo by Kristian Peters, with permission.

In the Appalachian Mountain stream bryophytes, the Uenoidae were represented by a completely different genus from the ones I found in publications, the only one being Neophylax, a genus that sometimes reached large numbers among the Trichoptera, but usually was absent (Glime 1968). Nevertheless, three species were represented: N. concinnus (Figure 53), N. consimilis (Figure 54), N. oligius (Figure 55). These were usually in the mat-forming bryophytes, a location permitted by their smaller size.

Figure 53. Neophylax concinnus larva, a moss dweller in mid-Appalachian Mountain streams. Photo by Bob Henricks, with permission.

Chapter 11-12: Aquatic Insects: Holometabola – Trichoptera, Suborders Integripalpia and Spicipalpia

Figure 54. Neophylax consimilis larva, a moss dweller in mid-Appalachian Mountain streams. Photo by Bob Henricks, with permission.

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Figure 56. Brachycentrus numerosus larva, a species like one that is common among Hygroamblystegium fluviatile in the Appalachian Mountain streams. Photo by Tom Murray, through Creative Commons.

Figure 55. Neophylax oligius larva, a moss dweller in midAppalachian Mountain streams. Photo by Tom Murray, through Creative Commons. Figure 57. Brachycentrus montanus adult, a species that lives among stream mosses. Photo by James K. Lindsey, with permission.

Phryganeoidea Brachycentridae Caddisflies

–

Humpless

Casemaker

The Brachycentridae are a Northern Hemisphere family (Holzenthal et al. 2010b). They eat algae and plankton (Neuswanger 2015b), but some also ingest bryophytes (Muttkowski & Smith 1929). These caddisflies build cases that resemble log cabins or cylinders made of tiny plant fragments (Holzenthal et al. 2010b), including bryophytes in some genera (Glime 1968). Often they are found among mosses (Bouchard 2004). When they emerge, they do so on the surface, which sometimes subjects them to 3-7 m of drifting (Neuswanger 2015b). Females may dive to lay eggs or land with spread wings on the surface to accomplish the task. Brachycentrus Larvae of Brachycentrus (Figure 56-Figure 59) species actually attach to the mosses (Armitage 1961; Glime 1968). Brachycentrus was one of only two genera of caddisflies that Muttkowski and Smith (1929) found among mosses in the trout streams of Yellowstone National Park, USA. Needham and Christenson (1927) reported Brachycentrus from mosses in streams of northern Utah, USA. In Europe, Krno (1990) found Brachycentrus montanus (Figure 56) among mosses in the River Rajcianka, Slavakia. In the Appalachian Mountains, B. cf. numerosus (Figure 56) occurred in clumps of the moss Hygroamblystegium fluviatile (Figure 4-Figure 5) (Glime 1968).

Gallepp (1977) considered Brachycentrus – B. americanus (Figure 58), B. occidentalis (Figure 59) – to be filter feeders, but Muttkowski and Smith (1929) found that mosses were among the food items in the gut, with one individual having 90% moss. Others had only algae and a few had aquatic insects.

Figure 58. Brachycentrus americanus larva, a moss consumer. Photo by Donald S. Chandler, with permission.

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Chapter 11-12: Aquatic Insects: Holometabola – Trichoptera, Suborders Integripalpia and Spicipalpia

Figure 59. Brachycentrus occidentalis larvae, a moss consumer species. Photo by Arlen Thomason, with permission.

riparioides (Figure 13) was the most frequent food, but both mosses and liverworts were eaten. Diatoms were also present in the gut, but they might have been eaten inadvertently along with the bryophytes. And in the Pyrénées Micrasema morosum behaves as a shredder and eats mosses (and periphyton) as well (Décamps & Lafont 1974). In the Pyrénées Décamps (1968) found that Micrasema morosum was abundant in the mosses Cratoneuron commutatum (Figure 61) and Bryum (Figure 26) and was the most abundant bryophyte-inhabiting caddisfly. At one station M. morosum comprised 56% of the Trichoptera fauna among mosses and at another it comprised 87.8% (Décamps 1967). Micrasema vestitum was abundant in Fontinalis squamosa (Figure 62) and in one location it comprised 69% of the Trichoptera fauna among the mosses.

Gallepp (1977) found that two species of Brachycentrus were more responsive to temperature and food availability than to the flow rate. Although casebuilding decreased with increasing temperature over the range of 4-17°C, B. occidentalis (Figure 59) grew faster as the temperature increased in the range of 4-27°C. Micrasema The larvae of the grazer genus Micrasema (Figure 60) (Gallepp 1977) are common among mosses (Glime 1968, 1994; Tada & Satake 1994). In the mid-Appalachian Mountain streams I was able to distinguish three different morphotypes (species?) among the bryophytes (Glime 1968). In fact, this genus seems to be almost restricted to that habitat (Hilsenhoff 1975). Tada and Satake (1994) found a species in this genus to be the most abundant insect taxon on mats of Platyhypnidium riparioides (Figure 13) in a cool mountain stream in central Japan. Among the bryophyte mats its density exceeded 100,000 individuals per square meter in November, an abundance that was 2.816.3 times as high as that on the bare rock bottom. At least one species of Micrasema (Figure 60) constructs a "log cabin" out of moss stems and leaves (Glime 1968).

Figure 61. Cratoneuron commutatum, home to several species of Micrasema. Photo through Creative Commons.

Figure 62. Fontinalis squamosa, home to several species of Micrasema larvae. Photo by David T. Holyoak, with permission. Figure 60. Micrasema charonis larva, a common mossdweller that often makes its case from mosses. Photo by Bob Henricks, with permission.

Chapman and Demory (1963) found that in two streams in Oregon, USA, this genus occurred only among mosses and liverworts where there was little detritus. They graze on periphytic algae during the first instar, but in later instars they are likely to be herbivore-chewers (shredders) on mosses and other small photosynthetic material (Chapman & Demory 1963; Aquatic Insects). In fact, Chapman and Demory (1963) found that Platyhypnidium

Décamps and Lafont (1974) demonstrated the change in moss substrate for Micrasema morosum as altitude changes in the Pyrénées. At 1940 m asl the dominant bryophytes were Brachythecium rivulare (Figure 63), Cratoneuron commutatum (Figure 61), and Hygrohypnum molle (Figure 64). At 1590 m asl dominance shifted to Fontinalis squamosa (Figure 62), Fissidens polyphyllus (Figure 65), and Platyhypnidium riparioides (Figure 13). At 1360 m asl Fissidens grandifrons (Figure 66) appeared and Platyhypnidium riparioides (Figure 13) remained in the stream flora. At

Chapter 11-12: Aquatic Insects: Holometabola – Trichoptera, Suborders Integripalpia and Spicipalpia

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550 m asl the dominant mosses were Brachythecium rivulare, Fissidens grandifrons, Platyhypnidium riparioides, and Chiloscyphus polyanthos (Figure 67), with a change in the Micrasema species to M. morosum, M. longulum, M. moestum, M. difficile, and M. minimum. At the lowest location of 430 m, asl Brachythecium rivulare, Cinclidotus fontinaloides (Figure 68), Fontinalis antipyretica (Figure 11), Platyhypnidium riparioides, and Cratoneuron filicinum (Figure 69) with Micrasema morosum once again the predominant species. The food of these Micrasema species consisted of fragments of mosses and periphytic algae, with some food unidentifiable. Figure 66. Fissidens grandifrons, home to larvae of several Micrasema species. Photo by Scot Loring, through Creative Commons.

Figure 63. Brachythecium rivulare, home to several species of Micrasema larvae. Photo by David T. Holyoak, with permission.

Figure 64. Hygrohypnum molle, home to several species of Micrasema larvae. Photo by Jan-Peter Frahm, with permission.

Figure 65. Fissidens polyphyllus, home for several species of Micrasema. Photo by David T. Holyoak, with permission.

Figure 67. Chiloscyphus polyanthos, home to lower elevation species of Micrasema larvae in the Pyrénées. Photo by Barry Stewart., with permission

Figure 68. Cinclidotus fontinaloides, home to lower elevation species of Micrasema larvae in the Pyrénées. Photo by David T. Holyoak, with permission.

Figure 69. Cratoneuron filicinum in Europe, home for many immature insects. Photo by Michael Lüth, with permission.

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In Japan, Micrasema uenoi (Figure 70) feeds on the leaves of Platyhypnidium riparioides (Figure 13) and the first instar larvae make their cases of its leaves (Kato 1995). The first two instars live in greater numbers among mosses than on cobble, but by third to fifth instars the numbers are about equal. When artificial mosses (glass wool) and cleaned mosses were introduced, these larvae reached normal densities in 15-30 days. Surprisingly, the density on the glass wool was 2-3 times that among the mosses, but it subsequently decreased quickly. Gut contents of those third to fifth instars on bryophytes was 80% moss; those on the glass wool contained litter and detritus instead. The larvae move about a lot between the pebbles and the mosses. Eggs were apparently absent on the mosses, suggesting that the hatchlings move there.

Figure 72. Micrasema wataga case with moss sprouts. A pupa is hiding inside. Photo by D. N. Bennett, with permission. Figure 70. Micrasema uenoi adult, a species whose larvae feed on leaves of Platyhypnidium riparioides in Japan. Photo by Takao Nozaki, with permission.

D. N. Bennett (pers. comm. 6 August 2013, 12 August 2014) observed Micrasema wataga (Figure 71-Figure 72) larvae eating moss (possibly Hygrohypnum montanum) leaves (Figure 71) in the Blue Ridge Mountains of Virginia, USA. They made their cases of the same moss, starting with a tiny cone of minute sand grains. The mosses closest to this cone part, hence the oldest, were no longer green, but those near the opening were still green. This can be a possible source of dispersal of fragments that break away from the unfinished cases. But a later observation showed that the mosses in the case actually sprouted there (Figure 72)! This case was apparently occupied by a pupa, ceasing the activity that could break off these sprouts before they attained sufficient size to exist on their own.

Figure 71. Micrasema wataga eating moss (Hygrohypnum montanum?). Photo by D. N. Bennett, with permission.

Adicrophleps hitchcockii This interesting larva makes its case from bryophytes. It was relatively common among Hygroamblystegium fluviatile (Figure 4-Figure 5) in Appalachian Mountain streams (Glime 1968). It appeared to have used costae from this moss in the construction of its cases. D. N. Bennett likewise collected larvae of the somewhat rare Adicrophleps hitchcockii (Figure 1, Figure 73) in several cold, rapid streams (1-10 m wide) from the aquatic leafy liverwort Scapania (Figure 74) growing in riffle areas (Henricks 2013; D. N. Bennett, pers. comm. September 2014). But the case is not made of liverworts, but rather it displays mosses. Wiggins (1977) described these as "4-sided, tapered, and constructed of pieces of moss arranged transversely; trailing ends frequently left attached to the moss pieces give the case a furry appearance."

Figure 73. Adicrophleps hitchcocki, a species that lives among bryophytes and makes its case from mosses. Photo by D. N. Bennett, with permission.

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Figure 76. Trichoptera eggs, often laid on bryophytes. Photo by Bob Armstrong, with permission.

Figure 74. Scapania undulata, home for Adicrophleps hitchcocki but not used for case building. Photo by Hermann Schachner, through Creative Commons.

The larvae of Eubasilissa regina (Figure 77) in Japan begin their construction days by making cases of liverworts, but as they develop they change to terrestrial leaf litter and move their abode from the liverworts to pools (Ito 1988).

Phryganeidae – Giant Casemakers This family with relatively large larvae lives mostly in lakes and rivers (Neuswanger 2015a). The pupae crawl from their watery location to shore to emerge. Females run across the water surface to lay their eggs. The larvae are most common among aquatic plants in ponds and marshes, but some occur in streams and others in temporary pools and deep in lakes (Holzenthal et al. 2007). Larvae are typically either predators or herbivores. This family is not common among the bryophytes. But, Yphria californica (Figure 75), a species restricted to the west coast states of USA, lays its eggs (Figure 76) underwater among mosses that dangle over the stream in the Sierra Nevada, North America (Erman 1984). To do that, the adult must swim underwater.

Figure 75. Yphria californica adult, a USA west coast species that lays its eggs among mosses. Biodiversity Institute of Ontario, through Creative Commons.

Figure 77. Eubasilissa regina adult, a large Japanese caddisfly for which the larvae begin their case construction using liverworts. Photo through Creative Commons.

Oligostomis ocelligera (Figure 78) lives in moist places such as under mosses where it is protected (Redell et al. 2009). It usually occupies positions with a mean distance of 6.1 cm below the surface.

Figure 78. Oligostomis ocelligera larva, a species that lives under mosses. Photo by Tom Murray, through Creative Commons.

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Chapter 11-12: Aquatic Insects: Holometabola – Trichoptera, Suborders Integripalpia and Spicipalpia

Hagenella clathrata is a rare caddisfly in Europe, inhabiting the disappearing bog habitat (Buczyńska et al. 2012). In particular, the species often occurs in bog pools that occur only in rapidly disappearing floating bogs, hence being dependent on the particular habitat created by Sphagnum (Figure 10) (Kleef et al. 2012).

Sericostomatoidea Beraeidae This family is scattered about the globe, being concentrated in the western Palaearctic Region (Eurasia from western Europe to the Bering Sea), but also occurs in Tanzania, Japan, and eastern North America (Hamilton 1985; Holzenthal et al. 2007). Adults have forewings that are only 4-6 mm long (Watson & Dallwitz 2003). Larvae live in springs, seeps, and small streams where they utilize a variety of substrates, including bryophytes (Hamilton 1985; Holzenthal et al. 2007). They eat plant and fungal material, but there seem to be no records of eating bryophytes. Beraea maura (Figure 79) represents this family in the River Rajcianka, Slovakia, where it inhabits the submerged bryophytes (Krno 1990). Unlike several members of the Limnephilidae and Rhyacophilidae, this species is not found above the water level in the wet mosses there. In the Pyrénées, Décamps (1968) found larvae of this family among mosses, but this family had a wide range of habitats in addition to the mosses.

Figure 80. Pycnocentrodes aureolus adult, member of a family (Conoesucidae) with bryophyte dwellers in the Australian region. Photo by Maurice, through Creative Commons.

Helicophidae This family of 6-14 mm length (Helicophidae 2015b) is mostly known from Australia, New Zealand, and New Caledonia, but also from southern South America and scattered locations in North America (Helicophidae 2015a). The larvae live in slow streams and are mostly detritivores (Helicophidae 2015b). Trichoptera are not as common in New Zealand as in other parts of the planet, but the Helicophidae are represented there, sometimes associated with mosses (Winterbourn & Gregson 1981). Zelolessica cheira (Figure 81) occurs among Fissidens rigidulus (Figure 82) in the torrential waters near the middle of stream channels in the Southern Alps (Cowie & Winterbourn 1979). Zelolessica cheira is usually associated with mosses and liverworts in rapid streams with a stable, rocky substrate (Winterbourn & Gregson 1981; Eward et al. 1994). The cases are curved, comprised variously of sand grains, liverworts, and mosses.

Figure 79. Beraea maura adult, a species that lives among submerged bryophytes as larvae. Photo from Biodiversity Institute of Ontario, through Creative Commons.

Conoesucidae Among the unfamiliar Trichoptera names (to those of us in the northern hemisphere), the Conoesucidae (Figure 80) is another of bryophyte-dwelling families from down under (Winterbourn & Gregson 1981). The family is endemic to Australia, New Zealand, and Tasmania (Johanson et al. 2009). Among the bryophyte dwellers is Confluens hamiltoni, an endemic on the North Island, New Zealand, where it is associated with mosses, liverworts, and algae in rapid-flow streams (Winterbourn & Gregson 1981). On the South Island, this species is replaced by C. olingoides, occupying conditions like those of C. hamiltoni.

Figure 81. Zelolessica larvae. Some members make their cases from bryophytes. Photo by Stephen Moore, Landcare Research, NZ, with permission.

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Figure 84. Gumaga sp. larva, a relatively immobile caddisfly. Photo from Biodiversity Institute of Ontario, through Creative Commons.

Figure 82. Fissidens rigidulus, home for Zelolessica cheira in torrential New Zealand waters. Photo by Bill & Nancy Malcolm, with permission.

Alloecentrella (Figure 83) is known from China, Australia, New Zealand, and the Antarctic. In New Zealand, Alloecentrella magnicornis and an unnamed species occur among mosses and liverworts in rocky streams where they build their cases using bryophytes (Eward et al. 1994).

Figure 85. Gumaga nigricula adult, a relatively immobile caddisfly in the larval stage. Photo from Biodiversity Institute of Ontario, through Creative Commons.

Figure 83. Alloecentrella sp. larva, a species that covers its case with mosses and liverworts. Photo by Stephen Moore, Landcare Research, NZ, with permission.

The Sericostomatidae live in both streams and lakes and mostly feed on leaf litter (Family Sericostomatidae 2015). They build slightly to strongly curved tubular cases from sand grains or just silk. Because of their interesting designs and strength, the Tupi-Guarani Indians in Brazil used the cases of Grumicha as adornment. Some of the moss dwellers are quite rare. Stern and Stern (1969) found the larvae of Sericostoma sp. (Figure 86) only among algae and mosses in a Tennessee, USA, springbrook. Sericostoma pedemontanum (Figure 86), a caddisfly of fast-running streams, refused Fontinalis antipyretica (Figure 11) when provided a diet of birch, beech, and oak leaves with it (Higler 1975). Birch was the preferred food.

Sericostomatidae – Bushtailed Caddisflies These caddisflies are of moderate size, with wings 815 mm long (Watson & Dallwitz 2011). This family is cosmopolitan except for the Australian region (Sericostomatidae 2015). Nevertheless, many of the genera are endemic to small areas of their continents. At least some larval members of the family move little. For example, more than 120,000 larvae of Gumaga nigricula (Figure 84-Figure 85) were released in pools of a California mountain stream and 87-93% of them remained within 4 m of the pools (Jackson et al. 1999). In this clever experiment, the larvae were provided with bright gold or magenta sand grains to complete their cases so that they could easily be tracked.

Figure 86. Sericostoma pedemontanum larva, a species that refused Fontinalis and chose various species of leaf litter in a feeding experiment. Photo by Massimo Del Guasta, with permission.

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Chapter 11-12: Aquatic Insects: Holometabola – Trichoptera, Suborders Integripalpia and Spicipalpia

SUBORDER SPICIPALPIA Glossosomatoidea Glossosomatidae Makers

–

Tortoise

or

Saddle-case

This worldwide family makes its larval cases from pebbles in the shape of a turtle shell (Glossosomatidae 2014). It is probably this structure that forces them to build a new case in each new instar, rather than adding to the old one as most caddisfly families do. These small to mediumsized larvae usually occur in cool mountain streams where they scrape algae from the rocks as their food. The female adults lay their eggs in gelatinous masses under rocks at the water surface or on floating objects, probably including mosses. The gelatinous material protects the eggs from desiccation. From Ceylon, Schmid (1958) reported Agapetus rawana (see Figure 87-Figure 90) from large, mossy rocks in the torrent. In the Appalachian Mountains, Glossosoma (Figure 91) larvae and pupae were often present among the bryophytes (Glime 1968).

Figure 89. Agapetus prepupa in larval case. Photo by Mark Melton, with permission.

Figure 90. Agapetus pupa removed from case. Photo by Mark Melton, with permission.

Figure 87. Agapetus fuscipes larva and case, a genus known from large, mossy rocks of torrents in Ceylon. Photo by J. C. Schou, with permission.

Figure 91. Glossosoma sp. larvae, showing its "turtle shell" case. Photo by Jason Neuswanger, with permission.

Hydroptiloidea Hydroptilidae Caddisflies

Figure 88. Agapetus fuscipes larvae showing the unusual shape of the case. Photo by Dragiša Savić, with permission.

–

Microcaddisflies,

Purse-case

This is a worldwide family, less than 5 mm long, that builds flattened cases often resembling an eyeglass case (Hydroptilidae 2015). The members of the family solve the problem of locating food by depositing their eggs near a suitable food source (Leader 1970). They typically feed on algae by sucking out the cell contents or by feeding on diatoms.

Chapter 11-12: Aquatic Insects: Holometabola – Trichoptera, Suborders Integripalpia and Spicipalpia

In the Appalachian Mountain streams where I worked, this tiny caddisfly is usually not very common, but Percival and Whitehead (1929) found them more commonly among mosses on stones than on other substrates in the UK. Hughes (1966) found them to be more abundant in open areas than in shaded ones, a factor that usually contrasts with bryophyte preferences. Percival and Whitehead (1929) found that the hydroptilids from mosses feed on algae and diatoms. The larvae of this family have mouthparts that are able to pierce and suck, enabling them to suck the contents from filamentous algae or to scoop up diatoms (Nielsen 1948). It is perhaps telling that at least in Denmark, the genera Agraylea (Figure 92), Hydroptila (Figure 93), Oxyethira (Figure 94-Figure 95), and Orthotrichia (Figure 96) are very common in eutrophic lakes (Nielsen 1948). This suggests that in streams we should look for the bryophyte dwellers deep within the mat where there is reduced flow. But even in the lakes these genera occupy vegetation near the surface. Agraylea and Orthotrichia occur in slowly flowing water, and this is where mosses can add possible niches. Orthotrichia often becomes coated in detritus and will pass one of its hind legs down the dorsal side of its abdomen to clean the tracheal gills there.

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Figure 94. Oxyethira larva, a moss dweller in Danish lakes. Photo by Stephen Moore, Landcare Research, NZ, with permission.

Figure 95. Oxyethira pupa. Photo by Stephen Moore, Landcare Research, NZ, with permission.

Figure 92. Agraylea sexmaculata larva, a genus that lives among bryophytes in slowly flowing water. Photo by Massino Del Guasta, with permission.

Figure 96. Orthotrichia sp larva and case, a species that lives among mosses in lakes. Photo by Urmas Kruus, with permission.

Figure 93. Hydroptila sparsa larvae, member of a genus that occurs among bryophytes in lakes and streams. Photo by Massimo Del Guasta, with permission.

Hydroptila (Figure 93) can build a case of detrital matter and sand grains in about four hours (Nielsen 1948). To increase the size of the case, the larva splits it open along the ventral edge, adding sand grains to the edge. The completed case, as in most members of the family, looks like a case for eye glasses (Figure 93) – the one with an

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Chapter 11-12: Aquatic Insects: Holometabola – Trichoptera, Suborders Integripalpia and Spicipalpia

open end – which is where the head protrudes in the caddisfly version. Some cases are built with algal filaments, especially in Agraylea (Figure 92), and I have observed cases made almost entirely of diatoms. In both Hydroptila and Agraylea the outer coating of sand or algae will wear off as the larva nears maturity, leaving only the smooth inner wall made of silk spun by the larva as it cements the case together. Orthotrichia (Figure 96) and Ithytrichia (Figure 97) species use only silk in the construction of their cases. These genera feed by sucking the contents out of algal cells.

Figure 98. Fontinalis dalecarlica, home to many insects. Photo by J. C. Schou, with permission.

Woodall and Wallace (1972) found Ochrotrichia sp (Figure 99) on moss-covered granite outcrops in the Appalachian, USA, streams that they studied. They considered the moss-covered rock outcrops to be the central factor influencing the distribution of this species in the area. In my own studies of the mid-Appalachian Mountain streams, this genus was not present, but I did occasionally find Mayatrichia, Neotrichia, and Stactobiella in addition to the more common ones discussed above under this family (Glime 1968).

Figure 97. Ithytrichia lamellaris larva & case, a genus that uses only silk in its case. Photo by Urmas Kruus, with permission.

When these four genera (Agraylea, Hydroptila, Orthotrichia, Ithytrichia) emerge, they split the pupal case, then move about until they find a protruding object to climb up and out of the water (Nielsen 1948). Once out they can flit about on the water surface and in the air. The moss-dwelling genus Oxyethira (Figure 94-Figure 95), including more than one species, comprised 44.5% of the Trichoptera fauna at the acid site in Frost's (1942) moss fauna study of the River Liffey, Ireland. It was absent at the alkaline site. Oxyethira frici lives in the angle between the leaf and the stem of the moss and pupates among the mosses, a behavior that is uncommon among caddisflies. By contrast, Ithytrichia lamellaris (Figure 97), a species almost restricted to mosses, was common at the alkaline site and absent from the acid site. It likewise lives in the angle between the leaf and the stem of the moss and pupates among the mosses. Both of these genera were present, but rarely, among the bryophytes of Appalachian Mountain mostly acid streams, USA (Glime 1968). They were more common on Fontinalis, where larvae of Oxyethira and Hydroptila sometimes decorated the branches of Fontinalis dalecarlica (Figure 98). From Ceylon, Schmid (1958) reported Chrysotrichia hapitigola, and Hydroptila kirilawela from large, mossy rocks in the torrent.

Figure 99. Ochrotrichia eliaga larva and case, a genus found on moss-covered granite outcrops in Appalachian streams. Photo by Trevor Bringloe, Biodiversity Institute of Ontario, through Creative Commons.

In a Tennessee, USA, springbrook, Ochrotrichia unio (see Figure 100) live among algae and mosses as larvae, then move to bare rocks to pupate (Stern & Stern 1969). In Great Britain, the larvae of this species feed on diatoms and other algae (Percival & Whitehead 1929).

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side by side with mosses such as Fontinalis (Figure 11). The species of liverwort depends on availability, with cases of Paleagapetus celsus from the eastern USA known from Scapania nemorea (Flint 1962; Glime 1978) (Figure 101), S. undulata (Glime 1978) (Figure 74), Plagiochila porelloides (Glime 1978) (Figure 102), Frullania sp. (Glime 1978) (Figure 103). In those I observed, the pieces of liverwort were cut into nearly circular pieces and cemented together along their margins, forming a case typical of many hydroptilids – the shape of an eyeglass case. Ito and Vshivkova (1999) described the pieces of liverworts comprising the cases of Palaeagapetus finisorientis from the Russian Far East similarly as being roughly rounded fragments.

Figure 100. Ochrotrichia larva, a genus in which some larvae live among mosses, then migrate to bare rocks to pupate Photo from California Department of Wildlife, through public domain.

Ptilocolepus Ptilocolepus granulatus is crenophilic (describing organism preferring spring environments but may also occupy similar habitats), living in montane to subalpine regions of central Europe (Waringer & Graf 2002). Wesenberg-Lund (1943) reported that Ptilocolepus granulatus lives in moss cushions and makes its case from moss fragments. Similarly, González et al. (2000) reported that P. extensus, an endemic on the Iberian Peninsula and a close relative, uses leaf pieces of several moss and liverwort species to make its final instar case. Unlike most of the Hydroptilidae, this case is flattened dorsiventrally, but still has the typical elongate-oval shape. In the Pyrénées, Thienemann (1950) and Décamps (1968) found Ptilocolepus granulatus among mosses and liverworts. These bryophytes also formed a significant portion of their food as well as construction material for their cases. Ito (1998) reported that this genus lives among, eats, and builds its cases from the leafy liverworts Chiloscyphus polyanthos (Figure 67) and Scapania undulata (Figure 74). Depisch (1999) and Ito and Higler (1993) all found that the species commonly lives among and feeds on the liverwort Scapania undulata. In Belgium Ptilocolepus granulatus uses Jungermannia riparia for food, but surprisingly, it also sometimes builds its case from the moss Fontinalis (Figure 11) (Ito & Higler 1993). Thus it is not surprising that Dittmar (1955) found it associated with Fontinalis. Ito and Higler found that it does not seem to feed on the moss, but later Ito (1998) states that it is the only species in the subfamily Ptilocolepinae that is able to feed on Fontinalis (and other mosses), attributing this ability to its large mandibles. Palaeagapetus Microcaddisflies such as Hydroptila (Figure 93) often attach their tiny homes to the moss leaves and stems, but Palaeagapetus in the same family constructs its home strictly out of leafy liverworts (Flint 1962; Glime 1978; Ito & Hattori 1986; Ito 1991), even when these are growing

Figure 101. Scapania nemorea, one of the species used for making cases of Palaeapetus celsus. Photo by Bernd Haynold, through Creative Commons.

Figure 102. Plagiochila porelloides, a species used by Palaeagapetus celsus for making its case. Photo by Hermann Schachner, through Creative Commons.

Figure 103. Frullania eboracensis, a terrestrial epiphytic species that may fall into the water and be used in the case of Palaeagapetus celsus. Photo by Bob Klips, with permission.

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Not only do members of this genus use liverworts in the construction of their cases, but the liverworts are also a primary food source (Botosaneanu & Levanidova 1987). In his review of four species of Palaeagapetus, Ito (1998) found that all of them used the liverworts Chiloscyphus polyanthos (Figure 67) and Scapania undulata (Figure 74) for food, housing, and case construction. It appears that all known members of the genus have this same strong dependence on leafy liverworts, including those in the eastern part of the former Soviet Union (Botosaneanu & Levanidova 1987), Japan (Ito & Hattori 1986; Ito 1988, 1991), and North America (Flint 1962; Glime 1978). In the western USA, Palaeagapetus nearcticus uses Scapania uliginosa for its case and food (Ito et al. 2014). The larvae pierce the cells and consume the liverwort one cell at a time. Ito and Vshivkova (1999) found that in the Palaeagapetus species they observed, the early instars fed on the contents of the liverwort cells, whereas the final instar cut off the leaves and apparently ingested them, reminiscent of human babies who also shift from sucking to chewing. Ito (1991) found that Palaeagapetus rotundatus feeds on the leaves of leafy liverworts Chiloscyphus polyanthos and Scapania undulata (Figure 74), but will not feed on the moss Platyhypnidium riparioides (Figure 13). Ito (1988) followed the life history of Palaeagapetus ovatus in a spring stream in Japan. He found that the density changed with season, reaching the highest in winter and being low in summer. Living with it was a predatory Trichoptera, Eubasilissa regina (Phryganeidae; Figure 77), that preyed upon it among the liverworts. We know more about this genus and its liverwort relationship through the description of a new species, Palaeagapetus ovatus, in Japan (Ito & Hattori 1986). This liverwort dweller fed exclusively on the leaves of the leafy liverwort Chiloscyphus polyanthos (Figure 67). Its fifth and final instar made the typical oval case from the leaves of this liverwort. And the females, within two days of emergence, laid 50-85 eggs on the leaves of this liverwort. The eggs do not form a mass and at 10.5-12°C they hatch in 21-23 days. Palaeagapetus nearcticus also deposits its orange eggs on liverwort leaves (Ito et al. 2014). More recently, Woods (2002) was surprised to find the thallose liverwort Riccardia chamedryfolia (Figure 104) moving in a slow, jerky motion on the sandy bottom of a pool in Wales. Investigation revealed that two matching pieces of the thallus had been cemented together by a caddisfly larva that was using it for a home (case). The larva was not identified but could have been a member of Hydroptilidae.

Figure 104. Riccardia chamedryfolia, a liverwort that some caddisflies use to make a case. Photo by Kristian Peters, with permission.

Figure 105. The caddisfly Scelotrichia willcairnsi (Hydroptilidae) with a case made of pieces of the moss Rhynchostegium brevinerve. Note the way pieces fit together as parallel rings. Photo courtesy of Andi Cairns.

Scelotrichia My email makes Christmas come all year-round. One of these nice surprises came when Andi Cairns sent me pictures of a caddisfly that was a bryological surprise. This new species, actually in a genus new to Australia, was Scelotrichia willcairnsi (Figure 105) living among the mosses in a waterfall (Figure 106). It was feeding on Rhynchostegium brevinerve (Figure 107), a new species previously thought to be Platyhypnidium muelleri and renamed by Huttunen and Ignatov (2010), in north-eastern Queensland, Australia. This microcosm was full of surprises!

Figure 106. Rhynchostegium brevinerve in Fishery Falls, Australia, home to Scelotrichia willcairnsi. Photo courtesy of Andi Cairns.

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Figure 109. Pieces of the moss Rhynchostegium brevinerve with numerous cases of the caddisfly Scelotrichia willcairnsi (Trichoptera: Hydroptilidae). Photo courtesy of Andi Cairns. Figure 107. Rhynchostegium brevinerve, home to the caddisfly Scelotrichia willcairnsi. Photo courtesy of Andi Cairns.

The Scelotrichia willcairnsi larva had a case (Figure 105) it had built by cementing moss leaf fragments together (Figure 108) – the same species of moss it was eating (Cairns & Wells 2008). It remained in this case to pupate, cementing it to the moss stems (Figure 109). When making a case, the larvae cut the leaves longitudinally, in parallel with the long axis of the leaf and its cells, giving them long pieces (Figure 108). Cairns and Wells described these: "neatly, the fragments fitted together, almost in rings." Ohkawa and Ito (2002) had already distinguished the types of cuts for leaves and for food in Scelotrichia ishiharai. This microcaddis uses the moss Rhynchostegium sp. (Figure 107-Figure 109) for food (Figure 110-Figure 111) and case building (Figure 105-Figure 109), likewise using different orientations for the two kinds of cuts.

When Cairns and Wells (2008) examined the gut contents, they discovered that these tiny caddisfly engineers cut the pieces of moss very differently for food than they did for cases. For food, they cut the leaves perpendicular to the long axis and across the cells (Figure 110-Figure 111). Such a cut would give the gut enzymes more access to the contents of the cells.

Figure 110. Pieces of the moss Rhynchostegium brevinerve from the gut of the caddisfly Scelotrichia willcairnsi (Hydroptilidae). Photo courtesy of Andi Cairns.

Figure 108. Pieces of the moss Rhynchostegium brevinerve from the case of the caddisfly Scelotrichia willcairnsi (Hydroptilidae). Photo courtesy of Andi Cairns.

Figure 111. Pieces of the moss Rhynchostegium brevinerve from the gut of Scelotrichia willcairnsi. The moss fragments are stained with Toluidine blue to make cell walls more evident. Note that cell contents appear to be gone in nearly all fragments, suggesting digestion. Photo courtesy of Andi Cairns.

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Chapter 11-12: Aquatic Insects: Holometabola – Trichoptera, Suborders Integripalpia and Spicipalpia

One of these larvae had included moss leaves, liverwort leaves, and even hornwort thallus all in one case (Chris Cargill, pers. comm. 30 March 2016). And all these pieces were still alive! Chris Cargill told me she later found discarded cases made of thalli from liverworts or hornworts and new thalli had started to grow from the case (Figure 112). I think we have just added a new means of bryophyte dispersal!

Figure 112. Scelotrichia willcairnsi caddis fly case old with living liverworts. Photo courtesy of Chris Cargill.

Elsewhere, in Papua New Guinea, Scelotrichia was similarly collected from mosses in the strong currents at the crest of a short waterfall (Wells 1990). They likewise made their cases of the moss leaves and later attached their pupal cases to the stems of the same species of moss. Wells found adults of two other species of Scelotrichia near waterfalls or soaked mosses. As in S. willcairnsi (Figure 105), the caddisfly larvae from Papua New Guinea had cut slivers of the moss down the long axis of the leaf, making the cells parallel to the length of the fragment. These differed from the pieces cut by Paleagapetus and Ptilocolepus, which were cut from leafy liverworts and glued together to resemble a patchwork quilt (Ito 1998; Ito & Higler 1993). It appears that cutting behavior can determine the type of bryophyte that is suitable for making the case.

Rhyacophiloidea Rhyacophilidae – Free-living Caddisflies This is a Northern Hemisphere family from the temperate parts of North America, Europe, and Asia, extending into India and the tropical areas of southeastern Asia (Kjer 2010). The larvae are 9-16 mm long and are green or brown, blending easily with the bryophytes (Bumble.org 2013). Don't be misled by the pink color they assume in preservative. Larvae of this family do not build cases (Figure 113), so they do not attach themselves to the substrate by gluing their cases like some caddisflies do. Their life cycle is one year, with two generations overlapping. The larvae prefer rapid, cold streams where they are able to stay themselves in the current by clinging to mosses or debris (Hilsenhoff 1975). Most are carnivorous, but a few are herbivorous. And some can live above the water level among wet

emergent mosses: Rhyacophila nubila (Figure 114), R. polonica, and R. tristis, whereas in the same River Rajcianka, Slavakia, these three species plus R. obliterata (Figure 115), R. philopotamoides, and R. vulgaris occur among the mosses under water (Krno 1990).

Figure 113. The free-living caddisfly, Rhyacophila, is a common member of the stream moss community. Its color is typically green, and it has large hooks that permit it to cling to mosses and other substrata to avoid being washed away by the fast-flowing water it inhabits. Its lack of a case permits it to traverse the internal chambers of the moss without getting caught by the branches. Photo by Janice Glime.

Figure 114. Rhyacophila nubila larva, a species that can live among mosses above or below the water surface. Photo by Niels Sloth, with permission.

In my studies of Appalachian Mountain stream mosses in Maryland and Pennsylvania, USA, the genus Rhyacophila was among the most common and constant of the caddisfly larvae among the bryophytes. Décamps (1967, 1968) found Rhyacophila laevis to be abundant among mosses in the Pyrénées. In a cool mountain stream of central Japan, Tada and Satake (1994) found that R. towadensis was significantly more abundant among the moss Platyhypnidium riparioides (Figure 13) than in bare rock areas. Many members of Rhyacophilidae most likely benefit both from the protection afforded by the bryophytes, but also from the resident fauna that serves as food, especially the numerous Chironomidae. In their study of four small Appalachian, USA, streams, Woodall and Wallace (1972) found larvae of Rhyacophila torva (Figure 125) (see also Roback 1975), R. nigrita (Figure 116), R. carolina (Figure

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122), R. minora (Figure 117) (see also Glime 1968), R. glaberrima (Figure 118), and R. fuscula (Figure 123Figure 124) among mats of mosses on rock outcrops. They fed on the Chironomidae larvae (Ross 1944) that shared the bryophyte habitat. In one of my collections from the mid-Appalachian Mountains I caught R. carolina in the act – it was preserved with a chironomid larva in its mouth. Although R. minora in a wooded Ontario, Canada, stream is typically carnivorous, early instars feed on plant material (Singh et al. 1984). This strategy works well until they gain the size and skill to be predators. Figure 117. Rhyacophila minora larva, an Appalachian Mountain stream bryophyte dweller. Photo from Biodiversity Institute of Ontario, through Creative Commons.

Figure 118. Rhyacophila glaberrima larva, a common species among mosses in the Appalachian Mountain streams. Photo by Donald S. Chandler, with permission.

Food

Figure 115. Rhyacophila obliterata adult, a species whose larvae are common among bryophytes. Photo by James K Lindsey, with permission.

Figure 116. Rhyacophila nigrita larva, a moss dweller in Appalachian Mountain streams. Photo by Donald S. Chandler, with permission.

Most Rhyacophila species are carnivores that do not make cases, but the Verrula group eat photosynthetic organisms with their hypognathous heads (oriented downwards), feeding on algae, diatoms, and particularly bryophytes (Smith 1968; Thut 1969). Cummins (1973) likewise reported that R. verrula in western North America is a herbivore and especially eats aquatic mosses (Slack 1936; Gerson 1982; Smith 1968). In his study of diets of the Rhyacophila species in constructed streams in western USA, Thut (1969) found that R. verrula feeds predominantly on aquatic mosses. This effect is intensified in winter when several mosses are dominant and diatoms are abundant. Interestingly, diatoms become more important in the fourth and fifth instars than they are in earlier instars. In a Tennessee cold springbrook, Rhyacophila lobifera larvae fed among the moss and algae, eating smaller caddisfly larvae, midge larvae, naiads of mayflies and stoneflies, detritus, and diatoms (Stern & Stern 1969). Slack (1936) also reported that one out of nine Rhyacophila dorsalis (Figure 119) had leaves of Fontinalis antipyretica (Figure 11) in the gut, but that it is primarily carnivorous. Nevertheless, one specimen contained only diatoms in the gut and the one with Fontinalis had only plant material. In a study in the English Lake District, Elliott (2005) found that early instars ate primarily diatoms (mostly Achnanthes spp., Figure 120), with bryophyte fragments also present in nearly all gut samples, but the bryophytes appeared to be undigested, displaying their chlorophyll. These bryophytes may have been eaten to obtain adhering diatoms. Both second and third instars would disappear into the bryophyte clumps to search for prey, but they returned to the surface of those

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clumps to consume their finds. Fourth and fifth instars fed only at night and used an ambush strategy to capture prey, which includes Baetis and Gammarus.

almost totally at night (Elliott 2005). They can feed on other insects inhabiting their moss habitat, such as Ephemeroptera (mayflies), Simuliidae (blackflies), and Chironomidae (midges). As they grow older, instars 4 and 5, they adopt an ambush strategy at dusk and dawn, catching such active prey as the mayfly Baetis and the scud Gammarus. During the night they used a searching strategy to capture the more sedentary prey, for example Chironomidae (midges) and Simuliidae (blackflies). Thut (1969) suggested that the high proportion of moss fragments in the diets of the herbivorous Rhyacophila was at least in part the result of seasonal changes in the available primary producers in streams. Bryophytes are available in winter when most of the algae are dormant in a resting stage. Substrate Preference

Figure 119. Rhyacophila dorsalis larva, a carnivorous species that sometimes has leaves of Fontinalis antipyretica in its gut. Photo by Walter Pfliegler, with permission.

Larvae of most of the predominantly carnivorous Rhyacophila dorsalis (Figure 119) occur among bryophytes [leafy liverwort Scapania sp. (Figure 74) and mosses Platyhypnidium riparioides (Figure 13) and Fontinalis antipyretica (Figure 11)] (Slack 1936). For less active prey they use a searching strategy (Chironomidae, Simuliidae). The percentage of larvae with bryophytes in the gut was much smaller than that of prey. It appears that this species changes its diet as it grows, but it may also be an opportunist regarding its diet. But if one considers that both the diatoms and bryophytes still had chlorophyll in their cells, it appears that even the first and second instar larvae may have been carnivores, eating these photosynthetic organisms by chance while attempting to capture prey. Instead, the first and second instar larvae eat copepods, rotifers, and tardigrades, common bryophyte inhabitants, but these require special preservation techniques in order to recognize them in gut samples. Instead of a shift from apparent herbivore to carnivore, Elliott (2005) demonstrated a shift in size of prey.

Rhyacophila species typically make their larval homes under rocks or among mosses (Bouchard 2004). They are able to use their claws (Figure 121) to anchor themselves or cling to the mosses, but also use them as they creep along in the stony stream bed (Badcock 1949). Percival and Whitehead (1929) found that Rhyacophila dorsalis (Figure 119) preferred thick mosses and Potamogeton on stones. Elliott (2005) found some larvae found under large stones, but most were among bryophytes growing on the upper surfaces of large stones [Scapania (Figure 74), Platyhypnidium riparioides (Figure 13), Fontinalis antipyretica (Figure 11)].

Figure 121. Rhyacophila fuscula larva showing anal hooks that cling to its substrate. Photo by Jason Neuswanger, with permission.

Figure 120. Achnanthes longipes. Chepurnov, through non-commercial license.

Photo by Victor

The caddis larvae of Rhyacophila dorsalis (Figure 119) begin their early instars by feeding equally day and night, but by the 4th to 5th instar they shift to feeding

In the Great Smoky Mountains National Park, R. montana lives in the films of water that flow over vertical rock faces, crevices, or among wet mosses (Parker et al. 2007). Rhyacophila evoluta and R. intermedia are characteristic of mosses in torrents in the Pyrénées (Décamps 1967). Rhyacophila evoluta has the ability to go into a cold-induced diapause at any stage in its development. This permits it to complete its development in one, two, or three years, depending on the temperatures. Some species seem to prefer liverworts and some to prefer mosses for their homes (locations, not cases). In the

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mid-Appalachian Mountain streams I found Rhyacophila cf. carolina (Figure 122) primarily among liverworts (Scapania undulata; Figure 74), whereas R. fuscula (Figure 121, Figure 123-Figure 124) predominated in Fontinalis dalecarlica (Figure 98) and R. torva (Figure 125) in Hygroamblystegium fluviatile (Figure 4-Figure 5) and Platyhypnidium riparioides (Figure 13). Rhyacophila invaria (Figure 126) occurred frequently among clumps of the moss Platyhypnidium riparioides (36% frequency) but was absent among Hygroamblystegium fluviatile clumps despite the frequent intermingling of these two mosses. It reached its greatest numbers in Scapania undulata.

Figure 122. Rhyacophila carolina larva, species that is common among clumps of the leafy liverwort Scapania undulata in Appalachian Mountain, USA, streams. Photo by Bob Henricks, with permission.

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Figure 125. Rhyacophila torva larva, a moss dweller in Appalachian Mountain streams. Photo by Trevor Bringloe, Biodiversity Institute of Ontario, through Creative Commons.

Figure 126. Rhyacophila invaria larva, a species that occupies both mosses and liverworts in Appalachian Mountain streams. Photo by Donald S. Chandler, with permission.

Summary

Figure 123. Rhyacophila fuscula larva, a moss dweller on boulders in the Appalachian Mountain streams. Photo by Donald S. Chandler, with permission.

Figure 124. Rhyacophila fuscula pupa. Henricks, with permission.

Photo by Bob

The Limnephilidae are mostly large and therefore are usually absent from the smaller mosses. However, sometimes several may occur within a clump of Fontinalis. The Brachycentridae are common among bryophytes. Some (Micrasema, Adicrophleps hitchcockii) use mosses in their cases and some also eat them. The genera Palaeagapetus and Scelotrichia, both in the Hydroptilidae, use bryophytes (exclusively?) for food and case construction, the former using leafy liverworts and the latter using mosses. In the same family, Ptilocolepus uses both mosses and liverworts for food and in case construction. The family Rhyacophilidae is a free-living caddisfly and is mostly carnivorous. However, some of the bryophyte dwellers eat bryophytes, whereas others use them as a place to capture prey. Other families that can be found among bryophytes less commonly include Odontoceridae, Goeridae, Limnephilidae, Lepidostomatidae, Oeconesidae (especially in New Zealand), Uenoidae, Phryganeidae, Beraeidae, Conoesucidae, Helicophidae, Sericostomatidae, and Glossosomatidae. Among these, the Limnephilidae and Phryganeidae have mostly large larvae that are unable to move about in most of the bryophytes but that can live among the large branches of Fontinalis species. Unlike the Coleoptera, this order is poorly represented in bogs and fens, but they are common in streams and less so in lakes.

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Acknowledgments As a graduate student I relied heavily on the expertise of Oliver Flint, Glenn Wiggins, Tom Waters, and Ken Cummins for both encouragement and identification help in my novice years. Thank you so much to D. N. Bennett for making a special trip to photograph live Micrasema wataga eating and showing the mosses sprouting on the case, just for this chapter! She also made additional trips to the field to determine the identity of the moss. David Tempelman helped me to obtain some of the images and permission for use and provided me with references on Ptilocolepus. Eileen Dumire proofread the chapter and helped me improve the clarity and readability.

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Pritchard, G. and Berté, S. B. 1987. Growth and food choice by two species of limnephilid caddis larvae given natural and artificial foods. Freshwat. Biol. 18: 529-535. Redell, L. A., Gall, W. K., Ross, R. M., and Dropkin, D. S. 2009. Biology of the caddisfly Oligostomis ocelligera (Trichoptera: Phryganeidae) inhabiting acidic mine drainage in Pennsylvania. USGS Publications Warehouse. Roback, S. S. 1975. New Rhyacophilidae records with some water quality data. Proc. Acad. Nat. Sci. Philadelphia 127(5): 45-50. Ross, H. H. 1944. The caddis flies, or Trichoptera, of Illinois. Ill. Nat. Hist. Surv. Bull. 23: 326 pp. Rousseau, E., Lestage, J., and Schouteden, H. 1921. Les Larves et Nymphes Aquatiques des Insectes dEurope. Vol. 1. Office de Publicite, Brussels. Schmid, F. 1958. Trichopteres de Ceylan. Arch. Hydrobiol. 54: 1-173. Singh, M. P., Smith, S. M., and Harrison, A. D. 1984. Life cycles, microdistribution, and food of two species of caddisflies (Trichoptera) in a wooded stream in southern Ontario. Can. J. Zool. 62: 2582-2588. Slack, H. D. 1936. The food of caddis fly (Trichoptera) larvae. J. Anim. Ecol. 5: 105-115. Smith, S. D. 1968. The Rhyacophila of the Salmon River drainage of Idaho with special reference to larvae. Ann. Amer. Entomol. Soc. 61: 655-674. Stern, M. S. and Stern, D. H. 1969. A limnological study of a Tennessee cold springbrook. Amer. Midl. Nat 82: 62-82. Suren, A. M. 1988. Ecological role of bryophytes in high alpine streams of New Zealand. Internat. Ver. Theor. Angew. Limnol. 23: 1412-1416. Suren, A. M. and Winterbourn, M. J. 1991. Consumption of aquatic bryophytes by alpine stream invertebrates in New Zealand. N. Z. J. Marine Freshwat. Res. 25: 331-343. Tada, M. and Satake, K. 1994. Epiphytic zoobenthos on bryophyte mats in a cool mountain stream, Toyamazawa. Jap. J. Limnol. 55: 159-164. Thienemann, A. 1950. Verbreitungsgeschichte der Süsswassertierwelt Europas. Binnengewässer 18: 809 pp. Thut, R. N. 1969. Feeding habits of larvae of seven Rhyacophila species with notes on other life-history features. Ann. Amer. Entomol. Soc. 62: 894-898. Wallace, J. B . and Ross, H. H. 1971. Pseudogoerinae: A new subfamily of Odontoceridae (Trichoptera). Ann. Entomol. Soc. Amer. 64: 890-894. Waringer, J. and Graf, W. 2002. Ecology, morphology and distribution of Ptilocolepus granulatus (Pictet 1834) (Insecta: Trichoptera) in Austria. Lauterbornia 43: 121-129. Watson, L. and Dallwitz, M. J. 2003 onwards. British insects: The families of Trichoptera. Version: 9 April 2007. Accessed 20 July 2007 at . Watson, L. and Dallwitz, M. J. 2011. British insects: The families of Caddis flies (Trichoptera). Accessed 20 January 2015 at . Wells, A. 1990. The hydroptilid tribe Stactobiini (Trichoptera: Hydroptilidae: Hydroptilinae) in New Guinea. Invert. System. 3: 817-849. Wesenberg-Lund, C. 1943. Biologie der Susswasserinsekten. Otto Koeltz, Kopenhagen. Wiggins, G. B. 1973. New systematic data for the North American caddisfly genera Lepania, Goeracea and Goerita (Trichoptera: Limnephilidae). Royal Ontario Mus. Life Sci. Contrib. 91: 1-33.

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Wiggins, G. 1977. Larvae of the North American Caddisfly Genera (Trichoptera). University of Toronto Press, Toronto, pp. 52-53. Wiggins, G. B. 2004. Caddisflies: The underwater architects. University of Toronto Press, Toronto, pp. 187-189. Wiggins, G. B. and Anderson, N. H. 1968. Contributions to the systematics of the caddisfly genera Pseudostenophylax and Philocasca with special reference to the immature stages (Trichoptera: Limnephilidae). Can. J. Zool. 4: 61-75. Wiggins, G. B. and Winchester, N. N. 1984. A remarkable new caddisfly genus from northwestern North America (Trichoptera, Limnephilidae, Limnephilinae). Can. J. Zool. 62: 1853-1858.

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Winchester, N. N., Wiggins G. B., and Ring, R. A. 1993. The immature stages and biology of the unusual North American Arctic caddisfly Sphagnophylax meiops, with consideration of the phyletic relationships of the genus (Trichoptera: Limnephilidae). Can. J. Zool. 71: 1212-1220. Winterbourn, M. J. and Gregson, K. L. D. 1981. Guide to the aquatic insects of New Zealand. Bull. Entomol. Soc. N.Z. 5: 1-80. Woodall, W. R. Jr. and Wallace, J. B. 1972. The benthic fauna in four small southern Appalachian streams. Amer. Midl. Nat. 88: 393-407. Woods, R. G. 2002. Thalloid liverwort on an involuntary subaquatic peripatetic substrate in Wales. Bull. Brit. Bryol. Soc. 78: 56-57.

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Glime, J. M. 2017. Aquatic Insects: Holometabola – Diptera, Suborder Nematocera. Chapt. 11-13a. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. Ebook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 19 July 2020 and available at .

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CHAPTER 11-13a AQUATIC INSECTS: HOLOMETABOLA – DIPTERA, SUBORDER NEMATOCERA TABLE OF CONTENTS DIPTERA – Flies .......................................................................................................................................... 11-13a-2 Suborder Nematocera............................................................................................................................. 11-13a-5 Nymphomyiidae.............................................................................................................................. 11-13a-6 Cylindrotomidae – Long-bodied Craneflies.................................................................................... 11-13a-6 Limoniidae – Limoniid Craneflies .................................................................................................. 11-13a-8 Pediciidae – Hairy-eyed Craneflies............................................................................................... 11-13a-11 Tipulidae – Craneflies ................................................................................................................... 11-13a-11 Anisopodidae – Wood Gnats, Window Gnats .............................................................................. 11-13a-19 Axymyiidae................................................................................................................................... 11-13a-19 Cecidomyiidae – Gall Midges, Gall Gnats ................................................................................... 11-13a-20 Mycetophilidae – Fungus Gnats ................................................................................................... 11-13a-20 Sciaridae – Dark-winged Fungus Gnats........................................................................................ 11-13a-20 Ceratopogonidae – Biting Midges, No-see-ums, Sand Flies......................................................... 11-13a-20 Summary .............................................................................................................................................. 11-13a-22 Acknowledgments................................................................................................................................ 11-13a-22 Literature Cited .................................................................................................................................... 11-13a-22

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CHAPTER 11-13a AQUATIC INSECTS: HOLOMETABOLA – DIPTERA, SUBORDER NEMATOCERA

Figure 1. Triogma trisulcata among mosses. This species makes its home among wet mosses of bogs and swamps and is effectively a moss mimic. Photo by J. C. Schou, with permission.

DIPTERA – FLIES Gerson (1969) suggested that the ancestral fly groups originated among mosses where it is always damp. Because the systematics of the fly groups are still poorly understood, I have divided the treatments into the two suborders, Nematocera and Brachycera. From there they are alphabetical within superfamilies, but the superfamilies are not delineated by name. Diptera adults are distinguished by having only two wings, as reflected in the name of Diptera (di = 2; pteron = wing). In place of the second pair of wings the flies have a pair of halteres (Figure 2), thoracic projections that resemble lollipops, one on each side of the thorax. In the larval stage, they are distinguished by having only fleshy prolegs (Figure 9) or no legs. They lack the chitinized, jointed thoracic legs found in most larval insects (Johannsen 1969).

Figure 2. Tipulidae showing two wings and halteres. Photo by Pinza, through Creative Commons.

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Gerson (1982) reported a number of bryophytedwelling Diptera of medical and veterinary importance because they bite. Among these are the sand flies [Psychodidae (see Chapter 13b; Quate 1955)], mosquitoes [Culicidae (see Chapter 13b; Fantham & Porter 1945)], black flies [Simuliidae (Figure 3); Snow et al. 1958)], biting midges [Ceratopogonidae (Figure 84-Figure 88; Séguy 1950)], and horse flies [Tabanidae (Figure 4; Teskey 1969)]. All of these are discussed in this chapter except Tabanidae. I found it only occasionally among bryophytes in Appalachian Mountain, USA, streams; the other studies I reviewed did not mention it.

Figure 5. Fontinalis antipyretica, home for numerous aquatic insects and suitable for larger ones. Photo from Projecto Musgo, through Creative Commons.

Figure 3. Simuliidae larvae in the rapid flow of a stream. Photo by Janice Glime.

Bryophytes accumulate coarse (CPOM), fine (FPOM), and ultrafine (UPOM) particulate organic matter that serves as a food source for their inhabitants (Habdija et al. 2004). These conditions favor small forms of oligochaetes, Diptera, and Coleoptera that comprise 64-99% of the macrophyte (plant – especially aquatic – large enough to be seen without a lens) individuals. Collector gatherers dominate in spring and summer, collector-filterers in autumn, and scrapers in winter. In a cool mountain stream in central Japan, five of the six taxa of Diptera identified (mostly at the level of family or subfamily) were significantly more abundant in clumps of the moss Platyhypnidium riparioides (Figure 6) than in areas of bare stones (Kato 1992). These included Limoniidae (Antocha spp.; Figure 7), Simuliidae (Figure 3), and Chironomidae [Figure 8; Tanypodinae, Diamesinae, Orthocladius spp.].

Figure 4. Chrysops divaricatus (Tabanidae) adult, an adult pest (horse fly) whose larvae sometimes live among the bryophytes. Photo by Kallema, through Creative Commons.

In streams, bryophytes are often important contributors to biodiversity. Flow rates are important in determining the type of Diptera able to live there. The abundance of Chironomidae (see Chapter 13b) is negatively correlated with flow rate as it approaches clumps of mosses (Fontinalis antipyretica; Figure 5), whereas the abundance of the smallest Simuliidae (Figure 3) is positively correlated (Linhart et al. 2002a). In the Plitvice Lakes National Park in the Dinaric karst region of Croatia, Čmrlec (2013) found that the Diptera families were least abundant in silt and that mosses were the preferred substrate. These correlations with speed and silt do not prevent both groups of species from living in the same bryophyte clump – the slow-water silt lovers live near the bottom while the fast-water silt avoiders live near the surface of the bryophyte clump.

Figure 6. Emergent but wet Platyhypnidium riparioides in Europe, a common home for Diptera. Photo by Michael Lüth, with permission.

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Figure 7. Antocha, a larva that inhabits the moss Platyhypnidium riparioides (Figure 6) in cool mountain streams in Japan. Photo by Bob Henricks, with permission.

In Alaska, Diptera dominate by an even larger proportion than in streams of temperate North America (Oswood 1989). The Chironomidae (Figure 8) exhibit a significant increase from south to north, whereas most other taxa (excluding Nemouridae) decrease. Figure 9. Deuterophlebia ventral side showing suction cups. Photo from Aquatic Bioassessment Laboratory , with permission.

Figure 8. Chironomidae larva, a common bryophytedwelling family whose numbers increase from south to north. Photo by Jason Neuswanger, with permission.

The Diptera have a variety of adaptations to their aquatic domicile of choice. For example, Bass and Cooling (1983) reported that Muscidae (Brachycera), Ichneumonidae (Hymenoptera), and Simuliidae (Figure 3) were associated with mosses below a reservoir in southern England. Both the larvae and pupae had posterior projections to anchor them to the mosses. Amos (1999) describes the role of the brook moss Fontinalis (Figure 5) in providing a safe habitat in the torrent, and this moss likes cold water (Glime 1987) where few tracheophytes persist. Here one can find many small invertebrates, but it seems still to be a challenge to stay put. The mountain midge larva (Deuterophlebiidae, Figure 9) survives the torrent by the use of strong suction to hold the rock. The suction cups of Deuterophlebia (Figure 9) are of little use among bryophytes, but are fantastic for adhering to "bare" rocks. Respiratory adaptations are numerous and will be discussed for the various families. The floating community includes only a few species of bryophytes, notably Ricciocarpos natans (Figure 10) and Riccia fluitans (Figure 11). In some cases, the Diptera associated with the thallose floating liverwort Ricciocarpos natans are the same ones found among floating tracheophytes such as Spirodela, Lemna minor (Figure 10), and Wolffia (Scotland 1934).

Figure 10. Ricciocarpos natans and Lemna minor, floating plants that can harbor surface-dwellers. Photo by Jan-Peter Frahm, with permission.

Figure 11. Riccia fluitans with pearling (oxygen bubbles produced by the plants), a floating community that provides cover and oxygen for aquatic insects. Photo by Christian Fischer, through Creative Commons.

Despite the number of families of Diptera among the bryophytes, and the presence of such mixed terrestrial/aquatic families as the Tipulidae (Figure 46-

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Figure 73, Figure 75, Figure 77-Figure 76), it is interesting that this order is poorly represented among the wet emergent mosses in the River Rajcianka in Slovakia (Krno 1990). Only the Psychodidae (see Chapter 13b) were able to take advantage of the safety of the emergent bryophytes there. On the other hand, fauna of the submerged mosses were represented by not only the Psychodidae, but also the Ceratopogonidae (Figure 84-Figure 88) and Simuliidae (Figure 3). Conspicuously absent in these eutrophic (referring to lake or other body of water rich in nutrients and thus supporting dense plant/algal populations) waters were the Tipulidae and Chironomidae (Figure 8). Occasionally, or perhaps frequently, the insects do something beneficial for the bryophytes they visit. In a study to determine the role of adult Diptera in dispersing algae and Protozoa, Revill et al. (1967) found that in addition to 21 species of viable algae and 5 of Protozoa, the washings from the four species of Diptera produced viable moss spores/protonemata as well. These transporting insects included Tipula triplex (Tipulidae; Figure 12), Bittacomorpha clavipes (Ptychopteridae, Figure 13), Chaoborus punctipennis (Chaoboridae, Figure 14-Figure 15), and Chironomus (Chironomidae; Figure 16).

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Figure 14. Chaoborus punctipennis adult, a species known to carry bryophyte spores/protonemata. Photo by Tom Murray, through Creative Commons.

Figure 15. Chaoborus sp., larva of one of the Diptera known to carry bryophyte spores/protonemata. Photo by Viridiflavus, through Creative Commons.

Figure 12. Tipula triplex adult, a cranefly known to disperse bryophyte spores or protonemata. Photo by Paul Rhine , through Creative Commons.

Figure 16. Chironomus dorsalis adult, an insect known to transport bryophyte spores or protonemata. Photo by James K. Lindsey, with permission.

Suborder Nematocera

Figure 13. Bittacomorpha clavipes adult, a phantom cranefly that carries bryophyte spores or protonemata. Photo by Matt Muir, through Creative Commons.

The name Nematocera means "thread horns" and refers to the long, threadlike antennae. These are elongated flies with thin, segmented antennae. The larvae are mostly aquatic and the family includes craneflies, gnats, midges, mosquitoes, and blackflies.

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Nymphomyiidae This is a family of tiny (2 mm) flies in the northern parts of the Northern Hemisphere, especially eastern North America and eastern and central Asia (Nymphomyiidae 2013). The adults are neotenic (retaining larval or immature characters in adulthood), with straplike wings having poor venation (few wing veins). They live in running waters, where they often are found on mosscovered rocks, and pupation (development process between larva and adult) usually occurs in the same place (Courtney 1994). Adults have aborted mouth parts and live only a short time, some dying while still in the copulatory (mating) position. Nymphomyia is the only genus currently listed in this family (Myers et al. 2014). It lives among aquatic mosses in small, rapid streams (Courtney 1994; Courtney et al. 1996). Not only larvae, but also often pupae and adults of Nymphomyia, live on rocky substrates covered with aquatic mosses such as Platyhypnidium riparioides (Figure 6), Fontinalis (Figure 5), and Hygroamblystegium (Figure 91) (Cutten & Kevan 1970; Adler et al. 1985).

Triogma trisulcata (Figure 17) larvae are inhabitants of semiaquatic mosses, especially in stagnant water in bogs (Brinkmann 1997). In contrast to the tracheal gill respiration of Phalacrocera replicata (Figure 18), another bryophyte dweller in this family, the larvae lie on the leaves of the moss in a position that places the spiracular disk (apparatus that contains the breathing openings called spiracles) at the level of the water surface. Like P. replicata, these larvae have appendages that match the color and mimic the morphology of the surrounding mosses. These have been variously interpreted as mimetic camouflage to protect them against enemies and as respiratory organs. It seems reasonable that both interpretations may be correct. The pupae remain in these same positions until a short time before the adults emerge (ecdysis). Just before ecdysis, they search for drier mosses. Eggs are laid singly on mosses just below the surface by females dipping the tip of the abdomen into the water to touch the leaves. The eggs are attached by an adhesive.

Cylindrotomidae – Long-bodied Craneflies The family Cylindrotomidae is often separated from the Tipulidae (Figure 46-Figure 73, Figure 75, Figure 77Figure 76), which I have chosen to do to make it easy to discuss its unique characters relative to bryophytes. These are of moderate size (11-16 mm) and yellowish to pale brownish as adults (Cylindrotominae 2014). Most larvae live among mosses – terrestrial, semiaquatic, and aquatic mosses (Cylindrotominae 2014), and feed on mosses and tracheophytes (plants with lignified vascular tissue) (Gelhous et al. 2007). The family occurs mostly in the Holarctic and Oriental Regions, but there are scattered records in southern South America, New Guinea, and Australia. The aquatic insects don't seem to have the elaborate camouflage known in some terrestrial insects, but some still do an excellent job at blending. The Cylindrotomidae in particular are bryophyte dwellers and are world-class mimics of that habitat – bryocamouflage! The larvae of Triogma trisulcata (Figure 1, Figure 17) are known for their mimicry in a Sphagnum (Figure 69) habitat, but they also occur in streams where the larvae attach to Fontinalis antipyretica (Figure 5) (Gerson 1969). The leaflike appendages most likely are equally useful in that habitat as camouflage.

Figure 17. Triogma trisulcata larva posterior showing flanges that make it almost invisible among Sphagnum. Photo by Walter Pfliegler, with permission.

Figure 18. Phalacrocera replicata larva, an effective moss mimic that develops among mosses. Photo through Wikimedia Commons.

Phalacrocera replicata (Figure 18) lives among Sphagnum (Figure 69), Fontinalis antipyretica (Figure 5), and Warnstorfia fluitans (Figure 19) (Brinkmann 1997). Larvae in this species find tufts of mosses, then attach themselves to the leaves and stalks by affixing the anterior part of the body using the mandibles (crushing organs in an arthropod's mouthparts) to grab onto the edge of a leaf. They then crawl by crooking the body and securing the dorsal hooks. They have backward-pointing appendages that presumably help prevent them from being swept away by the current. At this stage they have functional spiracles that they do not use. Instead, the long, filiform appendages along the body function as tracheal gills, supplemented by cutaneous (referring to outer cuticle of insect body) gas exchange. But when it is time for pupation, the larvae move to the water surface to expose their spiracles (external openings through which insects breathe) to the atmospheric air. To maintain this contact with surface air, the pupae hang beneath the surface film, using their

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respiratory horns, and cling to the stems of mosses or other plants with the appendages on the last of the abdominal segments, positioning their bodies horizontally.

Figure 19. Warnstorfia fluitans, one of the homes of larvae of Phalacrocera replicata. Photo by Michael Lüth, with permission.

Figure 21. Liogma nodicornis adult, a species whose green larvae have markings that make them look like the leafy mosses where they live. Photo by Ilona L., through Creative Commons.

Clymo and Hayward (1982) reported that Phalacrocera replicata feeds on Sphagnum (Figure 69). Miall and Shelford (1897) found that P. replicata (Figure 18) larvae eat Warnstorfia exannulata (Figure 20). They described pupae that attach to the moss leaves by dorsal appendages on posterior segments. The females lay about 60 eggs in axils (upper angle between leaf stalk or branch and stem from which it grows) of the moss leaves.

Figure 22. Rhytidiadelphus squarrosus, home and food for Liogma (Figure 21) and Triogma (Figure 17) larvae. Photo by Michael Lüth, with permission.

Figure 20. Warnstorfia exannulata, food for Phalacrocera replicata (Figure 18). Photo by Michael Lüth, with permission.

Byers (1961) reported that the larvae of Liogma (Figure 21) use bryophytes for their larval habitats. Larvae of the genera Liogma and Triogma (Figure 17) have a green color with markings that make them look like leafy mosses (Gerson 1969). These two genera live among and eat the mosses Rhytidiadelphus squarrosus (Figure 22) and Hypnum cupressiforme (Figure 23). Larvae of Triogma trisulcata (Figure 17) inhabit the brook moss Fontinalis antipyretica (Figure 5) in mountain streams (Alexander 1920). These larvae have appendages that resemble leaves on a branch, and the color is typically green and black.

Figure 23. Hypnum cupressiforme, home and food for Liogma (Figure 21) and Triogma (Figure 17) larvae. Photo by Li Zhang, with permission.

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Limoniidae – Limoniid Craneflies The Limoniidae (Figure 24) family is an offshoot of the Tipulidae and thus many of the taxa discussed here were originally reported as members of Tipulidae. They are a worldwide family, mostly aquatic, and of moderate size (Limoniidae 2015). Their feeding groups vary considerably, including phytophagous (eating plants), saprophagous (eating dead organisms), mycetophagous (eating fungi), and carnivorous (eating animals) species.

Figure 25. Geranomyia sp adult. Geranomyia rostrata larvae live among mosses and thallose liverworts in North and South America. Photo by Ted Kropiewnicki, through Creative Commons.

Figure 24. Limoniidae adults mating, a family with larvae that often live among mosses, some consuming them. Photo by Anki Engström at , with permission.

From Cape Town, South Africa, we have a report of the Limoniidae occupying mosses in the stream of an isolated mountain (Harrison & Barnard 1972). The genus Geranomyia rostrata (see Figure 25) lives among algae, wet mosses, and thallose liverworts in the eastern part of North and South America (Rogers 1927; Johannsen 1969). These larvae are greenish and translucent (allowing light but not clear images to pass through), slow movers, and herbivores on algae and moss (Johannsen 1969). Geranomyia sexocellata (see Figure 25) larvae live in a gelatinous tube made with minute sand grains and attached to mosses in waterways that are only trickles. By contrast, Dicranomyia capicola (syn. of Limonia capicola?; see Figure 26) larvae live among mosses at the edge of a rapidly flowing streamlet (Harrison & Barnard 1972) and larvae of Limonia sp. and Ormosia sp. (Figure 28) live among bryophytes in Appalachian Mountain streams (Glime 1968). Harrison and Barnard (1972) also found Elephantomyia aurantiaca (see Figure 29) larvae among the damp mosses and liverworts. Several researchers have reported Limonia species from bryophytes (Byers 1961; Hilsenhoff 1975; Suren 1991). Suren (1991) found that Limonia hudsoni (see Figure 27) apparently required more from the bryophytes than just a substrate. It failed to colonize the artificial bryophytes in his New Zealand stream studies. Instead, Suren and Winterbourn (1991) reported that it actually commonly consumes bryophytes. Apparently artificial ones couldn't fill the bill.

Figure 26. Dicranomyia modesta adult, member of a genus with some larvae that live among mosses at streambanks. Photo by James K. Lindsey, with permission.

Figure 27. Limonia wellingtonia, member of a genus with some moss-dwelling members. Photo by Stephen Moore, Landcare Research, NZ, with permission.

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Figure 30. Scapania undulata, home for several species of Hexatoma. Photo by Michael Lüth, with permission.

Figure 28. Ormosia adult, a genus whose larvae sometimes live among mosses. Photo by Malcolm Storey, through Creative Commons.

Figure 31. Hexatoma larva; some members of this genus eat mosses. Photo by Jason Neuswanger, with permission.

Figure 29. Elephantomyia westwoodii adult female; larvae live among damp mosses and liverworts. Photo by Robert Lord Zimlich, through Creative Commons.

An important use of bryophytes can be that of providing a place for them to emerge. Rhipidia maculata emerges from the stream bed and also from thin moss layers on exposed rocks (Needham 1908; Johannsen 1969). In my studies of Appalachian Mountain stream moss communities, both Hexatoma cf. longicornis and H. cf. spinosa occurred among the leafy liverworts Scapania undulata (Figure 30) (Glime 1968). Hexatoma (Figure 31Figure 32) is known to ingest mosses (Percival & Whitehead 1929), so perhaps it is looking for food.

Figure 32. Hexatoma (Eriocera) gravelyi male adult. Photo by Muhabbet Kemal, with permission.

Limnophila occurs among bryophytes in several locations (Alexander 1919; Hilsenhoff 1975). In the Appalachian Mountain streams several species occur among the bryophytes, including L. cf. macrocera (Glime 1968). Limnophila alleni (see Figure 33) lays its eggs

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among mosses (Alexander 1919). Lauga and Thomas (1978) found that Limoniidae in France were more likely to be found among bryophytes when it was time for pupation and molting. The same relationship was seen for members of Athericidae and Rhagionidae (Brachycera).

Figure 33. Limnophila larva, member of a genus known to lay eggs in mosses. Photo by Tom Murray, through Creative Commons.

Erioptera (Figure 34), Pseudolimnophila (Figure 35), and Pilaria (Figure 36) in Wisconsin, USA, use mosses among their larval substrata (Hilsenhoff 1975). Byers (1961) reported that the larvae of Erioptera and Gonomyia (Figure 37) use bryophytes as larval habitats. In the Appalachian Mountain streams (USA), one can find the genus Antocha (Figure 7) (Glime 1968), a genus found in similar habitats in Japan.

Figure 36. Pilaria sp. larva breathing apparatus, a genus that lives among Wisconsin mosses. Photo by Urmas Kruus, with permission.

Figure 37. Gonomyia adult, a genus whose larvae live among bryophytes. Photo by Joe Zito, through Creative Commons.

Figure 34. Erioptera sp. larva, a moss inhabitant. Photo courtesy of the State Hygienic Laboratory at the University of Iowa, with permission.

Figure 35. Pseudolimnophila sp. larva breathing apparatus, a genus that lives among Wisconsin mosses. Photo by Urmas Kruus, with permission.

Blanket bogs have their own fauna, some of which is unique. Larvae that live in these habitats in Dartmoor, UK, include Molophilus occultus (Figure 38) whose larvae seem to require areas of bare, wet peat where they live in litter and among mosses (Boyce 2011). But this genus can also be found among bryophytes in Appalachian Mountain, USA, streams (Glime 1968). Phylidorea squalens (Figure 39) larvae in the Dartmoor blanket bogs live in the bog pools.

Figure 38. Molophilus sp. larva, a larva that seems to require bare, wet peat. Photo by Erin Hayes-Pontius, through Creative Commons.

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may not eat. That's right, they are not giant mosquitoes and won't bite you! But they do look like giant mosquitoes, with long legs and bodies 7-35 mm long (Tipulidae 2014), but narrow. Unlike the Limoniidae, the Tipulidae (Figure 42) are mostly terrestrial. Their larval food choices include algae, microflora, and both living and decomposing plant matter, including wood.

Figure 39. Phylidorea squalens adult male, a species whose larvae live in bog pools. Photo by James K. Lindsey, with permission.

Pediciidae – Hairy-eyed Craneflies The Pediciidae occur in the temperate zones of both hemispheres (Kits 2005b). These are medium to large (2035 mm) flies (Pediciidae 2014) that resemble craneflies. Pedicia (Figure 40) (now placed in Pediciidae) is one of the craneflies found among mosses as larvae (Figure 41) in some streams in the Appalachian Mountains, USA (Glime 1968). Hilsenhoff (1975) reported the genus in Wisconsin, USA, where it includes mosses among its substrata.

Figure 42. The cranefly Tipula occurs frequently among leaf litter that it helps to shred by eating it, but it can also occur among submerged and moist moss clones where its ecological role is unknown. Photo by Janice Glime.

The Tipulidae accomplish most of their respiration by using a posterior respiratory apparatus (Figure 43-Figure 44) (Pritchard 1983). They have a single pair of spiracles located there. The spiracles can't be closed, but there are tiny hairs on the walls of the spiracle opening that reduces water loss. There also seems to be cuticular respiration.

Figure 43. Larva of Tipula showing respiratory apparatus at right. Photo from Beentree, through Creative Commons. Figure 40. Pedicia rivosa adult on Equisetum. Larvae of some species live among mosses in Appalachian Mountain streams. Photo by Niels Sloth, with permission.

Figure 41. Pedicia albivitta larva, member of a genus of moss dwellers. Photo by Jason Neuswanger, with permission.

Tipulidae – Craneflies This is a worldwide family that occupies a wide range of habitats as larvae, from water to mosses to dry logs (Hofsvang 1997). As adults they live only a few days and

Figure 44. Respiratory apparatus with spiracles of Tipula sp. Photo from Beentree, through Creative Commons.

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Egg-laying (Figure 45) of tipulids on bryophytes has been known for a long time. For example, Alexander (1919) reported that Tipula nobilis laid her eggs in moss. Females already have mature eggs when they emerge from the pupa and after copulation they deposit them on wet soil or algae, or drop them (Tipulidae 2014). These eggs are usually black and may have a thin thread that could help to attach them in the water.

(Stern & Stern 1969). Slightly farther north in the Appalachian Mountains, I found what appeared to be seven different species of Tipula among bryophytes in the 28 streams I studied, including Tipula collaris (Figure 47) (Glime 1968). At Barrow, Alaska, USA, Tipula carinifrons (Figure 48) is common in the dry moss hummocks (MacLean 1980).

Figure 45. Cranefly laying eggs in submerged mosses. Photo by Janice Glime.

Tipulidae adults look like giant mosquitoes because of their long legs (Figure 46). In some regions they are known as daddy-long-legs for the same reason, but these are not to be confused with the 8-legged daddy-long-legs that are arachnids. Many Tipulidae live among aquatic leaf litter and mosses as larvae. Likewise, most of them pupate in soil near water, in mosses, or in litter (Byers 1978, 1996; Erman 1984).

Figure 47. Tipula collaris adult, a species whose larvae live among bryophytes in Appalachian Mountain streams. Photo through Carnegie Museum of Natural History, through Creative Commons.

Figure 46. Tipula adult. Photo by Micka 972, through Creative Commons in .

Larvae of craneflies are highly susceptible to desiccation (Pritchard 1983) and bryophytes seem to be an important habitat for maintaining moisture in bog species and terrestrial species. Tipula montana burrows into mosses when it is disturbed (Smith et al. 2001). Dolichopeza (Figure 77) species select their moss habitat for its suitability for making burrows (Byers 1961). The cranefly larvae seem to prefer compact mosses rather than loose ones in the same species (Todd 1993). Tipula ignobilis occurs throughout the year among mosses on boulders in a Tennessee, USA, springbrook

Figure 48. Tipula carnifrons adult male, a common species in dry moss hummocks of Alaska. Photo by Ashley Bradford, through Creative Commons.

Byers (1961) listed bryophytes as the larval habitat of many Tipula species. The genus Tipula is typically a consumer of leaf litter. But mosses can be a major part of the diet in some species. Dangles (2002) found that in the four study streams of Vosges Mountains in northeastern France bryophytes comprised 96% of the diet of Tipula (Savtshenkia) (Figure 49).

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lanuginosum to the other mosses and often avoided Pleurozium schreberi when given a choice (Sphagnum girgensohnii was the least preferred). This avoidance of Pleurozium schreberi is likely because of the high phenolic content (compounds that taste bad, including tannic acid) of P. schreberi (Liao 1993; Glime 2006; Hribljan 2009; see chapter 10-3 on Isopoda in this volume).

Figure 49. Tipula (Savtshenkia) adult, a genus in which the larvae can eat considerable amounts of bryophytes. Photo by James K. Lindsey, with permission.

Tipulidae larvae commonly feed on mosses (Coulson 1962; Freeman 1967; MacLean 1980; Richardson 1981; Todd 1993), and these mosses often form a significant portion of the diet (Coulson 1962). Larvae of Tipula signata (Figure 50) feed on aquatic mosses (Hemmingsen 1965).

Figure 51. Racomitrium lanuginosum, a preferred food for Tipula montana. Photo by Michael Lüth, with permission.

Figure 50. Tipula signata adult male, a species whose larvae eat aquatic mosses. Photo by James K. Lindsey, with permission.

Tipula montana is a bog dweller and is surrounded by bryophytes as a larva. Smith et al. (2001) experimented with food preference in larvae of this species. The research team gave the larvae trials with five individual species of mosses, then with two-species pairs, to determine their growth responses and preferences. Larvae grew on diets of each of the five species of mosses [Racomitrium lanuginosum (Figure 51), Dicranum fuscescens (Figure 52), Sphagnum girgensohnii (Figure 53), Pleurozium schreberi (Figure 54), and Polytrichum commune (Figure 55)], but there was a wide range in which mean weights differed by a factor of two. The highest development rate, by far, was for larvae fed Pleurozium schreberi, with nearly 50% reaching the fourth instar, whereas fewer than 5% of those fed on the other moss species reached that stage (Figure 56). Pleurozium schreberi also was the best moss for promoting growth, with weight gain double that of larvae fed on Sphagnum girgensohnii (Figure 57). Nevertheless, there was little difference among the survivorships of the larvae fed on each on the five mosses (Figure 58). But the larvae preferred Racomitrium

Figure 52. Dicranum fuscescens, a moss with a high relative percentage of observations of being eaten by Tipula montana. Photo by Michael Lüth, with permission.

Figure 53. Sphagnum girgensohnii, the least preferred moss among choices given to Tipula montana. Photo by Michael Lüth, with permission.

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Figure 54. Pleurozium schreberi a moss that gives Tipula montana good growth performance but that is not preferred. Photo by Janice Glime.

Figure 57. Mean fresh weight (+ standard error) of larvae of Tipula montana, starting with second-instars, after 52 days on each of five moss species. Sample sizes appear above bars. Redrawn from Smith et al. 2001.

Figure 55. Polytrichum commune, a potential food avoided by Tipula montana. Photo by Michael Lüth, with permission. Figure 58. Percent survival of Tipula montana larvae fed on each of five moss species for 52 days. Sample sizes appear above bars. Redrawn from Smith et al. 2001.

Figure 56. Survival percentages of Tipula montana larvae, starting with second-instar larvae, entering fourth instar after 52 days of feeding on diets of five moss species. Sample sizes appear above bars. Redrawn from Smith et al. 2001.

Smith et al. (2001) issued a note of caution: The fecal indications of moss herbivory did not match the observational data. They suggested this may have been due to behavior differences between the larvae and the observers. The observers noted feeding behavior between 8:30 hours and 19:30 hours, but the larvae may have been feeding actively above ground at night, with daylight causing them to avoid the greater exposure on the sedge Carex bigelowii. This could explain the estimated lower percentage of Carex bigelowii in the observed diet in the field when using observations, and accounting for the higher percentage of Dicranum fuscescens (Figure 52) in the observations when compared to the ratio in the feces. Ratios of other mosses were similar using both methods. In the field, when Carex bigelowii was readily available, it was the clear choice compared to the mosses. The researchers also concluded that the bryophytes may be more important as a refuge than as a food source in nature. As pointed out by the researchers, experiments in which

Chapter 11-3a: Aquatic Insects: Holometabola – Diptera, Suborder Nematocera

development and growth on the sedge compared to those of the mosses would be instructive. It may be that the best growth is on a combination of these, with reduced growth or development resulting when no mosses are eaten. On the other hand, avoidance of predators may force the larvae to remain among the mosses and to eat them in the daytime. Several birds are primary predators on these larvae (Galbraith et al. 1993; Nethersole-Thompson 1966). Tipula subnodicornis (Figure 59) feeds on liverworts in British moorland blanket bogs and consumes large quantities of Sphagnum (Figure 53, Figure 69) leaves (Coulson 1962; MacLean 1980). MacLean estimates that more than 25% of the energy consumption may be derived from the living plants of Sphagnum.

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burned Calluna heath. Tipula montana in the upland moors feeds exclusively on mosses.

Figure 60. Tipula confusa adult; larvae eat mosses, preferring woodland species. Photo by Malcolm Storey, through Creative Commons .

Figure 59. Tipula subnodicornis adult, a cranefly whose larvae feed on liverworts in British blanket bogs but seem to have little preference in experiments with moss species. Photo by James K. Lindsey, with permission.

In the genus Tipula, later instars ingest only slightly more vegetable matter as they grow to larger and larger instars. Rather, the early and late instars ingest similarsized particles. In feeding experiments, Todd (1993) found that Tipula confusa (Figure 60) preferred woodland moss species, whereas T. subnodicornis (Figure 59) showed no preference between woodland and moorland mosses. Tipula confusa had a hierarchical preference among the 10 moss species offered, whereas T. subnodicornis showed much less hierarchy in food choices. Brindle (1960) noted that T. subnodicornis (Figure 59) typically associates with wet species such as those of Sphagnum (Figure 69) and Hypnum (Figure 23) in moorlands. Among 11 species Todd (1993) studied, 8 were moss consumers, with 7 of these in the same subgenus Savtshenkia (Tipula rufina (Figure 61), T. confusa, T. pagana (Figure 62), T. staegeri, T. limbata (Figure 63), T. alpium (Figure 64), and T. subnodicornis). Brindle (1960) had earlier observed that all the moss feeders known to him had four pairs of short anal papillae, whereas in wetter environments these papillae were longer. The eighth, T. montana is in the subgenus Vestiplex. In Great Britain, approximately onefourth of the 59 (Freeman 1967) members of Tipula feed on mosses. Even the invasive species Campylopus introflexus (Figure 65) is Tipula food in the recently

Figure 61. Tipula rufina adult, a species whose larvae eat small particle sizes of bryophytes. Photo by Malcolm Storey, through Creative Commons .

Figure 62. Tipula pagana male adult, a species whose larvae eat small bites of bryophytes. Photo by James K. Lindsey, with permission.

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Figure 63. Tipula limbata adult, a species whose larvae eat bryophytes in small bites. Photo by Derek Sikes, University of Alaska Museum, through Creative Commons.

The insect feces (excrement; waste material discharged from gut) reveal a great deal about the use of mosses as food (Todd 1993). The particle size remains the same in the feces as it was in the cut ingested portion (Pritchard 1983). Interior cells of the pieces are significantly less damaged (Todd 1993). Instead, digestion appears to be limited to the broken cells on the edges, with little or no damage caused by passage through the gut. This inability to obtain nutrients from the interior cells accounts for the consistency in small-sized particles from early to late instars. The particle sizes are significantly smaller for Tipula rufina (Figure 61), T. lateralis (Figure 66), and T. subnodicornis (Figure 59); T. paludosa (Figure 67) and T. oleracea (Figure 68) ingest significantly larger particles than any other species. These differences are at least partly explained by mandible size. Tipula paludosa has significantly larger mandibles and T. rufina has significantly smaller ones than any other species. In short, those species feeding on grass are generally larger and have longer mandibles than those species feeding on mosses.

Figure 64. Tipula alpium adult, a species whose larvae eat bryophytes in small bites. Photo by Malcolm Storey, through Creative Commons.

Figure 66. Tipula lateralis adult, a species whose larvae ingest small particle sizes. Photo by James K. Lindsey, with permission.

Figure 65. Campylopus introflexus, an invasive species that has become a food source for Tipula larvae in the Calluna heath. Photo by Michael Lüth, with permission.

Figure 67. Tipula paludosa larva, a bryophyte consumer. Photo by Roger S. Key, with permission.

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food, as Bryum (Figure 72) sp. and several seedlings were untouched.

Figure 68. Tipula oleracea, a bryophyte consumer that ingests large particles. Photo by Malcolm Storey, through Creative Commons .

Tipula has both terrestrial and aquatic members. Some of these in both habitats consume bryophytes. But Tipula subnodicornis (Figure 59) prefers the cottongrass Eriophorum vaginatum to the terrestrial moss Campylopus paradoxus and bog moss Sphagnum papillosum (Figure 69) (Todd 1993). However, in early winter (10 December to 9 January) the preference changes significantly from cottongrass to Sphagnum papillosum. It is interesting, however, that during the growing season there is a mix of Eriophorum vaginatum with S. papillosum where the larvae spend the most time.

Figure 70. Anthoceros agrestis, food source for Tipula larvae. Photo by Jan-Peter Frahm, with permission.

Figure 71. Phaeoceros carolinianus, food source for Tipula larvae. Photo by Michael Lüth, with permission.

Figure 69. Sphagnum papillosum, a moss that becomes a preferred food in winter for Tipula subnodicornis. Photo by Michael Lüth, with permission.

Bisang (1996) reports a rather bizarre experience in The Bryological Times. She had several cultures of Anthoceros agrestis (Figure 70) and Phaeoceros carolinianus (Figure 71), both hornworts. Using the same techniques as she had used previously, she cultured these in jars, keeping two in Switzerland and taking one to Sweden. To her surprise, one of the cultures in Switzerland and the one taken to Sweden virtually disappeared from the jar. They had not dried and sabotage seemed absurd. Careful examination revealed larvae 1.5 cm long with a breathing apparatus at the posterior end. The cultures were supporting a healthy colony of larvae of Tipula (Figure 42), craneflies. The hornworts seemed to be a preferred

Figure 72. Bryum capillare. A species of Bryum was refused as food by larvae of a species of Tipula. Photo by Aimon Niklasson, with permission.

The members of Tipula are among the few documented moss consumers, although there is much more consumption than is generally recognized. Todd (1993) suggested that the presence of cell wall bioflavonoids in bryophytes might function not only to resist fungal invasion (Geiger 1990), but also to discourage insect

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browsers. It is also possible that in some cases the fungi are needed to facilitate digestion, making mosses that lack them indigestible. Furthermore, lignin-like compounds in the bryophyte cell walls protect the cell wall compounds (cellulose, hemicellulose, and other kinds of polysaccharides) from hydrolytic attack (using a chemical reaction where something reacts with water and is changed into a new substance), preventing the consumers from using hydrolytic attack to extract cell contents, as demonstrated in Tipula abdominalis (Figure 75) (Martin et al. 1980). Nevertheless, in North America the genus Tipula (Figure 75) is able to hydrolyze proteins from unconditioned maple (Acer) leaves (Barlocher & Porter 1986). Suitable food sources often depend on pH of the gut (Martin et al. 1980). Very high and very low pH levels seem to work best. But Barlocher and Porter (1986) found that the larvae of Tipula caloptera (Figure 73) have a gut pH that is somewhat alkaline. Fungal carbohydrases ingested with the leaves do not remain active in the T. caloptera gut, but do in the nearly neutral pH of the amphipod Gammarus tigrinus and net-spinning caddis larva Hydropsyche betteni (Figure 74).

Figure 73. Tipula caloptera adult female. Larvae of this species have an alkaline gut that may help it digest plant material. Photo by Tom Murray, through Creative Commons.

Figure 74. Hydropsyche betteni larva, a species with a slightly alkaline gut and ability to keep fungal enzymes alive. Photo by Donald S. Chandler, with permission.

In Tipula abdominalis (Figure 75) the midgut has a pH near 11.5 in a narrow section where there is extremely high proteolytic activity (Martin et al. 1980). In addition to low pH created by Sphagnum (Figure 69) and other mosses, mosses are well known for their antibiotics (McCleary et al. 1960; McCleary & Walkington 1966), additional factors that might interfere with gut digestion.

Figure 75. Tipula abdominalis larva. Larvae have a high pH in the midgut. Photo by Tom Murray, through Creative Commons.

Dolichopeza (Figure 77) is a genus known from mosses in various parts of the world. Dolichopeza americana is generally considered to be a terrestrial larva (Byers pers. comm.), but in the Appalachian Mountain streams it occurs among the leafy liverworts (Scapania undulata; Figure 30) in small waterfalls in March and December (Glime 1968). Dolichopeza albipes (see Figure 77) is a white-footed ghost cranefly whose larvae live among the mosses and liverworts of the Ghyll woodlands in Sussex, UK (Roper 2001). But this genus also chooses mosses for home in South Africa (Harrison & Barnard 1972). Members of this genus are known to lay their eggs among bryophytes, giving these larvae their start in life among the bryophytes. Dolichopeza barnardi, D. hirtipennis, and D. peringueyi larvae live beneath and within cushions of wet mosses and liverworts at the sides of waterfalls in South Africa (Harrison & Barnard 1972). And in North America, the genus feeds on terrestrial mosses (Byers 1961). In the coastal tundra near Barrow, Alaska, Prionocera recta (Figure 76) is restricted to mossy depressions.

Figure 76. Prionocera turcica adult, relative of P. recta restricted to mossy depressions in the Alaskan tundra. Photo by Andre Vrigens, through Creative Commons.

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Figure 78. Philonotis fontana similar to seepage area where a member of Anisopodidae was eating and defecating bits of moss. Photo by Michael Lüth, with permission.

Figure 77. Dolichopeza carolus adult. Larvae of several species in this genus live among mosses, including at the sides of waterfalls. Photo by Tom Murray, through Creative Commons.

Many of the Tipulidae that inhabit mosses as larvae do so among terrestrial bryophytes and will be discussed in a separate chapter on Terrestrial Insects. Anisopodidae – Wood Gnats, Window Gnats This family is worldwide, but bryophytes are not a usual habitat. Most are small (4-12 mm) (Anisopodidae 2014). Fungi are typical foods, but it appears that at least some feed on micro-organisms, as I have observed. While looking for mosses one day, I found some (Philonotis fontana?; Figure 78) in a seepage area on a cliffside. There on one of its branches was a small larva eating away at the wet moss. But as I watched for awhile, I realized that the mosses were going into one end of the larva covered with detritus and coming out the other end clean and still bright green. I was unable to identify this single larva beyond family. The larvae of Sylvicola cinctus (Figure 79) was reported from mosses in Norway (Søli 1992). Perhaps there are other members of this small family hiding among the bryophytes.

Figure 79. Silvicola cinctus male adult, a species whose larvae live among bryophytes in Norway. Photo by Walter Pfliegler, with permission.

I have seen only one record from this little-known family. Axymyia furcata (Figure 80) is a semi-aquatic fly in its larval stage and is typically a wood inhabitant. However, Wihlm and Courtney (2011) found that the larvae often choose logs that are covered with mosses.

Axymyiidae This is a small family of six known species (Axymyiidae 2014). Its limited distribution is Holarctic and Oriental (Hauser 2008). The larvae live in decomposing wood (Axymyiidae 2014).

Figure 80. Axymyia furcata, a semi-aquatic larva that lives among mosses on logs. Photo by M. J. Hatfield, through Creative Commons.

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Cecidomyiidae – Gall Midges, Gall Gnats This family is worldwide with most records in the Northern Hemisphere. They are small flies, mostly 1-5 mm (Balaban & Balaban 2004). Most of these are gall makers, with their larvae living on the gall material, but some feed on plants and some on decaying matter. Hence, as one might expect, they are predominantly terrestrial, but there are aquatic exceptions. Although the Cecidomyiidae (Figure 81) are not typical bryophyte inhabitants, some do prefer mosses in torrents (Thomas 1980). Porricondyla ramadei was described as a new species from tufts of mosses in the turbulent waters of high Pyrénées streams. This is a poorly known fauna, and it is likely more insects may be discovered among the bryophytes there.

Because they live among litter and fungi, they are frequent in flower pots. They are small, up to 7 mm long.

Figure 82. Gnoriste sp. adult; larvae of Gnoriste apicalis live in saturated mosses. Photo from Biodiversity Institute of Ontario, through Creative Commons.

In Korea, Japan, China, and other parts of Asia, the shiitake mushroom business is important. To this end, studies on the pests of this delicacy are common. And sometimes we find that mosses are involved. Shin et al. (2012) found that one of the mushroom pests, Bradysia difformis (Figure 83), also occurs in moorland on peat moss.

Figure 81. Cecidomyiidae larva; some members of this family live among mosses in torrents. Photo by M. J. Hatfield, through Creative Commons.

Mycetophilidae – Fungus Gnats As the name implies, these flies live among fungi, hence making them most common in damp or sometimes wet habitats (Mycetophilidae 2014). They are worldwide, especially in forested areas (Kits 2005a). Although they are worldwide, most records are in the Northern Hemisphere (Mycetophilidae 2015). They typically feed on the fruiting bodies of the fungi (Mycetophilidae 2014). But some live among mosses and liverworts. Fungi are often moist, so it may not be so surprising that some of these fungus gnats have found bryophytes to be suitable habitats. Gnoriste apicalis (Figure 82) is a semi-aquatic species. The larvae are able to live in saturated moss clumps on lake shores (Lenz 1927; Johannsen 1969). The pale green coloring may help it to be inconspicuous as it feeds on detritus. It may also make a dense but delicate white web in which it lives in such habitats, with the web offering further camouflage. Sciaridae – Dark-winged Fungus Gnats As you might expect of a fungus gnat, these flies prefer moist sites and eat the fruiting bodies of mushrooms and various parts of other fungi (Sciaridae 2014). They are worldwide in distribution, including such extremes as deserts, sub-Antarctic islands, and altitudes over 4000 m.

Figure 83. Bradysia difformis, a shiitake mushroom pest whose larvae sometimes live on peat mosses of moorlands. Photo by David Pilling, with permission.

Ceratopogonidae – Biting Midges, No-see-ums, Sand Flies, Punkies Their small size (16 hr to 15%), indicating their use of animal-derived N through repeated trophic transfer by microbial activity since the original deposition. Figure 36. Falco peregrinus, Peregrine Falcon and guano on cliff edge where it perches. Photo by Mike Baird through Creative Commons.

But guano does not always favor the mosses. In the polar Mac. Robertson Land, guano has reached toxic levels, making the coastal slopes barren of mosses and lichens (Bergstrom & Seppelt 1990). This is largely due to Antarctic Petrels (Thalassoica antarctica; Figure 37) that breed along these slopes, with a mean nest density of 0.82 mˉ1 (Alonso et al. 1987)! But the area also serves as breeding grounds for Southern Fulmars (Fulmarus glacialoides; Figure 38) and Adélie Penguins (Pygoscelis adeliae; Figure 39). Figure 39. Pygoscelis adeliae, Adelie Penguin on Antarctica, illustrating the large number of birds that can create guano. Photo by Murray Foubister, with permission

Figure 37. Thalassoica antarctica, Antarctic Petrel flying. Photo by François Guerraz, through Creative Commons.

In the case of the Adelie Penguin (Pygoscelis adeliae; Figure 39), dung left 3000-8000 years ago remains, at least partly frozen in ice (Gill 2012). Mosses are able to derive nutrients from these deposits, giving them much needed resources that are so scarce in the sand and gravel substrate of Antarctica. Penguin rookeries on King George Island in the maritime Antarctic are an important source of nutrients and have a strong influence on the vegetation patterns and diversity (Smykla et al. 2007). The nutrient input, as guano, creates a zonation pattern. The first zone includes those areas under the immediate influence of fresh guano and trampling, supporting little or no vegetation. The second zone is adjacent to the first and is covered with nitrogen-loving green algae and sometimes Cyanobacteria. The third zone is dominated by Antarctic hair-grass. The fourth zone is dominated by mosses. The fifth and last zone under the rookery influence is dominated by lichens.

Peatland Habitats

Figure 38. Fulmarus glacialoides, Antarctic Fulmar roosting; their guano prevents establishment of bryophytes. Photo by Samuel Blanc, through Creative Commons.

Brewer (1967) pointed out that studies on bog vegetation were much more numerous than those on the animal populations. To help remedy this situation, he studied the breeding bird populations on two peatlands in lower Michigan. In the years 1961-1966 he noted 24 species of breeding birds in Portage Bog. These included the Song Sparrow (Melospiza melodia; Figure 40), Field Sparrow (Spizella pusilla; Figure 41), Yellowthroat (Geothlypis trichas; Figure 42), Yellow Warbler (Setophaga petechia; Figure 43), Nashville Warbler (Leiothlypis ruficapilla; Figure 44), Eastern Towhee

Chapter 16-1: Birds

(Pipilo erythrophthalmus; Figure 45), Brown-headed Cowbird (Molothrus ater; Figure 46), Catbird (Dumetella carolinensis; Figure 47), American Goldfinch (Carduelis tristis; Figure 48), Traill's Flycatcher (Empidonax traillii; Figure 49), Black-capped Chickadee (Poecile atricapillus; Figure 50), Mourning Dove (Zenaida macroura; Figure 51), Cedar Waxwing (Bombycilla cedrorum; Figure 52), Yellow-shafted Flicker (Colaptes auratus; Figure 53), Cardinal (Cardinalis cardinalis; Figure 54), Brown Thrasher (Toxostoma rufum; Figure 55), Ruby-throated Hummingbird (Archilochus colubris; Figure 56), Mallard (Anas platyrhynchos; Figure 57), Marsh Hawk (Circus cyaneus), Eastern Bluebird (Sialia sialis; Figure 58), Tree Swallow (Tachycineta bicolor; Figure 59), Robin (Turdus migratorius; Figure 60), Whip-poor-will (Caprimulgus vociferus; Figure 61), and Veery (Catharus fuscescens; Figure 62). Among these, the Mallards were the only species for which the researchers located a nest, and the nest occurred in three of the six years. About 425 pairs were located there per hectare. Brown-headed Cowbirds were the most dense and Song Sparrows were the most abundant, the latter having an average of 138 territorial males per hectare. Others with a density of more than 24 per hectare were Yellowthroats, Field Sparrows, Eastern Towhees, and, perhaps, Brown-headed Cowbirds.

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Figure 42. Geothlypis trichas, Yellowthroat, a species that commonly occurs in bogs during breeding season. Photo by Dan Pancamo, through Creative Commons.

Figure 43. Setophaga petechia, Yellow Warbler, a species that commonly occurs in bogs during breeding season. Photo by Dick Daniels, through Creative Commons.

Figure 40. Melospiza melodia, Song Sparrow, a species that commonly occurs in bogs during breeding season. Photo by Len Blumin, through Creative Commons.

Figure 41. Spizella pusilla, Field Sparrow, a species that commonly occurs in bogs during breeding season. Photo by Jeff Whitlock, through Creative Commons.

Figure 44. Leiothlypis ruficapilla, Nashville Warbler, a species that commonly occurs in bogs during breeding season. Photo by Jerry Oldeneffel, through Creative Commons.

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Figure 45. Pipilo erythrophthalmus, Eastern Towhee, a species that commonly occurs in bogs during breeding season. Photo by Ken Thomas, through Creative Commons.

Figure 48. Carduelis tristis, American Goldfinch, a species that commonly occurs in bogs during breeding season. Photo by MDF, through Creative Commons.

Figure 46. Molothrus ater, Brown-headed Cowbird, a species that commonly occurs in bogs during breeding season. Photo through Creative Commons.

Figure 49. Empidonax traillii, Willow Flycatcher, a species that commonly occurs in bogs during breeding season. Photo by Dominic Sherony, through Creative Commons.

Figure 47. Dumetella carolinensis, Grey Catbird, a species that commonly occurs in bogs during breeding season. Photo by Steve, through Creative Commons.

Figure 50. Poecile atricapillus, Black-capped Chickadee, a species that commonly occurs in bogs during breeding season. Photo by Zac Cota, through Creative Commons.

Chapter 16-1: Birds

Figure 51. Zenaida macroura, Mourning Dove, a species that commonly occurs in bogs during breeding season. Photo by R. L. Sivaprasad, through Creative Commons.

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Figure 54. Cardinalis cardinalis, Cardinal in snow in Pickerington, OH, a species that commonly occurs in bogs during breeding season. Photo courtesy of Eileen Dumire.

Figure 52. Bombycilla cedrorum, Cedar Waxwing, a species that commonly occurs in bogs during breeding season. Photo by Cephas, through Creative Commons. Figure 55. Toxostoma rufum, Brown Thrasher, a species that commonly occurs in bogs during breeding season. Photo by E. Monk, through Creative Commons.

Figure 53. Colaptes auratus, Yellow-shafted Flicker, a species that commonly occurs in bogs during breeding season. Photo by Minette Layne through Creative Commons.

Figure 56. Archilochus colubris, Ruby-throated Hummingbird, a species that commonly occurs in bogs during breeding season. Photo by Dan Pancamo, through Creative Commons.

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Figure 57. Anas platyrhynchos female (left) and male (right), a species that commonly breeds and nests in bogs. Photo by Richard Bartz through Wikimedia Commons. Figure 60. Turdus migratorius, Robin, a species that commonly occurs in bogs during breeding season. Photo by Dakota Lynch, through Creative Commons.

Figure 61. Caprimulgus vociferus, Whip-poor-will, a species that commonly occurs in bogs during breeding season. Photo by Jerry Oldeneffel, through Creative Commons. Figure 58. Sialia sialis, Bluebird male, a species that commonly breeds in bogs. Photo from Sandy's Photos 2009, through Creative Commons.

Figure 62. Catharus fuscescens, Veery, a species that commonly occurs in bogs during breeding season. Photo by Dominic Sherony, through Creative Commons.

Figure 59. Tachycineta bicolor, Tree Swallow, a species that commonly occurs in bogs during breeding season. Photo by John Benson, through Creative Commons.

In bogs studied by Brewer (1967), as the high thicket gave way to low thicket, some of the bird species changed, including the arrival of the Nashville Warbler (Leiothlypis ruficapilla; Figure 44) in 1965. The trees in the bog were not suitable for cavity-nesting birds during the study. Among these birds, Field Sparrows (Spizella pusilla; Figure 41) preferred open bog and Song Sparrows

Chapter 16-1: Birds

(Melospiza melodia; Figure 40) preferred thickets, as did the Towhee (Pipilo erythrophthalmus; Figure 45), Yellowthroat (Geothlypis trichas; Figure 42), and Catbird (Dumetella carolinensis; Figure 47). The number of species in the open bog was about 13, whereas in the thicket it was about 21. When examining peatlands on a larger scale, Niemi and Hanowski (1992) found 110 species of birds that frequented Minnesota peatlands. Brewer (1967) concluded that most of the birds came to the bog only for feeding. For example, Robins (Turdus migratorius; Figure 60) nested in the deciduous areas but came to the bog for feeding. This was especially true when berries were ripe, with both juveniles and adults coming to feed. Based on these habitat relationships, it is not surprising that most of the species in this bog were forest edge species. Brewer also considered it likely that some of the visitors, like the Meadowlark (Sturnella magna; Figure 63), mistook the open bog for an open field.

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This isolation causes the peatlands and their bird populations to behave with island dynamics. Among ten species of birds studied in detail, two rely primarily on peatlands for nesting sites. Bird species richness was primarily related to microhabitat richness and heterogeneity. The Palm Warbler (Dendroica palmarum; Figure 64) and Upland Sandpipers (Bartramia longicauda; Figure 65) depended on having larger, non-isolated peatlands.

Figure 64. Dendroica palmarum, Palm Warbler, a species that depends on large, non-isolated peatlands. Photo by Wolfgang Wander, through Creative Commons.

Figure 63. Sturnella magna, Eastern Meadowlark, a bird that may occasionally mistake an open bog or fen for an open field. Photo by Jim F. Bleak, through Creative Commons.

Brewer (1967) only observed birds in the Sugarloaf Bog for two years. This site had 26 breeding bird species during that time, with the average per year of about 20 species. The density was high, with about 675 males per hectare. The Black-capped Chickadee (Poecile atricapillus; Figure 50) was the most abundant, with about 100 males per hectare (compared to 10 at Portage Bog). Only nine species were common to both locations (Brewer 1967). In a larger study based on literature, Brewer found that there is little commonality among species of the open bog. Birds of the spruce forest, on the other hand, are similar to those of a cedar forest or a spruce thicket. It became clear that species of the bogs depended on the vegetation of that stand and on the vegetation of adjacent areas, as well as the geographic distribution of the species. Few birds were present in the winter, reflecting the poor winter food supply and insufficient cover. Calmé and Desrochers (1999, 2000) and Calmé et al. (2002) investigated the birds in 67 southern Quebec, Canada, peatlands. They expressed concern over the loss of peatlands to urban sprawl, agriculture, forestry, and peat mining, particularly in eastern Canada (Calmé & Desrochers 2000). This loss further fragments the peatlands, making natural re-introductions more difficult.

Figure 65. Bartramia longicauda, Upland Sandpiper, a species that depends on large, non-isolated peatlands. Photo by Johnath, through Creative Commons.

Calmé et al. (2002) found 17 species of birds that were significantly more frequent in peatlands than in the surrounding habitats. For some, the peatland was one of several habitats, but some were significantly more frequent in peatlands. In studying 28 southeastern Quebec, Canada, peatlands, Desrochers et al. (1998) found that harvesting

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effects on birds depended on the type of harvesting. Block harvesting had the least effect, presumably because it retained most of the topography and microhabitats. Vacuum harvesting, on the other hand, did alter the bird communities. Ten of the 28 species responded negatively to peatland perturbation. The Palm Warbler (Dendroica palmarum; Figure 64), in particular, was closely associated with the unperturbed sites. The Palm Warbler (Dendroica palmarum; Figure 64) is an area-sensitive bird and in southern Québec it is restricted to peatlands (Poulin 2002). The within-site habitat configuration strongly affects the physical efficiency of this species but not necessarily functional effectiveness. While it is clear that having a number of peatlands available is important to the Palm Warbler, the biological factors they provide remain elusive. When Lachance et al. (2005) investigated 16 peatlands in southern Quebec, Canada, they found 36 bird species and 154 plant species. They found that afforestation altered the vegetation structure in ways that changed the bird species composition. In particular, there were fewer mosses and shrubs, but more trees. One reason for the diminished number of birds in disturbed peatlands is the loss of eggs and nestlings to predation. Haddad et al. (2000) assessed the effects of harvesting peat mosses on the survival of bog-dwelling songbirds [Palm Warbler (Dendroica palmarum; Figure 64), Common Yellowthroat (Geothlypis trichas; Figure 42), Hermit Thrush (Catharus guttatus; Figure 66), and several species of sparrows (Passeridae; Figure 40-Figure 41)]. They found greater risk of nest predation in harvested bogs.

Figure 67. Tipula, leatherjacket larva, a genus that is eaten in great numbers by birds in bogs. Photo by Rasbak, through Creative Commons.

Effects on Structure

Bryophyte

Community

Birds can have considerable influence on bryophyte communities, especially in Arctic wetlands. We have already seen that guano from seabirds can provide nutrients that are otherwise limiting. And Pheasants (Figure 68) can disrupt the community while searching for food (Erkamo 1976).

Figure 68. Phasianus colchicus, Pheasant, a forager that can disturb bryophytes while foraging. Photo by Hugh J. Griffiths, through Creative Commons.

In the Arctic, geese (Figure 69) can play a role in community structure (Jasmin et al. 2008). Although one might expect such feeding disruption to reduce the number of species, Jasmin and coworkers found greater bryophyte species richness following 11 years of goose presence, compared to that in goose exclosures. The non-protected areas exhibited more variation in time and space than within the exclosures, promoting greater coexistence of bryophyte species at the microscale of 1 cm. Figure 66. Catharus guttatus, Hermit Thrush, a species that loses more eggs to predation in harvested bogs than in undisturbed bogs. Photo by Cephas, through Creative Commons.

Another possibility to explain loss of birds on harvested peatlands is disruption of the habitat of food organisms. Diptera larvae, especially the cranefly Tipula (Figure 67), live and pupate among the mosses in the peatland (MacLean 1980). The birds consume 35-70% of annual production of Tipula carinifrons and consume 50% of adults at peak emergence. The cranefly larvae feed on liverworts in these bogs (Coulson & Whittaker 1978). Paasivirta et al. (1988) likewise noted the importance of emerging insects for feeding birds in peatlands.

Figure 69. Chen caerulescens, migratory Snow Geese, foraging. Photo by Bradley Davis, through Creative Commons.

Chapter 16-1: Birds

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Conservation Issues Agricultural areas might actually help bird species diversity in tropical forests (Hughes et al. 2002; Sekercioglu et al. 2007). Although we typically think of deforestation for agriculture as being detrimental to bird diversity, researchers found that most of the 144 bird species used the agricultural areas for foraging, often travelling several kilometers from their forest home (Hughes et al. 2002). They estimated that 46% of the native birds were using the agricultural countryside in southern Costa Rica. The authors suggest that diversity will suffer less if tall trees and edge habitats are maintained. In an effort to understand how to protect birds with minimal effort, we have often chosen indictor species (Simberloff 1998). Unfortunately, these are not as indicative as we might hope. It is difficult to know what species should be the indicator and on just what it should indicate. Simberloff suggested instead that the species should be an "umbrella species,... one that needs such large tracts of habitat that saving it will automatically save many other species." A flagship species is typically a charismatic large vertebrate, such as the panda or a snowy owl (Anonymous, USDA; Simberloff 1998). It is useful because it causes both public interest and sympathy (Simberloff 1998). It suffers some of the same problems – it may not be in an area that protects many other species, and it might be expensive to protect. And management of one flagship species may conflict with that of managing another. "The recognition that some ecosystems have keystone species whose activities govern the well-being of many other species suggests an approach that may unite the best features of single-species and ecosystem management. If we can identify keystone species and the mechanisms that cause them to have such wide-ranging impacts, we would almost certainly derive information on the functioning of the entire ecosystem that would be useful in its management." Even keystone species can get complicated. As seen in a Colorado subalpine ecosystem, there may be subtle interdependencies (Daily et al. 1993). The Red-naped Sapsuckers (Sphyrapicus nuchalis; Figure 70) actually have two keystone roles. Their excavation activities to make nests in fungus-infected aspens are essential to two species of swallows, and when they drill sap wells into willows they nourish not only themselves, but also make this rich food source available to Hummingbirds (Figure 56), Orange-crowned Warblers (Vermivora celata; Figure 71), chipmunks (Tamias striatus), and other sap robbers. Thus for this community to persist, it requires the complex interactions of sapsuckers, willows, aspens, and a heartwood fungus. As an example, the penguin (Figure 39) can be a keystone species in the maritime Antarctic (Barcikowski et al. 2005). We have seen above that the guano produced by the penguins can form the base for an entire community by providing an important supplement to the rare nutrients. In areas where the guano enriches the substrate with nutrients originating in the ocean, the grasses Colobanthus quitensis (Figure 72) and Deschampsia antarctica (Figure 73) predominate. Where the guano is absent, mosses such as Polytrichum piliferum (Figure 74) predominate.

Figure 70. Sphyrapicus nuchalis, Red-naped Sapsucker, a keystone bird species. Photo by Dominic Sherony, through Creative Commons.

Figure 71. Vermivora celata, Orange-crowned Warbler, a species that depends on the Red-naped Sapsucker as a keystone species. Photo by Linda Tanner, through Creative Commons.

Figure 72. Colobanthus quitensis, a dominant Antarctic species in areas enriched by guano. Photo by John Clark, through Creative Commons.

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travel in the digestive system of birds (Behling et al. 2016). On Navarino Island, at the Cape Horn Biosphere Reserve, these researchers recovered bryophyte diaspores from fecal samples from the Upland Goose (Chloephaga picata; Figure 34) and the White-bellied Seedsnipe (Attagis malouinus). Viability remains to be established.

Figure 73. Deschampsia antarctica, a dominant Antarctic species in areas enriched by guano. Photo by John Clark, through Creative Commons.

Figure 75. Taeniopygia guttata, Zebra Finch, a pet that is an incessant nest builder and uses mosses, among other things. Photo from Sky High Butterfly, through Creative Commons.

Figure 74. Polytrichum piliferum, a moss that avoids areas with guano in the maritime Antarctic. Photo by Bob Klips, with permission.

To put this in a bryological perspective, we may find that a species is dependent on mosses in spring before herbaceous plants are available or in winter when tracheophytes cease growing. The bryophytes might depend on one or more species of birds for the bulk of their dispersal. Or the bryophytes might serve as emergency foods during years when the weather is not suitable for good productivity of other, more preferred foods. With so many possibilities, we have just begun to understand the interrelationships.

Davison (1976) describes the role of birds in the dispersal of mosses. Indeed, it was not the nest-building activities, but feeding activities that caught his attention. Where leaf litter is somewhat scarce, such as older beech woods, and mosses are abundant, foraging requires that the birds poke around among the mosses. Blackbirds (Turdus merula, Figure 76) in particular foraged among Mnium hornum (Figure 25) and Polytrichastrum formosum (Figure 77), breaking the plants and scattering them much like the Japanese do when planting a moss garden. Davison reports that within a two-month period these birds moved 34 clumps of moss from one place to another within an area of about 5 m2, but also brought to the area an additional 18 pieces.

Dispersal Agents If you have ever reared Zebra Finches (Taeniopygia guttata; Figure 75), you know that they are incessant nestbuilders. It was impossible to keep mosses in my garden room when I had finches because these mosses were prime nest-building material. But as you would also observe, not all selected mosses made it to the nest. Pieces would fall as the birds flew, and even the nest itself would occasionally lose pieces, but fragments would especially get dropped beneath the nest as the building progressed, in some cases deliberately as the birds determined that piece to be too recalcitrant to become part of the architecture. In addition to fragments and propagules travelling among feathers, it is also possible for bryophyte parts to

Figure 76. Turdus merula (Blackbird), a species that forages among Mnium hornum and Polytrichastrum formosum. Photo by Mario Modesto Mata through GNU Free Documentation.

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Figure 79. Hypnum cupressiforme, a species thrown about by Blackbirds in displacement activity. Photo by Jan-Peter Frahm, with permission.

Figure 77. Polytrichastrum formosum, a moss where Blackbirds forage, disturbing the moss. Photo by David T. Holyoak, with permission.

But it appears that might not be the only reason to cause Blackbirds (Turdus merula; Figure 76) to scatter bryophytes. Robin Stevenson reports (Bryonet 25 April 2010) observing a male of this same species of bird throwing clumps of mosses off a roof, alternately with mid air attacks by another Blackbird – a classic example of displacement! There was too much activity to discern if both birds were moss throwers. Apparently the two were fighting over territory or some other disagreement and the mosses were handy objects to throw from their rooftop habitat. In this case, the lucky roof mosses were Grimmia pulvinata (Figure 78), Hypnum cupressiforme (Figure 79), and Syntrichia montana (Figure 80). When on the ground they threw cockle shells and other things.

Figure 80. Syntrichia montana (Intermediate Screw-moss), a species thrown about by Blackbirds in displacement activity. Photo by Barry Stewart, with permission.

Figure 78. Grimmia pulvinata, a moss thrown about by a Blackbird during a territorial competition. Photo by Michael Lüth, with permission.

In another instance, Davison (1976) found spores of a moss on the feet of a dead Song Thrush (Turdus philomelos; Figure 81). Although most of the scavenging activity probably only transports moss fragments and spores for short distances, spores might occasionally be transported by feet, feathers, and beaks to considerable distances following such activity.

Figure 81. Turdus philomelos, Song Thrush, a bird known to carry mosses on its feet. Photo by Brian Eversham, with permission

But birds are imperfect in their industrious movement of moss from natural substrate to nest. Bits fall, and hence alight in a new location. This facilitated dispersal, while

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somewhat random, can be quite helpful in moving rarely fruiting mosses about. Chmielewski and Eppley (2019) found that when birds use bryophyte-covered areas for foraging and gathering nesting material, they can acquire propagules on their legs, feet, and tails. The researchers successfully germinated propagules from among the 242 propagules and 1512 spores they collected from 224 birds, comprised of bird 34 species. They found the tail feathers to be the greatest dispersal agents among bark and foliage species. Hence, birds are potential dispersal agents. The Pintail Duck (Anas acuta; Figure 82) is a likely agent of dispersal of Riccia rhenana (Figure 83) (McGregor 1961). In this liverwort, the older parts die, but the apices survive two months of drought and five weeks submersion in ice, making it likely that they would survive transport among the feathers of the Pintail Duck.

Figure 84. Tetraplodon mnioides with mature capsules; this species may be distributed by birds. Photo by Richard Caners, with permission.

Des Callaghan filmed a site where the White Wagtail (Motacilla alba; Figure 85) frequently perches on a particular branch. That branch is covered by Splachnum vasculosum (Figure 86-Figure 87). Does the bird simply like the soft moss and its location? Is the moss dispersed by the feathers and feet of the birds? Or might it be deposited in feces, indicating the birds ate the capsules?

Figure 82. Anas acuta, Northern Pintail male and female, agents of aquatic bryophyte dispersal, especially Riccia rhenana. Photo by J. M. Garg, through Creative Commons.

Figure 85. Motacilla alba alba, White Wagtail, a species that spends much time on a branch with Splachnum vasculosum in Wales. Photo by Luis Garcia, through Creative Commons.

Figure 83. Riccia rhenana, a species dispersed by pintail ducks. Photo by Štĕpán Koval, with permission.

Lewis et al. (2014b) suggested that Tetraplodon (Figure 84) species were distributed long-distances by birds. They reasoned that the absence of wind patterns to account for their distribution in the New World and the sensitivity of the spores to extreme environmental conditions, bird dispersal, probably on feathers, was the most reasonable explanation. In support of this possibility, Lewis et al. (2014a) demonstrated bryophyte diaspores among the feathers of transequatorial migrant birds.

Figure 86. Splachnum vasculosum growing on a branch next to a stream and the site where the White Wagtail, Motacilla alba, prefers to perch. Photo courtesy of Des Callaghan.

Chapter 16-1: Birds

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grows around the active burrows of shearwaters (Puffinus griseus; Figure 89) and diving petrels.

Figure 87. Splachnum vasculosum capsules. Photo by Dick Haaksma, with permission.

In some way the petrels and other sea birds seem to be responsible for the locations of members of Calymperaceae in the Chathams and other areas around New Zealand. Fife and Lange (2009) suggest dispersal by birds. They consider it likely that the sea birds may have contributed to dispersal of the moss Calymperes tenerum (Figure 88) on the Chatham Islands and the Kermadecs to the north and east of New Zealand, respectively. Peter de Lange (pers. comm. 12 June 2017) reported that until 80100 years ago, Tube Nose Petrels, especially Pterodroma spp. (Figure 92-Figure 93), were influential, but Broadbilled Prions (Pachyptila vittata) and shearwaters (Puffinus griseus; Figure 89) also were common in the areas where Calymperes grows now, but that these birds disappeared 80-100 years ago.

Figure 89. Puffinus griseus, Sooty Shearwater, a possible dispersal agent for Calymperes tenerum (Figure 88). Photo from USGS photograph by Jonathan Felis, through public domain.

Figure 90. Syrrhopodon, a genus that might be dispersed by sea birds in islands around New Zealand. Photo by Jan-Peter Frahm, with permission.

Figure 88. Calymperes tenerum, a species that may have been dispersed long distance by the Shearwater. Photo by JanPeter Frahm, with permission.

Later, de Lange (Peter de Lange, pers. comm. 12 June 2017) found Syrrhopodon armatus (Figure 90-Figure 91) on the smallest of the main Chatham Island, Rangatira. This island is free of predators and supports a million plus seabirds. The S. armatus grows on tree trunks that are used by the petrels and Broad-billed Prions (Pachyptila vittata) as runways. They also grow around the burrows of these birds, especially those of the Chatham Petrel (Pterodroma axillaris). On Rabbit Island, Syrrhopodon

Figure 91. Syrrhopodon armatus leaf, a possible propagule carried by sea birds to islands around New Zealand. Photo from Natural History Museum, London, through Creative Commons.

In addition to these islands, on the Chatham island of Rekohu and the Pitt island of Rangiuria, Calymperes (Figure 88) is found only in locations there the

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pterodromids once had dense nesting locations, as indicated by remains of their burrows (Peter de Lange, pers. comm. 12 June 2017). At the location where de Lange first found C. tenerum (Figure 88) there are still seabirds, including Taiko (Pterodroma magentae), a critically endangered species (Fife 2009). In New Zealand at Te Paki, Calymperes (Figure 88) again is associated with Pterodroma nigripennis (Figure 92) and P. gouldi (Peter de Lange, pers. comm. 12 June 2017). And on Raoul Island, all the locations found by de Lange were also in areas frequented by the Kermadec Petrel (Pterodroma neglecta neglecta; Figure 93) until the rats wiped them out early in the 20th Century. As on the Chatham Islands, the birds used the trees with Calymperes (Figure 88) as runways.

Based on what we know about these seabirdCalymperaceae relationships there are three plausible explanations for the relationships. The birds may fertilize the bark with guano, thus providing nitrogen for the mosses. The birds may serve as dispersal agents. The mosses may provide foraging substrate for the birds. Felicisimo et al. (2008) provided evidence that the Cory's Shearwater (Calonectris diomedea; Figure 94) follows wind patterns that could explain dispersal patterns. Cameron et al. (2006) have suggested that Buller's Shearwater (Puffinus bulleri; Figure 95) best explains the presence of the fern Asplenium pauperequitum on the Chatham Islands group, a distance of 1245 km from its nearest neighbor. This bird is a New Zealand endemic species and has large breeding populations on the Poor Knights Islands where Asplenium pauperequitum was originally described (Allan Fife, pers comm. 12 June 2017). In the Chathams it does not breed, but it is a regular visitor. Any and all of these explanations for the Calymperaceaeseabird associations may be true.

Figure 94. Calonectris diomedea, Cory's Shearwater flying, permitting it to disperse bryophytes over long distances. Photo by A. H. Kopelman, through Creative Commons. Figure 92. Pterodroma nigripennis, a species that seems to be associated with Calymperes (Figure 88) and may disperse it. Photo by Christopher Watson, through Creative Commons.

Figure 95. Puffinus bulleri, Buller's Shearwater, a species that might disperse mosses to islands near New Zealand. Photo by Tom Tarrant, through Creative Commons. Figure 93. Pterodroma neglecta, Kermadec Petrel, a species always found with Calymperes on Raoul Island. Photo by Lance Andrewes, through Creative Commons.

On the Poor Knights Islands, Jessica Beever has similarly collected Syrrhopodon armatus (Figure 90Figure 91) associated with a heavily burrowed petrel area (Allan Fife, pers. comm. 12 June 2017).

Chmielewski (2015) sought to support these suggestions by culturing propagules found on birds caught with mist nets. Using cotton swabs, he sampled feet, legs, and flight feathers. The spores obtained were cultured on nutrient agar. The resulting bryophyte plants were identified by PCR amplification and Sanger sequencing of the trnL region of the chloroplast genome. We shall have

Chapter 16-1: Birds

to look forward to the revelation of these species when this work is published. Dispersal of bryophytes by birds is discussed in more detail in subchapters 4-9 and 4-11 of Volume 1.

Soft Landings Pole jumpers have sand pits or mats to protect them when they land. To me it seems reasonable that birds might choose soft landing sites as well. Birds in captivity often get a condition known as bumblefoot (Figure 96) (Halliwell 1975; Hawkey et al. 1985), but the condition can occur in wild populations, albeit much less commonly (Gentz 1996). Bumblefoot can be caused by rough perches, sandpaper on the perch, sharp corners, dirty perches, or all perches of the same size. In the wild these problems are largely absent, explaining the scarcity of bumblefoot in nature. Do wild birds select landing spots on the basis of the presence of the spongy bryophytes and lichens (Figure 97)?

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breeding grounds, using moss hummocks as watch towers, throwing them in displacement behavior, bathing among them, and getting dry on them. On the other hand, the birds may help the bryophytes as dispersal agents and by providing fertilizer as guano. Or they may seriously disturb them during their foraging. Others provide so much guano that the bryophytes are intolerant of it. Soft bryophytes might also help to prevent bumblefoot in wild birds.

Acknowledgments Thank you to Brian Dykstra for sending me the wonderful thesis on birds and epiphytes by Adrian Wolf, as well as other references and personal observations. David Dumond shared the references he got from Bryonet. Thank you to Allan Fife for helping me get the details on the Shearwater dispersal story. Thank you to Janet Marr for a critical reading of the manuscript.

Literature Cited

Figure 96. Eagle bumblefoot, a common condition for birds of prey in captivity. Photo by Richard Jakowski, through Creative Commons.

Figure 97. Bird on moss perch – Is it a sentinel, or just cooling its feet on the moss? Photo by Ervin Gjata, through public domain.

Summary Birds interact with bryophytes by foraging among them, eating them, eating capsules, getting a drink, building nests or parts of nests with them, using them as

Alonso, J. C., Johnstone, G. W., Hindell, M., Osborne, P., and Guard, R. 1987. Las aves del Monolito Scullin, Antarctica Oriental (67°47′S 66°42′E). In: Castellri, J. (ed.). Actas del segundo simposio Español de estudios Antarcticos. Consejo superior de investigaciones cientificas, Madrid, pp. 375-386. Armstrong, E. A. 1955. The Wren. Collins, London. Barcikowski, A., Lyszkiewicz, A., Loro, P., Rektoris, L., Smyka, J., Wincenciak, A., and Zubel, P. 2005. Keystone species and ecosystems functioning: The role of penguin colonies in differentiation of the terrestrial vegetation in the Maritime Antarctic. Ecol. Quest. 6: 117-128. Behling, E., Caviness, T., Lewis, L. R., Jiménez, J. E., Goffinet, B., and Rozzi, R. 2016. Dispersal of bryophyte diaspores following ingestion by birds. Abstract 224. Botany 2016 poster, Savannah, GA, USA. 30 July – 3 August 2016. Bergstrom, D. M. and Seppelt, R. D. 1990. The lichen and bryophyte flora of Scullin Monolith, Mac. Robertson Land. Polar Rec. 26: 44-45. Bokhorst, S., Huiskes A. H. L., Convey, P., and Aerts, P. 2007a. Climate change effects on organic matter decomposition rates in ecosystems from the Maritime Antarctic and Falkland Islands. Global Change Biol. 13: 2642-2653. Bokhorst, S., Huiskes, A., Convey, A., and Aerts, R. 2007b. External nutrient inputs into terrestrial ecosystems of the Falkland Islands and the Maritime Antarctic region. Polar Biol. 30: 1315-1321. Brewer, R. 1967. Bird populations of bogs. Wilson Bull. 79: 371-396. Brightsmith, D. 1999. Stealth Conures of the genus Pyrrhura. Bird Talk Magazine, Electronic publication . Calmé, S. and Desrochers, A. 1999. Nested bird and microhabitat assemblages in a peatland archipelago. Oecologia 118: 361-370. Calmé, S. and Desrochers, A. 2000. Biogeographic aspects of the distribution of bird species breeding in Québec's peatlands. J. Biogeogr. 27: 725-732. Calmé, S., Desrochers, A., and Savard, J.-P. L. 2002. Regional significance of peatlands for avifaunal diversity in southern Québec. Biol. Conserv. 107: 273-281.

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Cameron, E. K., Lange, P. J. de, Perrie, L. R., Brownsey, P. J., Campbell, H. J., Taylor, G. A., Given, D. R., and Bellingham, R. M. 2006. A new location for the poor knights spleenwort (Asplenium pauperequitum, Aspleniaceae) on the Forty Fours, Chatham Islands, New Zealand. N. Z. J. Bot. 44: 199-209. Chmielewski, M. W. 2015. Abundance, species composition, and retention times of bryophyte spores on avian surfaces in a Pacific Northwest USA forest. (abstract). Internat. Assoc. Bryol. Conf. Chile, p. 5. Bryophytes on the Wing: First Evidence of Widespread Bryophyte Spore Vectoring on Bird Surfaces. Chmielewski, M. W. and Eppley, S. M. 2019. Forest passerines as a novel dispersal vector of viable bryophyte propagules. Proc. Roy. Soc. B 286: 20182253. . Cocks, M. P., Balfour, D. A., and Stock, W. D. 1998. On the uptake of ornithogenic products by plants on the inland mountains of Dronning Maud Land, Antarctica, using stable isotopes. Polar Biol. 20: 107-111. Coulson, J. C. and Whittaker, J. B. 1978. Ecology of moorland animals. In: Production Ecology of British Moors and Montane Grasslands. Springer, Berlin, Heidelberg, pp. 5293. Daily, G. C., Ehrlich, P. R., and Haddad, N. M. 1993. Double keystone bird in a keystone species complex. Proc. Natl. Acad. Sci. 90: 592-594. Davis, R. C. 1981. Structure and function of two Antarctic terrestrial moss communities. Ecol. Monogr. 51: 125-143. Davison, G. W. H. 1976. Role of birds in moss dispersal. British Birds 69: 65-66. Desrochers, A., Rochefort, L., and Savard, J.-P. L. 1998. Avian recolonization of eastern Canadian bogs after peat mining. Can. J. Zool. 76: 989-997. Emslie, S. and Woehler, E. 2005. A 9000-year record of Adélie penguin occupation and diet in the Windmill Islands, East Antarctica. Antarct. Sci. 17: 57-66. Erkamo, V. 1976. Warikset kallioiden sammalpeitteen turmelijoina. [Crows disturbing the moss cover of rocks in Helsinki.]. Luonnon Tutkija 80(2): 57-58. Erskine, P., Bergstrom, D., Schmidt, S., Stewart, G., Tweedie, C., and Shaw, J. 1998. SubAntarctic Macquarie Island - a model ecosystem for studying animal-derived nitrogen sources using 15N natural abundance. Oecologia 117: 18793. [FEMAT] Forest Ecosystem Management Assessment Team. 1993. Forest Ecosystem Management: An Ecological, Economic, and Social Assessment. Portland, OR. US Department of Agriculture, Forest Service, US Department of Commerce, National Oceanic and Atmospheric Administration, US Department of the Interior, Bureau of Land Management, US Fish and Wildlife Service, National Park Service, Environmental Protection Agency. Fife, A. 2009. Calymperes tenerum Müll. Hal. (Calymperaceae) on the Chatham Islands, New Zealand. Australasian Bryol. Newslett. 57: 14-16. Fife, A. J. and Lange, P. J. de. 2009. Calymperes tenerum Müll. Hal. (Calymperaceae) on the Chatham Islands, New Zealand. Australasian Bryol. Newslett. 57: 14-16. Gentz, E. J. 1996. Fusobacterium necrophorum associated with bumblefoot in a wild great horned owl. J. Avian Med. Surg. 10: 258-261. Gill, Victoria 2012. Antarctic moss lives on ancient penguin poo. BBC Nature News, 5 July 2012, last updated at 01:36. Accessed at .

Gradstein, S. R., Hietz, P., Lücking, R., Lücking, A., Sipman, H. J. M., Vester, H. F. M., Wolf, J. H. D., and Gardette, E. 1996. How to sample the epiphytic diversity of tropical rain forests. Ecotropica 2: 59-72. Haddad, S., Desrochers, A., and Savard, J.-P. L. 2000. Artificial nest predation bogs: Does peat harvest increase risk? Ecoscience 7: 32-37. Halliwell, W. H. 1975. Bumblefoot infections in birds of prey. J. Zoo Anim. Med. 6(4): 8-10. Hawkey, C., Samour, H. J., Henderson, G. M., and Hart, M. G. 1985. Haematological findings in captive Gentoo Penguins (Pygoscelis papua) with bumblefoot. Avian Pathol. 14: 251256. Hodkinson, I. D., Coulson, S., Webb, N. R., Block, W., Strathdee, A. T., and Bale, J. S. 1994. Feeding studies on Onychiurus arcticus (Tullberg) (Collembola: Onychiuridae) on West Spitsbergen. Polar Biol. 14: 17-19. Hollén, L. I., Bell, M. B., and Radford, A. N. 2008. Cooperative sentinel calling? Foragers gain increased biomass intake. Current Biol. 18: 576-579. Hughes, J. B., Daily, G. C., and Ehrlich, P. R. 2002. Conservation of tropical forest birds in countryside habitats. Ecol. Lett. 5: 121-129. Jasmin, J.-N., Line Rochefort, L., and Gauthier, G. 2008. Goose grazing influences the fine-scale structure of a bryophyte community in Arctic wetlands. Polar Biology 31: 10431049. Jofre, J., Goffinet, B., Marino, P., Raguso, R. A., Nihei, S. S., Massardo, F., and Rozzi, R. 2011. First evidence of insect attraction by a Southern Hemisphere Splachnaceae: The case of Tayloria dubyi Broth. in the Reserve Biosphere Cape Horn, Chile. Nova Hedw. 92: 317-326. Koponen, A. 1990. Entomophily in the Splachnaceae. J. Linn. Soc. Bot. 104: 115-127. Kuc, M. 1996. Briofityczne wieze obserwacyjne ptakow z Ziemi Ognistej na ogolnym tle powiazan mszakow ze zwierzetami. [Bryophyte watch-towers of birds from Tierra del Fuego in general context of bryo-animal relationships.] Fragm. Flor. Geobot. Ser. Polon. 3: 395-398. Lachance, D., Lavoie, C., and Desrochers, A. 2005. The impact of peatland afforestation on plant and bird diversity in southeastern Québec. Ecoscience 12: 161-171. Langevin, A. E. 2015. Avian guano as a nutrient input to cliffface ecosystems in western North Carolina. M. S. Thesis, Appalachian State University, Boone, N. C. Lee, Y., Hs, L., and Yoon, H. 2009. Carbon and nitrogen isotope composition of vegetation on King George Island, maritime Antarctic. Polar Biol. 32: 1607-1615. Lendrem, D. W. 1983a. Predation risk and vigilance in the blue tit (Parus caeruleus). Behav. Ecol. Sociobiol. 14: 9-13. Lendrem, D. W. 1983b. Sleeping and vigilance in birds. I. Field observations of the mallard (Anas platyrhynchos). Anim. Behav. 31: 532-538. Lewis, L. R., Behling, E., Gousse, H., Qian, E., Elphick, C. S., Lamarre, J.-F., Bêty, Liebezeit, J., Rozzi, R., and Goffinet, B. 2014a. First evidence of bryophyte diaspores in the plumage of transequatorial migrant birds. PeerJ 2:e424. Lewis, L. R., Rozzi, R., and Goffinet, B. 2014b. Direct longdistance dispersal shapes a New World amphitropical disjunction in the dispersal-limited dung moss Tetraplodon (Bryopsida: Splachnaceae). J. Biogeogr. 41: 2385-2395. MacLean, S. F. Jr. 1980. The detritus-based trophic system. In: Brown, J., Miller, P. C., Tieszen, L. L., and Bunnell, F. L. (eds.). An Arctic Ecosystem: The Coastal Tundra at Barrow,

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Alaska. Dowden, Hutchinson & Ross, Inc., Stroudsburg, PA, pp. 411-457. McGregor, R. L. 1961. Vegetative propagation of Riccia rhenana. Bryologist 64: 75-76. Metcalfe, N. B. 1984. The effects of habitat on the vigilance of shorebirds: Is visibility important? Anim. Behav. 32: 981985. Metcalfe, N. B. and Furness, R. W. 1984. Changing priorities: the effect of pre-migratory fattening on the trade-off between foraging and vigilance. Behav. Ecol. Sociobiol. 15: 203-206. Nadkarni, N. M. 1994. Diversity of species and interactions in the upper tree canopy of forest ecosystems. Amer. Zool. 34: 70-78. Nadkarni, N. M., Schaefer, D., Matelson, T. J., and Solano, R. 2004. Biomass and nutrient pools of canopy and terrestrial components in a primary and a secondary montane cloud forest, Costa Rica. Forest Ecol. Mgmt. 198: 223-236. Niemi, G. J. and Hanowski, J. A. M. 1992. Bird populations. Chapt. 8. In: Wright, H. E., Coffin, B. A., and Aaseng, N. E. (eds.). The Patterned Peatlands of Minnesota. Commissioner of Natural Resources, pp. 111-129. Paasivirta, L, Lahti, T., and Peraetie, T. 1988. Emergence phenology and ecology of aquatic and semi-terrestrial insects on a boreal raised bog in Central Finland. Holarct. Ecol. 11: 96-105. Perkins, R. C. L. 1903. Vertebrata. Aves. In: Sharp, D. (ed.). Fauna Hawaiiensis or the Zoology of the Sandwich (Hawaiian) Isles. C. J. Clay and Sons, Cambridge University Press Warehouse, London. Poulin, M. 2002. Reserve networks for a peatland bird: The importance of within-site habitat configuration and offnetwork habitat loss (abstract). Norges Tekn. – Naturvitensk. Univ. Vitenskapsmus. Rapp. Bot. 2002-3: 21. Sekercioglu, C. H., Loarie, S. R., Oviedo Brenes, F., Ehrlich, P. R., and Daily, G. C. 2007. Persistence of forest birds in the

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Costa Rican agricultural countryside. Conserv. Biol. 21: 482-494. Simberloff, D. 1998. Flagships, umbrellas, and keystones: Is single-species management passé in the landscape era? Biol. Conserv. 83: 247-257. Smykla, J., Wolek, J., and Barcikowski, A. 2007. Zonation of vegetation related to penguin rookeries on King George Island, maritime Antarctic. Arct. Antarct. Alp. Res. 39: 143151. Steere, W. C. 1976. Ecology, phytogeography and floristics of Arctic Alaskan bryophytes. J. Hattori Bot. Lab. 41: 47-72. Sullivan, K. A. 1984. The advantages of social foraging in downy woodpeckers. Anim. Behav. 32: 16-22. Takaki, N. 1957. [Certain mosses are utilized by birds and insects.] Misc. Bryol. Lichenol. 12: 1-2. Vanderpuye, A. W., Elvebakk, A., and Nilsen, L. 2002. Plant communities along environmental gradients of high-Arctic mires in Sassendalen, Svalbard. J. Veg. Sci. 13: 875-884. Wasley, J., Robinson, S. A., Turnbull, J. D., King, D. H., Wanek, W., and Popp, M. 2012. Bryophyte species composition over moisture gradients in the Windmill Islands, East Antarctica: Development of a baseline for monitoring climate change impacts. Biodiversity 13: 257-264. Watson, E. V. 1964. The Structure and Life of Bryophytes. Hutchinson Univ. Libr., London, 192 pp. Wickler, W. 1985. Coordination of vigilance in bird groups. The "Watchman's Song" hypothesis. Ethology 69: 250-253. Wolf, A. L. 2009. Bird use of epiphyte resources in an oldgrowth coniferous forest of the Pacific Northwest. Master's Thesis, Evergreen State College, WA, USA. Wuczyński, A., Dajdok, Z., Wierzcholska, S., and Kujawa, K. 2014. Applying red lists to the evaluation of agricultural habitat: Regular occurrence of threatened birds, vascular plants, and bryophytes in field margins of Poland. Biodiv. Conserv. 23: 999-1017.

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Glime, J. M. 2017. Birds and Bryophytic Food Sources. Chapt. 16-2. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. eBook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 19 July 2020 and available at .

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CHAPTER 16-2 BIRDS AND BRYOPHYTIC FOOD SOURCES TABLE OF CONTENTS Capsules ........................................................................................................................................................... 16-2-2 Ptarmigans................................................................................................................................................. 16-2-5 Grouse ....................................................................................................................................................... 16-2-7 Titmice ...................................................................................................................................................... 16-2-7 Kōkako ........................................................................................ 16-2-8 Fruit Mimicry by Capsules? ...................................................................................................................... 16-2-9 Bird Color Vision............................................................................................................................. 16-2-10 Leafy Plants.................................................................................................................................................... 16-2-11 Ducks and Food Availability .................................................................................................................. 16-2-12 Geese ....................................................................................................................................................... 16-2-12 Blood Pheasant........................................................................................................................................ 16-2-16 Kakapo .................................................................................................................................................... 16-2-17 Turkeys?.................................................................................................................................................. 16-2-18 Dispersal ................................................................................................................................................. 16-2-18 Nutritional Value of Bryophytes .................................................................................................................... 16-2-18 Palatability ..................................................................................................................................................... 16-2-20 Foraging ......................................................................................................................................................... 16-2-20 Ground Foragers ..................................................................................................................................... 16-2-20 Arctic Foraging Effects ........................................................................................................................... 16-2-20 Foraging on Epiphytes ............................................................................................................................ 16-2-21 Juncos .............................................................................................................................................. 16-2-27 Weaver Birds ................................................................................................................................... 16-2-28 Tropical Birds .................................................................................................................................. 16-2-28 Jamaican Blackbird .......................................................................................................................... 16-2-29 Summary ........................................................................................................................................................ 16-2-29 Acknowledgments .......................................................................................................................................... 16-2-30 Literature Cited .............................................................................................................................................. 16-2-30

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Chapter 16-2: Birds and Bryophytic Food Sources

CHAPTER 16-2 BIRDS AND BRYOPHYTIC FOOD SOURCES

Figure 1. Branta bernicla hrota, Brant, juvenile foraging; foods include bryophytes. Photo by MPF, through Creative Commons.

Many birds do depend on bryophytes for food. Some eat the leafy gametophytes, especially in the Arctic. Others use the more nutrient-rich capsules. And others, probably many more than we know, forage for macroinvertebrates among the bryophytes, especially epiphytes.

Capsules A. J. Grout, one of the earliest of North American bryologists, observed birds pecking the capsules of Polytrichum commune (Figure 2), a story retold by Lewis Anderson (Bryonet 10 April 2003). To this story, Frank Cook (Bryonet 15 May 2001) contributed his own observations of White-throated Sparrows (Zonotrichia albicollis; Figure 3) "vigorously nipping the capsules from Polytrichum in a white pine (Pinus strobus; Figure 4) stand in Algonquin Park, Ontario.

Figure 2. Polytrichum commune capsules, food for Whitethroated Sparrows (Zonotrichia albicollis) and Norwegian Grouse (Tetrao urogallus?) chicks. Photo by Bob Klips, with permission.

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and Polytrichum (Figure 2) are eaten by the Norwegian Grouse chicks (Tetrao urogallus?; Figure 6), apparently as the main food, whereas other kinds of capsules are eaten by Scottish Red Grouse (Lagopus lagopus scotica; Figure 7) (Lid & Meidell 1933). The Wyoming Sage Grouse (Centrocercus urophasianus; Figure 8) eats small amounts of moss, Snow Buntings (Plectrophenax nivalis; Figure 9) eat Bryum algovicum capsules (Figure 10), and the Moorhen (Gallinula chloropus; Figure 11), Blackbird (Turdus merula; Figure 12), Song Thrush (Turdus philomelos; Figure 13), and Fieldfare (Turdus pilaris; Figure 14) all eat mosses. In Britain, the Blue Tits (Cyanistes caeruleus; Figure 15) and Marsh Tits (Poecile palustris; Figure 16) feed on capsules of Dicranoweisia cirrata (Figure 17) (Betts 1955). Catherine La Farge reported on Bryonet (15 January 2008) that high Arctic moss capsules are consumed by lemmings and Arctic hares. Thus it would not be surprising if birds also consume them when the capsules are still green.

Figure 3. Zonotrichia albicollis, White-throated Sparrow, a consumer of Polytrichum capsules. Photo by Dorothy Pugh, with permission.

Figure 5. Bryum arcticum with capsules that serve as food for Norwegian Grouse (Tetrao urogallus?) chicks in Norway. Photo by Michael Lüth, with permission.

Figure 4. Pinus strobus (white pine) forest, Pennsylvania. Photo by Nicholas T., through Creative Commons.

Richardson (1981) reported moss-feeding by mammals and birds in northern areas. Capsules of Bryum (Figure 5)

Figure 6. Tetrao urogallus, Norwegian Grouse female, on moss. Chicks of this species eat capsules of Bryum and Polytrichum. Photo by Honza Sterba, through Creative Commons.

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Figure 10. Bryum algovicum with capsules that are eaten by the Snow Bunting. Photo by Barry Stewart, with permission.

Figure 7. Lagopus lagopus scotica, Red Grouse, a species that eats moss capsules. Photo by MPF, through Creative Commons.

Figure 11. Gallinula chloropus, Moorhen, a moss consumer. Photo from Anemone Projectors, through Creative Commons.

Figure 8. Centrocercus urophasianus, Greater Sage Grouse, a consumer of small amounts of mosses. Photo by Gordon Sherman, with online permission.

Figure 9. Plectrophenax nivalis, Snow Bunting, a herbivore on the capsules of Bryum pendulum. Photo by Cephas, through Creative Commons.

Figure 12. Turdus merula, a Blackbird that eats mosses. Photo by Mario Modesto Mata through GNU Free Documentation.

Chapter 16-2: Birds and Bryophytic Food Sources

Figure 13. Turdus philomelos, Song Thrush, in Cambridgeshire, a bird that eats mosses. Photo by Brian Eversham, with permission.

Figure 14. Turdus pilaris, Fieldfare, a bird that eats mosses. Photo by Frankie Fouganthin, through Creative Commons.

Figure 15. Cyanistes caeruleus, Blue Tit, in winter, a bird that eats capsules of Dicranoweisia cirrata. Photo through public domain.

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Figure 16. Poecile palustris, Marsh Tit, a species that eats capsules of Dicranoweisia cirrata. Photo by Luc Viatour, through Creative Commons.

Figure 17. Dicranoweisia cirrata with capsules that are eaten by Blue Tits and Marsh Tits. Photo from BioPix, through Creative Commons.

Dan Norris (Bryonet, 22 November 1995 & 19 November 2006) reported that the Green Eastern Rosella Parrot (Platycercus eximius; Figure 18) in Tasmania selects the green, but mature, capsules of Polytrichum juniperinum (Figure 19) on clay soil banks as a primary food source. He watched the parrots for over an hour, then examined the area to find that they clipped the setae at 45º angles and left a miniature forest of setae with a litter of calyptrae that were split off, falling 5-10 mm to the right of the sporophyte. The number of barren setae suggested that harvest in this manner was widespread. Further examination on other clay banks of the island revealed that similar patterns were common in the forested mid-elevation habitats throughout the island. Ptarmigans In northern Europe and Alaska, the Willow Ptarmigan (Lagopus lagopus; Figure 20-Figure 21, Figure 23) chicks consume moss capsules of Polytrichum s.l. (Figure 19) and Pohlia (Figure 22) (Weeden 1969; Gardarsson & Moss

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1970; Spidsø 1980; Martin & Hik 1992). Pullianen and Eskonen (1982) considered that moss capsules could be a source of high quality food in this Arctic environmental at a time when they were too small to handle large food items.

Figure 18, Platycercus eximius diemenensis, Green Eastern Rosella Parrot male, a species that selects green capsules of Polytrichum juniperum as food. Photo by J. J. Harrison, through Creative Commons.

Figure 19. Polytrichum juniperinum mature capsules that are still green under the calyptra, providing food for the Green Eastern Rosella Parrot (Platycercus eximius). Photo by Ian Sutton, through Creative Commons.

1974). In two cases the large numbers of capsules consumed suggest food selection rather than accidental ingestion (Martin & Hik 1992).

Figure 21. Lagopus lagopus lagopus, Willow Ptarmigan in winter plumage. Chicks of this species eat capsules of Polytrichum and Pohlia. Photo through Creative Commons.

Figure 22. Pohlia nutans with capsules. Capsules from this genus are eaten by the Willow Ptarmigan in the North. Photo by Michael Lüth, with permission.

Martin and Hik (1992) found the crops of Willow Ptarmigan chicks (Lagopus lagopus; Figure 23) stuffed with capsules of the moss Distichium inclinatum (Figure 24). The researchers suggested that the sporophytes might be easily accessible forage for these chicks. Could the capsules possibly act as grinding agents for other foods?

Figure 20. Lagopus lagopus lagopus, Willow Ptarmigan in summer plumage. Chicks of this species consume mosses. Photo by George Lesard, through Creative Commons.

The consumption of these moss capsules by Willow Ptarmigan chicks appears to be a regular event every spring as the capsules appeared in the diet in three consecutive years (Martin & Hik 1992). It is likely that they supply needed lipids; they contain about 20% lipids, a level higher than that in the other available vegetation (Pakarinen & Vitt

Figure 23. Lagopus lagopus lagopus cf pullus, Willow Ptarmigan juvenile, a consumer of moss capsules of Polytrichum and Pohlia. Photo by Walter Pfliegler, with permission.

Chapter 16-2: Birds and Bryophytic Food Sources

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Figure 24. Distichium inclinatum with capsules. Willow Ptarmigan chicks eat the capsules and they can be found in the crops of the birds. Photo by Michael Lüth, with permission.

Grouse Grouse (Tetraoninae) chicks (Figure 7) are known to eat moss capsules (Richardson 1981). In fact, the clutch size and mean egg weight are dependent on the food of the mother (Naylor & Bendell (1989). The two most preferred foods were the trailing arbutus (Epigaea repens; Figure 25) and capsules of Polytrichum (Figure 19), and their availability was important, but not the size of the hen or her scaled body weight. Egg size, on the other hand, was not related to spring diet, but was instead related to the size of the hen. Therefore, the spring diet was important in providing the nutrients required for clutch formation.

Figure 26. Baeolophus, Crested Titmouse, a genus that grazes on the tips of mosses, perhaps to eat capsules. Photo by Dick Daniels, through Creative Commons.

Betts (1955) considered that in oak woodlands the Great Tit (Parus major; Figure 27) and the Blue Bit (Cyanistes caeruleus; Figure 15) can compete for food with the Coal Tit (Periparus ater; Figure 28) and the Marsh Tit (Poecile palustris; Figure 29). Using gizzard analyses, she determined that the Great Tit and Blue Tit had different diets, with the former feeding mostly on adult insects, especially weevils, and the Blue Tit on scale insects, small larvae, and pupae. The Coal Tit fed mostly on small, free-living insects and scales. The Marsh Tit ate mostly adult insects, scales, and a few larval forms. But in winter the diet changed. The Blue Tit consumed large numbers of capsules from the moss Dicranoweisia cirrata (Figure 30), ignoring the capsules of all other species. It had so many capsules in its gizzard that the gizzard was a vivid green (300-450 capsules per gizzard). One Coal Tit had consumed a few capsules and one Marsh Tit had 233 capsules in the gizzard.

Figure 25. Epigaea repens, one of the two most preferred foods of grouse chicks. Photo by Fritz Flohr Reynolds, through Creative Commons.

Titmice Titmice eat moss capsules in the temperate zone (Richardson 1981). Haftorn (1954) on five occasions observed the Crested Titmouse (Baeolophus sp.; Figure 26) on snow-free rocks with mosses. The birds were pulling at the tips of the moss and Haftorn surmised that they were probably eating the capsules.

Figure 27. Parus major, Great Tit, a consumer of adult insects. Photo by Francis Franklin, through Creative Commons.

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Figure 28. Periparus ater, Coal Tit, a species that feeds on small, free-living insects and scales, but consumes large numbers of moss capsules in winter. Photo by David Kesl, through Creative Commons.

Figure 29. Poecile palustris, Marsh Tit, a species that switches to eating moss capsules in the winter. Photo by Luc Viatour, through Creative Commons.

Figure 31. Parus cristatus, Crested Titmouse, a species that harvests mosses in early winter. Photo by Jiří Duchoň, through Creative Commons.

Kōkako The Kōkako/Blue-wattled Crow (Callaeas wilsoni; Figure 32) in New Zealand feeds on moss capsules (Jessica Beever, Bryonet 2 May 2003, based on observations by personnel from the Department of Conservation). Of 912 observations, 26 were feeding on moss capsules. When it was a good year for tracheophytes, only 3 out of 217 observations were of capsule feeding, but in a poor-fruit year, this increased to 6 out of 178 on mosses. These are probably within normal variation, but it suggests that the moss capsules may serve as an emergency food. The Kōkako forage along the branches, snipping off the capsules with the edge of the beak. Although they also feed on invertebrates from the bark and mosses, their action in obtaining the mosses by deliberate cutting is different from the pecking used to obtain insects. Eating the capsules is no accident. The Kōkako (Callaeas wilsoni) make their greatest use of mosses in spring and summer (3%) when the capsules are most abundant, but they also may consume some in winter (0.75%) (Jessica Beever, Bryonet 2 May 2003, based on observations by personnel from the Department of Conservation). The actual consumption may be larger as it is more difficult to observe moss feeding than that on bright-colored fruits.

Figure 30. Dicranoweisia cirrata with capsules that provide winter food for the Blue Tit (Cyanistes caeruleus; Figure 15). Photo from BioPix, through Creative Commons.

In Norway, one might see the Crested Tit (Parus cristatus; Figure 31) pulling on moss tips that are free from snow on rocks in December (Haftorn 1954).

Figure 32. Callaeas wilsoni, Kōkako, a bird that feeds on moss capsules. Photo by Duncan, through Creative Commons.

Chapter 16-2: Birds and Bryophytic Food Sources

Fruit Mimicry by Capsules? Michael Lüth (Bryonet 16 January 2008) has observed that some members of the Splachnaceae change their odor as they mature. Tetraplodon mnioides (Figure 33) has violet-colored capsules that smell like blueberries when the capsules are still closed. Once the capsules open, the odor changes to the smell of dung. A similar change occurs in Splachnum ampullaceum (Figure 34). When this species has immature capsules, the capsules have a strong, sweet odor like berries. But once the capsule opens it smells like dung. Could it be that in these early fruity stages the capsules are eaten by the local fauna, including birds? Patricia Geissler once expressed the idea that birds eat the capsules of Voitia nivalis (Figure 35) that occur among the buds of Salix herbacea (Figure 36), an early season food for some of the Arctic birds. If so, this is another potential dispersal mechanism. One might be able to make some interesting observations from within a duck blind, or using time-lapse photography.

Figure 33. Tetraplodon mnioides with mature capsules that might be eaten by the local fauna. Photo by Richard Caners, with permission.

Figure 34. Splachnum ampullaceum, showing capsules that resemble some of the nearby fruits. Photo by Michael Lüth, with permission.

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Figure 35. Voitia nivalis with capsules on Svalbard. These capsules resemble fruits of Salix herbacea (Figure 36) and may be eaten along with them. Photo by Michael Lüth, with permission.

Figure 36. Salix herbacea fruits in Austria, resembling capsules of Voitia nivalis. Photo by El Grafo, through Creative Commons.

While in Tasmania in December for the Australasian Bryological Workshop, Paddy Dalton and Rod Seppelt showed their fellow bryologists Pleurophascum grandiglobum (Figure 37), a moss of the button grass plains in SW Tasmania. Allison Downing (Bryonet 18 January 2008) was "intrigued by the capsules (Figure 37), which are extremely large, globular, cleistocarpous, and on quite long setae, and was curious about dispersal, particularly the possibility that this species might be dispersed by birds. The capsules are light green, fading to pale yellow, and to me, had much in common with the fruits of many Epacridaceae (Ericaceae) and also of Persoonia (Proteaceae; Figure 38) that grow in this area." Emma Pharo stated that there are a number of birds that do feed on the ground in the button grass plains (Allison Downing, Bryonet 18 January 2008). The birds might not gain any nutrition from the capsules and their contents, but mimicry is used by many plants for pollination so why not for dispersal? The New Zealand species of Pleurophascum, similarly, has globular fruits that become orange/red with maturity, and the color (red, orange) would make them even more attractive to birds.

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Chapter 16-2: Birds and Bryophytic Food Sources

Figure 37. Pleurophascum grandiglobum with capsules that are large and may be eaten by birds and dispersed by them. Photo by Christopher Taylor, Australian National Botanic Gardens, with online permission. Figure 40. Tayloria gunnii with capsules, possible mimics of some of the fruits in the Ericaceae. Photo by Christopher Taylor, Australian National Botanic Gardens, with online permission.

Figure 38. Persoonia levis fruit; Pleurophascum grandiglobum capsules (Figure 37) mimic these and may be eaten by some of the same bird species. Photo by John Tann, through Creative Commons.

Michael Lüth's comment about Tayloria (Figure 39Figure 41) reminded Downing that three species of Tayloria, T. octoblepharum (Figure 39), T. gunnii (Figure 40), and Tayloria tasmanica (Figure 41), all with abundant and conspicuous capsules, grow in the same habitat as Pleurophascum (Figure 37). Perhaps they, too, are fragrant (like the fruits of some Ericaceae) in their early stages of development and dispersed by birds before they reach the 'dung'-smelling stage of their life cycle. Figure 41. Tayloria tasmanica with capsules, possible mimics of some of the fruits in the Ericaceae. Photo by Paddy Dalton, with permission.

Bird Color Vision

Figure 39. Tayloria octoblepharum with capsules, possible mimics of some of the fruits in the Ericaceae. Photo by Janice Glime.

To understand bird choice based on color, it is necessary to understand how birds see color. Most studies on bird responses to color have assumed that they see colors the same way as humans do (Bennett et al. 1994). However, this is not true. The human eye design is different from that of birds and has different spectral abilities. Birds have four types of cones in the retina, compared to our three (Finger & Burkhardt 1994). Among their differences, at least some birds are able to see UV light, and feathers of some birds reflect UV light (Bennett & Cuthill 1994).

Chapter 16-2: Birds and Bryophytic Food Sources

Using gene coding for UV- or violet-absorbing opsin in the retina, Ödeen & Håstad (2003) were able to assess color sensitivities on living birds. Their color vision can be put into two classes: short-wavelength sensitivity biased toward violet and another biased toward UV. The violet sensitivity is ancient among birds, and sensitivity to UV has evolved independently in four evolutionary lines. Many members of the orders Psittaciformes (parrots) and Passeriformes (perching birds) present UV-sensitive type color vision, but within the Passeriformes, the Corvidae (Jays, Magpies, & Crows) and Tyrannidae (Tyrant Flycatchers) do not. At least some members of Laridae (Skuas, Gulls, Terns, & Skimmers – Charadriiformes) and Struthionidae (flightless birds – Struthioniformes) likewise have UV-sensitive vision. Birds of prey (Accipitridae & Falconidae – Falconiformes), on the other hand, have the violet type. The colorations of songbirds are significantly more conspicuous to other songbirds than they are to raptors and covids in the coniferous and deciduous forests (Finger & Burkhardt 1994; Håstad et al. 2005). This difference permits the Passeriformes to advertise their colors for mating purposes while not advertising to the raptors (birds of prey) that are their predators. In addition to their cones birds have a complex of oil droplets in their retinas that may alter the color hues they perceive and that may also alter brightness and saturation (Bennett et al. 1994). Bennett and coworkers caution us that color is a product of the perception of the observer. This brings us to the question of bird choice of bryophyte capsules and leafy stalks based on color. We know that bryophytes often serve as emergency food. Consider the observation of Bennett and Théry (2007) that plants are most likely to produce conspicuous fruit colors at times when frugivorous bird abundance is low. By contrast, if seeds, or bryophyte spores, are dispersed by birds, then I would think it would be beneficial for the fruits and capsules if they were bright-colored when it is appropriate for dispersal. But capsules are not the only parts of bryophytes that are eaten. As you will soon see, leafy parts are as well. And we know that at least some bryophytes have fluorescent cell walls. For example, the bulbils of Pohlia are fluorescent under UV light (Nordhorn-Richter 1984). The value of this fluorescence for dispersal by birds remains unexplored.

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Figure 42. The Red-throated Loon, Gavia stellata, and young. This species actually eats the leafy bryophytes in the Pacific Northwest, USA. Photo by David Karnå, through Creative Commons.

Figure 43. Lagopus leucura, White-tailed Ptarmigan, Rocky Mountains, Alberta, a species that eats leafy bryophytes in the Arctic. Photo by John Hill, through Creative Commons.

Leafy Plants It is uncommon for birds to use leafy bryophytes for food, but they may do so when food is scarce (Sillett 1994; Rhoades 1995; Wolf 2009). Among the few birds that actually eat the leafy bryophytes, we know that the Redthroated Loon (Gavia stellata; Figure 42), Brant (Branta bernicla; Figure 1), White-tailed Ptarmigan (Lagopus leucura; Figure 43), Willow Ptarmigan (Lagopus lagopus lagopus; Figure 44), and Rock Ptarmigans (Lagopus muta; Figure 45) all eat bryophytes in the Pacific Northwest, USA (Palmer 1962; Martin & Hik 1992; Braun et al. 1993; Hannon et al. 1998).

Figure 44. Lagopus lagopus lagopus, Willow Ptarmigan, with summer plumage, sitting on its dinner plate of leafy bryophytes. Photo by George Lesard, through Creative Commons.

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Chapter 16-2: Birds and Bryophytic Food Sources

Figure 47. Branta canadensis, Canada Geese and goslings. This species avoids eating the moss Fontinalis. Photo by Janice Glime. Figure 45. Lagopus muta, Rock Ptarmigan in summer plumage, a species that eats leafy bryophytes. Photo by Böhringer Friedrich, through Creative Commons.

Ducks and Food Availability For ducks, bryophytes are not a preferred food. Ringnecked Ducks (Aythya collaris; Figure 46) in temporary wetlands use mostly plants, but those in more permanent wetlands choose animal foods for half their diet. The period during pre-laying and laying is an important time for females to obtain protein, and in the northern long days of Minnesota, USA, the females may feed up to 19 hours a day to obtain needed protein. However, when their usual food sources are unavailable, Ring-necked Ducks (Aythya collaris) may eat bryophytes (Hohman 1985). In 1980, reduced protein content in Class II juveniles seemed to be the result of a large percentage of aquatic mosses and caddisflies in cases. In that year, aquatic mosses comprised 18% of the diet, whereas in other years there were only trace amounts.

Figure 46. Aythya collaris, Ring-necked Duck male, a species that obtains protein from mosses. Photo by Alan Vernon, through Creative Commons.

Geese Geese seem to have a love-hate relationship with mosses as a food source. Sometimes they are essential to the diet, but in other times and places, they are deliberately avoided. The Canada Goose (Branta canadensis; Figure 47) selectively consumes the riverweed Podostemum ceratophyllum (Figure 48) over the moss Fontinalis novaeangliae (Figure 49) in a riverine system, despite the dominance (89% of biomass) of moss in that system. This preference may have been due to the presence of C18 acetylenic acid, octadeca-9,12-dien-6-ynoic acid in the mosses, a compound that deters crayfish feeding.

Figure 48. Podostemum ceratophyllum, a flowering plant species that is preferred over mosses as food by Canada Geese. Photo by Alan Cressler, with permission.

Figure 49. Fontinalis novae-angliae protecting invertebrates from Canada Goose grazing because the geese won't eat it. Photo by John Parker, with permission.

By contrast, polar and alpine habitats seem to encourage the consumption of bryophytes, including by geese (Longton 1992). Gloutney et al. (2001) report that at Karrak Lake, NT, Canada Geese (Branta canadensis; (Figure 47), Lesser Snow Geese (Chen caerulescens caerulescens; Figure 50) and Ross's Geese (Chen rossii; Figure 51) eat primarily mosses, chickweed (Stellaria spp.; Figure 52), and sedges (Carex spp.; Figure 53). In the Svalbard breeding season, mosses form a considerable part of the diet of Barnacle Geese (Branta leucopsis; Figure 54) (Prop et al. 1980).

Chapter 16-2: Birds and Bryophytic Food Sources

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Figure 50. Chen caerulescens, Lesser Snow Geese, grazing on sedges. Photo by Walter Siegmund, through Creative Commons.

Figure 53. Carex aquatilis var. minor in water; members of this genus are eaten by several species of geese. Photo by Jeffery M. Saarela, through Creative Commons.

Figure 51. Chen rossii, Ross's Goose, grazing on sedges. Photo by Andrew C., through Creative Commons.

Figure 54. Branta leucopsis, Barnacle Goose, grazing. This species grazes largely on mosses in the Arctic. Photo by Arthur Chapman, through Creative Commons.

Figure 52. Stellaria humifusa; members of this genus are eaten by several species of geese. Photo by Lynn J. Gillespie, through Creative Commons.

Barnacle Geese (Branta leucopsis; Figure 54) arrive in Spitzbergen, Scandinavia, after a long migration, but before flowering plants are available (Prop & Vulink 1992). Thus mosses are eaten heavily during pre-laying and laying periods (62% in feces) (Fox & Bergersen 2005). The young goslings also consume the mosses, and sampling revealed that 27 out of 28 samples of adult and gosling droppings contained mosses (Prop & Vulink 1992). Snow Geese (Chen caerulescens caerulescens; Figure 50) and Pink-footed Geese (Anser brachyrhynchus; Figure 55) consume mosses to a lesser extent than the Barnacle Geese. It is interesting that moss in the diet increased as the temperature increased (Fox et al. 2006).

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Chapter 16-2: Birds and Bryophytic Food Sources

Figure 55. Anser brachyrhynchus, Pink-footed Geese, foraging among grasses. Photo by Brian Eversham, with permission.

The Barnacle Goose (Branta leucopsis; Figure 54) grazes the top layer of mosses when the Calliergon (Figure 56) is still frozen (Prop & de Vries 1993). Along the water's edge, the geese dug for large lumps of mosses, consuming them as soon as they appeared. Fortunately, the mosses were a nearly inexhaustible food supply, but the geese seemed to prefer them when they were still anchored in ice. That made it possible for them to scrape the upper, most nutritious part with their bills without having to attempt separating them from their lower parts that were sealed in ice. Grasses began to grow when the moss beds began to thaw and within one week the young leaves appeared and were immediately consumed by the geese. During the earliest stages of this thaw, the geese fed on forbs (herbaceous flowering plant other than grass) and xerophytic mosses on the few snow-free patches. Then the forbs became the dominant food for about ten days. Then the moss meadows became available and the females switched to feeding on mosses, with their forbs proportion dropping to only 50%. As they became more available, graminoids gradually took on more importance in the diet of both males and females. However, at that time the proportion of mosses in the male diet was greater than that of females, both making great use of mosses in the moss meadows for food.

Figure 56. Calliergon cordifolium, a genus that is grazed by Barnacle Geese (Branta leucopsis; Figure 54) when the moss is still encased in ice. Photo by Janice Glime.

One factor in determining suitable food is retention time (Prop & Vulink 1992). Since plant cell walls are difficult to digest, and bryophytes have a higher cell wall to cell content ratio, the bryophytes are more difficult to digest than herbaceous foods. The Barnacle Goose (Branta leucopsis; Figure 54) increased its retention time 2-4-fold as the short days of winter increased to the continuous light of summer in their Arctic breeding area. This permitted greater digestion of their food from 37% in winter to 56% in summer and allowed them to expand their food choices to include bryophytes – often the only food available in their summer range. Competition may force some geese to eat mosses. When Barnacle Geese (Branta leucopsis; Figure 54) and Pink-footed Geese (Anser brachyrhynchus; Figure 55) coexist during molting time, their diet of sedges and grasses shifts to include more mosses, especially in the Barnacle Goose, reaching 33% of the diet, whereas mosses only reached 17% of the Pink-footed Goose diet (Madsen & Mortensen 1987). The Pink-footed Goose seems to be able to keep the Barnacle Goose from feeding in the preferred sedge and grass food patches. Mosses are suboptimal for both nutrients and fiber content compared to sedges and grasses. Ardea and Sage (1982; Sage & Ardea 1982) note that the Barnacle Geese (Branta leucopsis; Figure 54) begin eating mosses as soon at they arrive in their Arctic breeding grounds. The authors suggest that this is necessary for them to build up arachidonic acid, a fatty acid in cell membranes. This notion is supported by Prins (1982). Several species of geese are known to eat mosses in their Arctic breeding grounds, including the Snow Goose (Chen caerulescens; Figure 50), Pink-footed Goose (Anser brachyrhynchus; Figure 55), Barnacle Goose, and Brant Goose (Branta bernicla; Figure 1). Prins suggested that the arachidonic acid helped to keep the membranes pliable as they move about on the frozen Arctic ground. The Canada Goose (Branta canadensis; Figure 47) instead eats horsetails (Equisetum; Figure 57), which are likewise rich in arachidonic acid, but mosses have the highest contents known.

Figure 57. Equisetum arvense, a source of arachidonic acid for Canada Goose (Branta canadensis). Photo by MPF, through Creative Commons.

Chapter 16-2: Birds and Bryophytic Food Sources

When snow melt is delayed, as it has been recently along Hudson Bay shores, a predicted outcome of global warming, as many as 100,000 Snow Geese (Chen caerulescens caerulescens; Figure 50) stay for weeks instead of 1-2 days as in the past. The result is devastation of salt marsh and wetland plants, and only the moss carpet seems able to grow. In the high Andes of sub-Antarctic South America, Attagis malouinus (White-bellied Seedsnipe; Figure 58), Chloephaga picta (Upland Goose; Figure 59), and C. poliocephala (Ashy-headed Geese; Figure 60) frequently consume bryophytes (Russo et al 2020). The fragments, including both leafy stems and capsules, occurred in 84.6% of the seedsnipe (26 samples) and 90.9% of the Chloephaga goose fecal samples (22 samples; Figure 61). At least one of the Chloephaga species consumes the mosses Polytrichum strictum (Figure 62) and Notoligotrichum trichodon (Figure 63). Of 11 collected goose droppings, more than 50% contained fragments of the Polytrichaceae. Such consumption suggests the possibility of dispersal of this moss family in bird feces.

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Figure 60. Chloephaga poliocephala, sub-Antarctic bird that eats mosses on Ushuaia, Tierra del Fuego, Argentina. Photo through Creative Commons.

Figure 61. Chloephaga feces with mosses in it. courtesy of Nick Russo, modified by Janice Glime.

Photo

Figure 58. Attagis malouinus in mountain area of Patagonia, a sub-Antarctic bird that eats mosses. Photo courtesy of Sebastian Saiter.

Figure 59. Chloephaga picta, a sub-Antarctic bird that eats mosses. Photo by Peter Prokosch, through Creative Commons.

Figure 62. Male plants of Polytrichum strictum, a common food of Attagis malouinus, Chloephaga picta, and Chloephaga poliocephala. Photo by Kristian Peters, through Creative Commons.

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Chapter 16-2: Birds and Bryophytic Food Sources

distinguish which bryophytes were being consumed, the researchers were able to identify Actinothuidium hookeri (Figure 65), Funaria hygrometrica (Figure 66), Hedwigia ciliata (Figure 67), Homomallium connexum (see Figure 68), Pogonatum perichaetiale (Figure 69), and Rhytidium rugosum (Figure 70). It appeared that the birds preferred mosses that were soft and easily fragmented for ease of swallowing. On the other hand, some of these mosses may help to grind food in the gizzard. Grasses were also eaten in large supply, but since they were abundant, it did not appear that the mosses served as emergency food or a source of fiber. Furthermore, it did not appear that the mosses were eaten as a source of insects because the insects were in low supply. Hence, it appears that the mosses were a preferred food. Figure 63. Notoligotrichum trichodon with capsules; both leafy stems and capsules are common foods of Attagis malouinus, Chloephaga picta, and Chloephaga poliocephala. Photo by Bernard Goffinet, with permission.

Blood Pheasant The Blood Pheasant (Ithaginis cruentus; Phasianidae; Figure 64) is protected in China, where it lives in shrublands on high, cold plateaus. Mosses are an important part of its diet (Shi & Li 1985; Nan et al. 2011). Yao (1992) dissected 46 gizzards to analyze for food preferences. This revealed 32 species of mosses, comprising 22 genera and 14 families. The preferred mosses comprised 24-54% of the content, second preference comprised 11-17%, third preference 4-9%, and those occasionally eaten comprised less than 2.1%.

Figure 65. Actinothuidium hookeri, food of the Blood Pheasant (Ithaaginis cruentus). Photo by Li Zhang, with permission.

Figure 64. Ithaginis cruentus, Blood Pheasant, a species for which mosses are an important diet component. Photo from EOL China Regional Center, through Creative Commons.

Other foods of the Blood Pheasant include grasses, and both mosses and grasses are taken during prolonged feeding expeditions in which the birds bob up and down like a slow sewing machine needle at the rate of 50 pecks per minute (Nan et al. 2011). In 528 observations, all individuals consumed mosses. Although it was difficult to

Figure 66. Funaria hygrometrica capsules, food for the Blood Pheasant. Photo by Frank Vincentz, through Creative Commons.

Chapter 16-2: Birds and Bryophytic Food Sources

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Figure 67. Hedwigia ciliata drying, a species eaten by the Blood Pheasant. Photo by Janice Glime. Figure 70. Rhytidium rugosum, food for the Blood Pheasant. Photo by Michael Lüth, with permission.

Kakapo On Stewart Island, the third largest island of New Zealand, the Kakapo (Strigops habroptilus; Figure 71) "plucks" the mast of the moss Dicranoloma (Figure 72), the sedge Oreobolus, the grass Centrolepis, the flowering plant Astelia, and the Asteraceae member Celmesia (Best 1984). Signs on Dicranoloma were rare, typically represented as foliage that had been pulled from the ground.

Figure 68. Homomallium incurvatum; H. connexum is among the mosses consumed by the Blood Pheasant. Photo by Hermann Schachner, through Wikiwand.

Figure 69. Pogonatum perichaetiale with capsules. This species is eaten by the Blood Pheasant. Photo by Li Zhang, with permission.

Figure 71. Strigops habroptilus, Kakapo, camouflaged among leaves in NZ. The coloration camouflages it among the vegetation, including while it feeds among bryophytes. Photo by Mnolf, through Creative Commons.

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Chapter 16-2: Birds and Bryophytic Food Sources

Figure 74. Vanellus vanellus, Northern Lapwing, a bird that consumes bryophytes. The bryophytes can remain viable in the feces. Photo by Andreas Trepte, through Creative Commons. Figure 72. Dicranoloma billardieri in NZ, a species often pulled up by the Kakapo. Photo by Jan-Peter Frahm.

Turkeys? Glover and Bailey (1949) reported that turkey droppings indicated that bryophytes formed a common food source from January to April in the beech-birchmaple-hemlock forest. However, it appears that the "mosses" in this case were instead actually Lycopodium, referred to elsewhere in the paper as a bryophyte. Dispersal The birds in some cases return the "favor." The Mallard, Anas platyrhynchos (Figure 73) and Lapwing Vanellus vanellus (Figure 74) both eat bryophytes. Wilkinson et al. (2017) found a large fragment of the moss Didymodon insulanus (Figure 75) in the feces of the Mallard in Cumbria, England, and similarly in the Lapwing feces. These fragments were cultured and proved to be viable. This suggests that consumption of bryophytes by birds can in some cases be a means of dispersal. Could this be more true for species that benefit from guano deposits?

Figure 73. Anas platyrhynchos, Mallards, birds that eat bryophytes. The mosses can remain live in the feces. Photo courtesy of Eileen Dumire.

Figure 75. Didymodon insulanus, a moss that can survive the digestive tract of Mallards and Lapwings Photo by David T. Holyoak, with permission.

Nutritional Value of Bryophytes These records raise the question of nutritional value of bryophytes. Why do birds eat bryophytes? Sugawa (1960) found that puppies and chickens will eat the pendent moss Neodicladiella pendula that is pulverized and used as a food additive. These animals seemed to suffer no ill effects. In fact, they gained more weight than the controls. Sugawa found that these mosses contained considerable Vitamin B2. Mosses can have high contents of vitamins, especially B2 (Sugawa 1960; Margaris & Kalaitzakis 1974). The greatest known use of bryophytes as food for birds occurs in the Arctic tundra. In these mosses, the caloric content is ~4.5-5.0 kcal gˉ1 (Pakarinen & Vitt 1974). The flowering plants consist of about 15% protein and 5% fats, whereas mosses have about 4% protein and 2% fats. Much of the moss biomass is bound in lignin-like compounds. Sugars in these mosses comprise ~1.5%. These sugars include mannose, melibiose, maltose, and deoxyribose in the mosses Syntrichia princeps (Figure 76), Rhynchostegium sp. (Figure 77), Platyhypnidium riparioides (Figure 78), and Homalothecium spp. (Figure 79) (Margaris & Kalaitzakis 1974).

Chapter 16-2: Birds and Bryophytic Food Sources

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Figure 76. Syntrichia princeps with capsules. Photo by Michael Lüth, with permission. Figure 79. Homalothecium lutescens Europe 2 Michael Lüth, with permission.

Figure 77. Rhynchostegium alopecuroides. Michael Lüth, with permission.

Photo by

Figure 78. Platyhypnidium riparioides with capsules, an emergent aquatic moss. Photo by Michael Lüth, with permission.

Forman (1968) examined caloric values of thirteen bryophyte species from Mt. Washington, NH, USA. Values for fresh bryophytes varied from 3747 cal g-1 dry weight for Dicranella heteromalla (Figure 80) to 4305 cal g-1 in Thuidium delicatulum (Figure 81). But then, spinach has only 0.23 cal g-1 of fresh spinach (1 cup) (Wikipedia 2017). When species were transplanted to a high-temperature and high-humidity environment, the caloric content decreased. On the other hand, bryophyte species that originated from the coniferous and northern hardwoods forests all had higher caloric values than those from the higher alpine area or the lowland oak forest. On Mt. Washington, the bryophytes are among those plants with the lowest caloric values. Mosses can affect the nutritional value of forbs and grasses in Arctic wetlands (Kotanen 2002). Moss presence did not prevent the rapid uptake of nitrogen by other forage species. However, most of added N nevertheless ended up in the moss layer. Hence, the mosses are able to divert N away from the tracheophyte forage plants and into longlasting peat. This sequestering can make it more difficult for freshwater tracheophyte forage plants to recover from excessive foraging by Snow Geese (Chen caerulescens atlantica; see Figure 50). On the other side of the coin, the Snow Geese fertilize the moss layer in the polygon fens (Pouliot 2006).

Figure 80. Dicranella heteromalla, a moss with ~3700 cal g-1 dry weight. Photo by Michael Lüth, with permission.

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Chapter 16-2: Birds and Bryophytic Food Sources

Figure 81. Thuidium delicatulum, a moss with ~4300 cal g-1 dry weight. Photo by Michael Lüth, with permission.

Solheim et al. (1996) showed that grazing geese had a significant impact on nitrogen fixation in the Arctic Svalbard. In areas with grazing there was 10X as much N fixation as in areas with no grazing. Bird droppings under cliffs likewise increased N fixation. Atmospheric pollutants are having a large impact on the N content of bryophytes. Pitcairn et al. (1995) found that atmospheric N deposition caused a significant rise in tissue N of 38% in central Scotland to 63% in Cumbria during just two decades. Crafford and Chown (1991) suggested that herbivory by curculionid beetles on bryophytes originated in response to an absence of flowering plants during glacial periods. For birds, it appears that Arctic birds that eat bryophytes likewise have occupied a feeding niche that at least during part of the year is devoid of flowering plants.

Palatability Bryologists for a long time assumed that bryophytes were inedible. This could result from bad taste, low nutrient value, or toxic effects. But, in fact, bryophytes are eaten. To humans they may taste terrible, with Crum (1973) describing Dicranum (Figure 82) as having a strong, somewhat peppery taste, Rhodobryum giganteum (Figure 83) as having a sickening sweet taste, and most tasting like raw green beans. But are these the tastes registered by the birds? Feeding preference tests of birds with choices of leafy bryophytes and capsules seem to be lacking. Are there species preferences? Does color matter? Do they provide some essential nutrient that is more abundant in bryophytes than in other foods?

Foraging As already discussed in earlier chapters, many invertebrates reside among the bryophytes. These include grubs, beetles, bugs, worms, mites, spiders, and other macroinvertebrates. Many of these organisms are desirable food for birds. Hence, many birds forage among bryophytes, and some are specially adapted for this bryophyte foraging behavior.

Figure 82. Dicranum scoparium with capsules, a moss in a genus Crum described as tasting peppery. Photo by Janice Glime.

Figure 83. Rhodobryum giganteum, a moss with a sickening sweet taste. Photo by David Long, with permission.

Ground Foragers The Common Blackbird (Turdus merula; Figure 12) forages among mosses when snow still covers part of the ground (see film by Shutterstock 2017). It is likely that other early arrivals take advantage of the moss fauna when most insects are in the egg or pupal stage, often hidden under bark or in the soil and immobile. Arctic Foraging Effects In the Arctic breeding grounds, mosses are typically the dominant vegetation. The thickness of the moss mats influence the temperature of the underlying soil (van der Wal et al. 2001). Herbivores, including birds, can reduce that mat thickness by trampling, consumption, or foraging. When Barnacle Geese (Branta leucopsis; Figure 54) and reindeer were excluded from areas with moss cover at Spitsbergen, the moss mat increased in thickness and the soil temperature was reduced by 0.9°C. In all sites, the soil temperature was negatively correlated with the thickness of the moss mat. This temperature change had no effect on the moss growth rate, but the Arctic meadow-grass (Poa arctica; Figure 84) and polar cress [Cardamine pratensis (= C. nymanii); Figure 85] experienced a 50% reduction in biomass on the chilled soils.

Chapter 16-2: Birds and Bryophytic Food Sources

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the Pacific Northwest, USA, 44% of the foraging among epiphytes was on bryophytes. These were mostly pendant bryophytes (Figure 86), followed by foliose lichens (Figure 87), then appressed bryophytes (Figure 88). In these forests, 20% of the bryophyte foraging was on the abundant moss Isothecium myosuroides (Figure 86). The bark insectivorous birds were the most frequent foraging guild on the bryophyte and lichen substrates.

Figure 84. Poa arctica, an Arctic grass that diminishes in cover at lower temperatures. Photo by R. J. Soreng, through Creative Commons.

Figure 86. Isothecium myosuroides, most common epiphytic moss foraged by birds in the Pacific Northwest. Photo by Dale Vitt, with permission.

Figure 85. Cardamine pratensis, a species that has less growth at lower soil temperatures. Photo by Aiwok through Creative Commons.

Arctic foraging can have detrimental effects on the plants in this fragile ecosystem, but at times they benefit the bryophytes. The Lesser Snow Goose (Chen caerulescens caerulescens; Figure 50) in the Arctic coastal region can be very destructive while foraging among roots and rhizomes for grubs and other food (Jefferies 1988). At the rate of foraging exhibited, Jeffries estimated that the sedge meadow would convert to a moss carpet in about five years. Foraging on Epiphytes Bryophytes are often torn up by foraging birds, presumably in search of insects and other invertebrates. In

Figure 87. Flavoparmelia caperata, a foliose lichen like those foraged by birds in the Pacific Northwest. Photo by Robert Klips, with permission.

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Chapter 16-2: Birds and Bryophytic Food Sources

Figure 88. Hypnum imponens on log, an appressed bryophyte like those that are less preferred for foraging by birds in the Pacific Northwest. Photo by Janice Glime.

As an example, we know that the Blue Tit (Cyanistes caeruleus; Figure 15) eats larvae of Erannis (Lepidoptera) in winter (Betts 1955) – a moth associated with forests with lots of bryophyte cover (Kiadaliri et al. 2005). Females of at least some species of Erannis lay eggs under mosses as well as in crevices, making this a good foraging site for birds hunting larvae. Wolf (2009) questioned the value of epiphyte foraging to birds in coniferous forests of the Pacific Northwest. Of the 735 foraging records, ~30% occurred on epiphytic substrates. The data indicated selectivity by the Chestnutbacked Chickadee (Poecile rufescens; Figure 89), Redbreasted Nuthatch (Sitta canadensis; Figure 90), Brown Creeper (Certhia americana; Figure 91), Hairy Woodpecker (Picoides villosus; Figure 92), and Gray Jay (Perisoreus canadensis; Figure 93). Furthermore, the position in the canopy influenced their choices. In the mid and upper crown, lichens were preferred, whereas in the lower crown the bryophytes were preferred. Weikel and Hayes (1999) suggested that the bryophyte cover may house more arthropods that serve as food, but at the same time they hide the arthropods, making them less available to these birds.

Figure 90. Sitta canadensis, Red-breasted Nuthatch, a species that forages among epiphytic bryophytes in the Pacific Northwest. Photo by Matt MacGillivray, through Creative Commons.

Figure 91. Certhia americana, Brown Creeper, on a tree where it often forages among mosses and lichens. Photo by Walter Siegmund, through Creative Commons.

Figure 89. Poecile rufescens, Chestnut-backed Chickadee, a species that typically forages among epiphytic bryophytes in the Pacific Northwest, USA. Photo by Walter Siegmund, through Creative Commons.

In the Pacific Northwest coniferous forests of Washington and Oregon, USA, eleven species of birds use the bryophytes for foraging (Wolf 2009). However only four bird species comprised 79% of the foraging records. These were the Pacific Winter Wren (now named Troglodytes pacificus; Figure 94; 33 records), Brown Creeper (Certhia americana; Figure 91; 13 records), Gray

Chapter 16-2: Birds and Bryophytic Food Sources

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Jay (Perisoreus canadensis; Figure 93; 14 records), and Chestnut-backed Chickadee (Poecile rufescens; Figure 89; 13 records). Among these, the Brown Creeper (Certhia americana), Hermit Thrush (Catharus guttatus; Figure 95), and Winter Wren used the bryophytes in more than 20% of their foraging excursions.

Figure 95. Catharus guttatus, Hermit Thrush, a species that frequently forages among bryophytes. Photo by Cephas, through Creative Commons.

Figure 92. Picoides villosus, Hairy Woodpecker, a species that forages among epiphytic mosses. Photo by Will Pollard, through Creative Commons.

The behavior differed among these birds (Wolf 2009). The Brown Creeper (Certhia americana; Figure 91) and Hairy Woodpecker (Picoides villosus; Figure 92) hung vertically or upside-down on the epiphytes as they probed, hammered, pecked, or otherwise inspected the epiphytic bryophytes, using mostly prostrate mosses (esp. Hypnum; Figure 96) on the bole. The arthropods that are the victims of their searches use the epiphytes for refuge, forage, rest, aestivation, and thermoregulation (Richardson & Young 1977; Rhoades 1995; Shaw 2004). The dense mats accumulate soil, providing further habitat for invertebrates (Winchester & Ring 1996). The birds contribute a selection pressure that selects for cryptic coloration and other forms of camouflage in the arthropods (Richardson & Young 1977).

Figure 93. Perisoreus canadensis, Gray Jay, a species that forages among epiphytic bryophytes. Photo by Franco Folini, through Creative Commons.

Figure 96. Hypnum cupressiforme, a common epiphytic genus for foraging by Brown Creepers and Hairy Woodpeckers. Photo by Jan-Peter Frahm, with permission.

Figure 94. Troglodytes pacificus, Pacific Wren, a forager among bryophytes. Photo by Carly Lesser & Art Drauglis, through Creative Commons.

With the wide range of bryophytes in the Neotropics, certainly some are better sources of food items than others. The Ochraceous Wren and Common Bush-Tanager forage among the dead organic matter and bryophytes more frequently than they do among other (tracheophyte) epiphytes (Nadkarni & Matelson 1989). In Costa Rica, The Ruddy Treerunner (Margarornis rubiginosus; Figure 97) is an epiphyte specialist, foraging on bryophytes (Sillett 1994). The Spot-crowned Woodcreeper (Lepidocolaptes affinis; Figure 98) is a Central American foraging specialist on bryophytes and

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foliose lichens, but the bryophytes were used less proportionately than lichens.

Figure 99. Blue-capped Ifrita, Ifrita kowaldi, a poisonous bird that lives in mossy forests where it forages among midstory mosses. Photo by Jerry Oldenettel, through Creative Commons.

Figure 97. Margarornis rubiginosus, Ruddy Treerunner, a species that specializes on foraging among bryophytes. Photo by Dominic Sherony, through Creative Commons.

Figure 100. New Guinea Highlands, Papua New Guinea. Photo from eGuide Travel, through Creative Commons.

Figure 98. Lepidocolaptes affinis, Spot-crowned Woodcreeper, foraging among mosses. Photo by Carmelo López Abad, through Creative Commons.

The Blue-capped Ifrita (Ifrita kowaldi; Figure 99), a poisonous bird, is restricted to the highlands of New Guinea (Figure 100), mostly above 2000 m asl (Dumbacher et al. 2000). They live in mossy, moist montane forests, where they behave much like the nuthatches, foraging for insects and worms among mosses, on tree trunks, and on major branches in the midstory of the forest. They are rarely seen alone, typically travelling in groups of up to six individuals.

Pendant bryophytes (Figure 101) can protect some arthropods from foragers. These arthropods are able to dwell at some distance from the branch, away from the perches of the birds (Wolf 2009). These mosses are too unstable for many kinds of birds to perch. Among the birds that were not deterred by the pendant branches, the Pacificslope Flycatcher (Empidonax difficilis; Figure 102) used a sally, hover, and glean foraging behavior to capture insects on the dangling bryophytes. The Chestnut-backed Chickadee (Poecile rufescens; Figure 89) used short flights and hops to forage, but occasionally hovered or hung from the bryophytes to snatch an insect from the pendant portion. Furthermore, 70% of the nests of this species contained bryophytes (Dahlsten et al. 2002). Peterson et al. (1989) sampled trunk-surface arthropods from American beech (Fagus grandifolia; Figure 103) and sugar maple (Acer saccharum; Figure 104). The arthropod resources did not differ significantly between trees. Furthermore, they were not correlated with bark texture or bryophyte cover.

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Figure 101. Pseudobarbella mollisima, a pendant moss in Japan. Photo by Janice Glime. Figure 104. Acer saccharum autumn leaves and trunk. Photo by Janice Glime.

Figure 102. Empidonax difficilis, Pacific-slope Flycatcher, a species that is able to forage among dangling mosses. Photo by Ron Knight, through Creative Commons.

Figure 105. Phasianus colchicus, Pheasant, a species that often disturbs bryophytes while foraging. Photo by Gary Noon, through Creative Commons.

Figure 103. Fagus grandifolia forest in winter. Photo by Dcrjsr, through Creative Commons.

Pheasants (Phasianus colchicus; Figure 105) do not seem to have any particular use for the mosses themselves, but the mosses seem to be in their way on the forest floor of a wetland forest (Wiegers 1983). When they are foraging, they turn the bryophyte cover upside down in search of food. Following these events, some mosses, including Dicranum scoparium (Figure 106) and Mnium hornum (Figure 107), that were turned upside down develop into moss balls.

Figure 106. Dicranum scoparium, a moss that gets turned upside down by foraging pheasants. Photo by J. C. Schou, through Creative Commons.

Rod Seppelt (Bryonet 26 February 2013) has observed Skuas (Catharacta lonnbergi; Figure 108) upturning upland moss polsters of Ditrichum strictum (see Figure 109) on subAntarctic islands, searching for earthworms. It is puzzling because there are easier food items available than these relatively small worms.

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Figure 107. Mnium hornum, a moss that gets turned upside down by foraging pheasants. Photo by Kristian Peters, through Creative Commons.

Figure 110. Cyanocitta stelleri, Steller's Jay, a species that forages on mosses on oaks in the Pacific Northwest, USA. Photo by Alan D. Wilson, through Creative Commons.

Figure 108. Catharacta lonnbergi, Skua, on nest on South Georgia, a species that upturns mosses to forage. Photo by Christo Barrs, through Creative Commons.

Figure 111. Aphelocoma californica, Scrub Jay, a species that tears up mosses on oak trees. Photo by Minette Layne, through Creative Commons.

Figure 109. Ditrichum gracile; D. strictum is commonly upturned by foraging Skuas on sub-Antarctic islands. Photo by Hermann Schachner, through Creative Commons.

In Eugene, Oregon, USA, the Steller's Jay (Cyanocitta stelleri; Figure 110) tears up mosses from the oaks as it forages for arthropods that hide there (Wagner 2013). In other locations it is Crows (Figure 112) and Scrub Jays (Aphelocoma californica; Figure 111).

Crows (Corvus; Figure 112) are among those birds that can be quite destructive to bryophytes. Erkamo (1976) reported that some animal had upturned mosses on flat, open rocks in Finland. These mosses were typically only a few cm across, but some were up to 10-15 cm. Since the observations are indirect, based only on the upturned mosses, it is possible that voles, pheasants, seagulls, or crows were responsible, but crows seemed most likely. Erkamo has, at other times, seen crows engaging in such activity, presumably searching for insects or worms. Birds keep bryophytes from growing well on red wood ant (Formica rufa group; Figure 113) mounds due to the bird foraging activity on the ants (Heinken et al. 2007). Motley and Bosanquet (2004) reported a neglected flower pot that contained Petalophyllum ralfsii (Figure 114). Meanwhile, the surface had been colonized by various species of moss and the thallose liverwort Aneura

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(Figure 115). The surprise came when birds attacked the bryophytes, pulling them out and most likely taking them for nesting material. But they were selective. They avoided taking the P. ralfsii.

Figure 115. Aneura pinguis, a bryophyte among those collected by birds, presumably for nesting material. Photo by Michael Lüth, with permission.

Figure 112. Corvus corax, Crow, a species that is destructive of bryophytes while foraging. Photo by Ingrid Taylar, through Creative Commons.

Juncos The Dark-eyed Junco (Junco hyemalis; Figure 116) in the Pacific Northwest, USA, is most active in the low understory, but it may go to the upper canopy to search for prey items among the lichens (Wolf 2009). But they may also forage on Dicranum sp. (Figure 82, Figure 106) and Isothecium (Figure 86), where Wolf observed them on a horizontal tree bole and branch of Tsuga heterophylla (Figure 117) at 0.7 m and 3 m respectively.

Figure 113. Formica rufa sideview, an ant that builds mounds and birds keep bryophytes from growing on them. Photo by Richard Bartz, through Creative Commons. Figure 116. Junco hyemalis, Dark-eyed Junco, a species that forages on Dicranum sp. and Isothecium. Photo by Factumquintus, through Creative Commons.

Figure 114. Petalophyllum ralphsii, a species that is avoided when birds collect bryophytes for nests. Photo by Michael Lüth, with permission.

Figure 117. Tsuga heterophylla (hemlock) forest, home of the Dark-eyed Junco. Photo by Willow & Monk, through Creative Commons.

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Weaver Birds In the Udzungwa Mountains of Tanzania, the disturbed humid forest serves as home for at least 70 species of birds (Fjeldså 1999). Many of the birds search for their food among the epiphytic lichens, mosses, and ferns in the mature forests. The Tasmanian Mountain Weaver, Ploceus nicolli (Figure 118), is a vulnerable species that occurs in the tall forest of the Eastern Arc Mountains. It is associated with locations having large cover of epiphytic mosses and lichens.

Table 1. Percentage (and total number) of foraging visits to epiphytes by birds that probed moss mats and dead organic matter in the Monteverde field study, 1 July to 28 August 1985. Frequent foragers had 10 or more foraging visits recorded during the study period. Infrequent foragers had less than 10 foraging visits recorded. From Nadkarni and Matelson (1989).

Frequent foraging visits (> 10 foraging visits) White-throated Mountain-gem, Lampornis castaneoventris95 (150) Ochraceous Wren, Troglodytes ochraceus 89 (19) Common Bush anager, Chlorospingur ophthalmicus 57 (511) Olive-striped Flycatcher, Mionectes olivaceus 46 (37) Slate-throated Redstart, Myioborus miniatus 45 (47) Prong-billed Barbet, Semnornis fiantzii 30 (23) Golden-browed Chlorophonia, Chlorophonia callophrys 33 (187) House Wren, Troglodytes aedon 26 (57) Three-striped Warbler, Basileuterus tristriatus 20 (10) Mountain Robin, Turdus plebejus < 10 (146)

Infrequent foragers (< 10 total foraging visits) Spotted Barbtail, Premnoplex brunnescens

Figure 118. Ploceus velatus, Southern Masked Weaver and nest; P. nicolli lives in areas with a large cover of epiphytic mosses. Photo by Chris Eason, through Creative Commons.

Tropical Birds In the tropics, some birds use epiphytes as their feeding substrates. These include at one end of the spectrum those birds that choose the substrate where they prefer to feed, and at the other end the birds choose the prey item, going to the substrate if it potentially has that prey organism. In Costa Rica, Sillett (1994) studied eight species that use epiphytes among their feeding substrates. Four species were epiphyte specialists. These included two that chose bryophytes: Ruddy Treerunner (Margarornis rubiginosus; Furnariidae; Figure 97) on just bryophytes and Spot-crowned Woodcreeper (Lepidocolaptes affinis; Dendrocolaptidae; Figure 98) on bryophytes and lichens. Orians (1969) and Remsen (1985) have provided evidence of bryophyte utilization by tropical birds, but otherwise, little documentation of this tropical resource exists. In Neotropical Costa Rica, Nadkarni and Matelson (1989) report three birds that feed upon bryophyte inhabitants (Table 1). The Emerald-chinned Hummingbird (Abeillia abeillei; Figure 119) and Amethyst-throated Hummingbird (Lampornis amethystinus; Figure 120) feed upon insects associated with the mosses and other bryophytes. The Rufous-tailed Hummingbird (Amazilia tzacatl; Figure 121) utilizes the flowers that are anchored in the bryophytic substrate. In fact, the Ochraceous Wren (Troglodytes ochraceus; Figure 122) and Common BushTanager (Chlorospingus ophthalmicus; Figure 123) foraged in mosses more frequently than expected. Avian resources nestled among the bryophyte mats include fruits, flowers, seeds, water, and invertebrates.

Figure 119. Abeillia abeillei, Emerald-chinned Hummingbird, a tropical bird that feeds on insects associated with bryophytes. Photo by Scott Bowers, through Creative Commons.

Figure 120. Lampornis amethystinus, Amethyst-throated Hummingbird, a tropical bird that feeds on insects associated with bryophytes. Photo by Juan Carlos Pérez M., through Creative Commons.

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In subtropical evergreen forests, Dinesen (1995, 1997) reported on Shelley's Greenbul (Arizelocichla masukuensis; Figure 124). These birds found most of their food among the epiphytic mosses.

Figure 121. Amazilia tzacatl, Rufous-tailed Hummingbird, a bird that feeds on flowers that are anchored in bryophytes. Photo by Brian Gratwicke Creative Commons.

Figure 124. Shelley´s Greenbul, Arizelocichla masukuensis, a species that forages among epiphytic mosses. Photo by Per Holmen, with permission.

Jamaican Blackbird Another tropical bird, the Jamaican Blackbird, Nesopsar nigerrimus (Figure 125), lives in the moist montane of Jamaica above 515 m (Cruz 1978). Its food includes insects, and its foraging behavior among the epiphytes, dead leaves, and moss-covered tree trunks and branches seems to be part of its adaptive evolution on the island. Its shorter legs, more curved claws, and longer, narrower bill adapt it for arboreal rummaging in crevices and among bryophytes. Figure 122. Troglodytes ochraceus, Ochraceous Wren, on mosses, a location where it forages. Photo by Annika Lindqvist, through Creative Commons.

Figure 125. Nesopsar nigerrimus, Jamaican Blackbird, foraging amid lichens. Photo by Dominic Sherony, through Creative Commons.

Summary Figure 123. Chlorospingus ophthalmicus, Common Bush Tanager, on bryophytes where it forages. Photo by Cephas, through Creative Commons.

Both capsules and leafy portions of bryophytes are eaten by some birds. This is particularly true in polar climates where tracheophytes are scarce or absent. These birds include grouse and pheasants, as well as

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song birds. Even some parrots feed on capsules of Polytrichum. In tundra regions, the ptarmigan and grouse chicks often depend on bryophytes, especially the high quality food of capsules. Some birds use bryophyte capsules as emergency food, and one might describe all use of bryophytes as emergency food, although in some habitats, the emergency is long-lived. This capsule feeding can be seasonal, can depend on a bad year for tracheophytes, or can be used in a habitat with low productivity. Use of color by birds to locate food is a topic wide open for research. Several hypotheses have suggested that members of the Splachnaceae with their brightly colored capsules and fruity odors may get dispersed as a result of attracting birds. This may also occur for the moss Pleurophascum. The ability of most songbirds and some others may enable the birds to see UV reflections that we have not discovered for capsules, or to locate bulbils and other bryophyte structures. Leafy plants may be eaten as well, including by some diving birds and ptarmigans. Blood Pheasants, in particular, seem to consume large quantities of leafy bryophytes. In other cases, antiherbivory compounds keep the birds away, protecting the invertebrates living among the bryophyte branches. On the other hand, bryophytes may provide high concentrations of some vitamins, and one study on caloric content indicates that levels in leafy bryophytes may be high. Bryophytes can compete for nutrients, especially nitrogen, making the forbs less nutritious. Some birds may use the bryophytes to obtain arachidonic acid in preparation for winter. The high ratio of cell wall to cell contents requires a long retention time of consumed bryophytes. This can reduce the feeding rate, causing the birds to remain quiet and less conspicuous. On the other hand, it might provide the bryophytes with a means of long-distance dispersal; some bryophytes survive passage through the digestive tract. Perhaps the greatest food contribution of the bryophytes is through foraging. Many invertebrates reside there. This can be good or bad for the birds, with some specializing on bryophyte foraging and others unable to locate the invertebrates hidden by the bryophytes. Among these, the hanging bryophytes require the greatest specialization by the bird foragers, thus providing a safe haven for many invertebrates. On the other hand, the birds disturb the bryophytes on the ground and elsewhere, providing possible dispersal.

Acknowledgments Thank you to Brian Dykstra for sending me the wonderful thesis on birds and epiphytes by Adrian Wolf, as well as other references and personal observations. Thank you to Marcel Schrijvers-Gonlag for supplying me with the pertinent part of the Haftorn (1954) paper. David Dumond shared the references he got from Bryonet. Bernard Goffinet has kindly alerted me to new literature on the subjectt and sent me an advance copy of his paper on birds

eating sporophytes in the sub-Antarctic. Thank you to Janet Marr for a critical reading of the manuscript.

Literature Cited Ardea and Sage, B. 1982. Why the Arctic dwellers gather moss. New Sci. 95: 152. Bennett, A. T. and Cuthill, I. C. 1994. Ultraviolet vision in birds: what is its function? Vision Res. 34: 1471-1478. Bennett, A. T., and Théry, M. 2007. Avian color vision and coloration: multidisciplinary evolutionary biology. Amer. Nat. 169: S1-S6. Bennett, A. T. D., Cuthill, I. C., and Norris, K. J. 1994. Sexual selection and the mismeasure of color. Amer. Nat. 144: 848860. Best, H. A. 1984. The foods of Kakapo on Stewart Island as determined from their feeding sign. N. Zeal. J. Ecol. 7: 7183. Betts, M. M. 1955. The food of titmice in oak woodland. J. Anim. Ecol. 24: 282-323. Braun, C. E., Martin, K., and Robb, L. A. 1993. White-tailed Ptarmigan. In: Poole, A., Stettenheim, P., and Gill, F. (eds.). The Birds of North America, No 68, The Academy of Natural Sciences, Philadelphia, The American Ornithologists' Union, Washington, DC. Crafford, J. E., and Chown, S. L. 1991. Comparative nutritional ecology of bryophyte and angiosperm feeders in a sub‐Antarctic weevil species complex (Coleoptera: Curculionidae). Ecol. Entomol. 16: 323-329. Crum, H. 1973. Mosses of the Great Lakes Forest. Contrib. Univ. Mich. Herb. 10: 1-404. Cruz, A. 1978. Adaptive evolution in the Jamaican Blackbird Nesopsar nigerrimus. Ornis Scand. 9: 130-137. Dahlsten, D. L., Brennan, L. A., McCallum, D. A., and Gaunt, S. L. 2002. Chestnut-backed Chickadee (Poecile rufescens). In: Poole, A. and Gill, F. (eds.). The Birds of North America, no. 689. Academy of Natural Sciences; Philadelphia, and American Ornithologists’ Union, Washington, D. C. Dinesen, L. 1995. Seasonal variation in feeding ecology of Shelley's Greenbul in subtropical evergreen forests. Afr. J. Ecol. 33: 420-425. Dinesen, L. 1997. Succeshistorier blandt afrikanske regnskovsfugle. Naturens Verden 1997: 178-186. Dumbacher, J. P., Spande, T. F., and Daly, J. W. 2000. Batrachotoxin alkaloids from passerine birds: A second toxic bird genus (Ifrita kowaldi) from New Guinea. Proc. Natl. Acad. Sci. USA 97: 12970-12975. Erkamo, V. 1976. Warikset kallioiden sammalpeitteen turmelijoina. [Crows disturbing the moss cover of rocks in Helsinki.]. Luonnon Tutkija 80(2): 57-58. Finger, E. and Burkhardt, D. 1994. Biological aspects of bird colouration and avian colour vision including ultraviolet range. Vision Res. 34: 1509-1514. Fjeldså, J. 1999. The impact of human forest disturbance on the endemic avifauna of the Udzungwa Mountains, Tanzania. Bird Conserv. Internat. 9: 47-62. Forman, R. T. 1968. Caloric values of bryophytes. Bryologist 71: 344-347. Fox, A. D. and Bergersen, E. 2005. Lack of competition between barnacle geese Branta leucopsis and pink-footed geese Anser brachyrhynchus during the pre-breeding period in Svalbard. J. Avian Biol. 36: 173-178.

Chapter 16-2: Birds and Bryophytic Food Sources

Fox, A. D., Francis, I. S., and Bergersen, E. 2006. Diet and habitat use of Svalbard Pink-footed Geese Anser brachyrhynchus during arrival and pre-breeding periods in Adventdalen. Ardea 94: 691-699. Gardarsson, A. and Moss, R. 1970. Selection of food by Icelandic ptarmigan in relation to its availability and nutritive value. In: Watson, A. (ed.). Animal populations in relation to their food resources. Blackwell Scientific Publications, London, U.K, pp. 41-71. Gloutney, M. L., Alisauskas, R. T., Afton, A. D., and Slattery, S. M. 2001. Foraging time and dietary intake by breeding Ross's and Lesser Snow Geese. Oecologia 127: 78-86. Glover, F. A. and Bailey, R. W. 1949. Wild Turkey foods in West Virginia. J. Wildlf. Mgmt. 13: 255-265. Haftorn, S. 1954. Contribution to the food biology of tits especially about storing of surplus food. Part I. The Crested Tit (Parus c. cristatus L.). K. norske vidensk. Selsk. Skr. 1953 4: 1-122. Hannon, S. J., Martin, K., and Schieck J. O. 1988. Timing and reproduction in two populations of Willow Ptarmigan in northern Canada. Auk 105: 330-338. Håstad, O., Victorsson, J., and Ödeen, A. 2005. Differences in color vision make passerines less conspicuous in the eyes of their predators. Proc. Natl. Acad. Sci. USA 102: 6391-6394. Heinken, T., Rohner, M.-S., and Hoppert, M. 2007. Red wood ants (Formica rufa group) disperse bryophyte and lichen fragments on a local scale. Nova Hedw. 131: 147-163. Hohman, W. L. 1985. Feeding ecology of ring-necked ducks in northwestern Minnesota. J. Wildlf. Mgmt. 49: 546-557. Jefferies, R. L. 1988. Pattern and process in Arctic costal vegetation in response to foraging by lesser snow geese. In: Werger, M. J. A., Aart, P. J. M. van der, During, H. J., and Verhoeven, J. T. A. (eds.), Plant Form and Vegetative Structure, SPB Academic Publishing, The Hague, The Netherlands, pp. 281-300. Kiadaliri, H., Ostovan, H., Abaei, M., and Ahangaran, Y. 2005. Investigation on the behaviour treat of leaf feeder moth (Erannis defoliaria Clerck) and natural enemies in forests of the in west of Mazandaran province. J. Agric. Sci. 11: 145159. Kotanen, P. M. 2002. Fates of added nitrogen in freshwater Arctic wetlands grazed by snow geese: The role of mosses. Arct. Antarct. Alp. Res. 34: 219-225. Lid, J. and Meidell, O. 1933. The food of Norwegian Grouse chicks (Lagopus lagopus L.). Nytt. Mag. Naturvidensk. 73: 75-114. Longton, R. E. 1992. The role of bryophytes and lichens in terrestrial ecosystems. In: Bates, J. W. and Farmer, A. M. (eds.). Bryophytes and Lichens in a Changing Environment. Clarendon Press, Oxford, pp. 32-76. Madsen, J. and Mortensen, C. E. 1987. Habitat exploitation and interspecific competition of moulting geese in East Greenland. Ibis 129: 25-44. Margaris, N. S. and Kalaitzakis, J. 1974. Soluble sugars from some Greek mosses. Bryologist 77: 470-472. Martin, K. and Hik, D. 1992. Willow Ptarmigan chicks consume moss sporophyte capsules (Polluelos de Lagopus lagopus Consumen Cápsulas de Esporofitos de Musgos). J. Field Ornithol. 63: 355-358. Motley, G. S. and Bosanquet, S. D. S. 2004. Petalophyllum ralfsii inland in Carmarthenshire (and in a plant pot in Monmouthshire!). Field Bryol. 83: 15-17. Nadkarni, N. M. and Matelson, T. J. 1989. Bird use of epiphyte resources in neotropical trees. Condor 91: 891-907.

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Nan, W., Tan, B. C., Davison, G. W. H. 2011. Bryophytes as a major dietary component for the Blood Pheasant, Ithaginis cruentus. J. Bryol. 33: 257-260. Naylor, B. J. and Bendell, J. F. 1989. Clutch size and egg size of spruce grouse in relation to spring diet, food supply, and endogenous reserves. Can. J. Zool. 67: 969-980. Nordhorn-Richter, G. 1984. Primary fluorescence of mosses. Leitz-Mitt. Wiss. Tech. 8: 167-170. Ödeen, A. and Håstad, O. 2003. Complex distribution of avian color vision systems revealed by sequencing the SWS1 opsin from total DNA. Molec. Biol. Evol. 20: 855-861. Orians, G. H. 1969. The number of bird species in some tropical forests. Ecology 50: 783-801. Pakarinen, P. and Vitt, D. H. 1974. The major organic components and calorie contents of high Arctic bryophytes. Can. J. Bot. 52: 1151-1161. Palmer, R. S. 1962. Handbook of North American birds. Vol 1, Loons through Flamingos. Yale University Press, New Haven, CT. Peterson, A. T., Osborne, D. R., and Taylor, D. H. 1989. Tree trunk arthropod faunas as food resources for birds. Ohio J. Sci. 89: 23-25. Pitcairn, C. E. R., Fowler, D., and Grace, J. 1995. Deposition of fixed atmospheric nitrogen and foliar nitrogen content of bryophytes and Calluna vulgaris (L.) Hull. Environ. Pollut. 88(2): 193-205. Pouliot, R. 2006. New IAB members present themselves: Rémy Pouliot. Bryol. Times 121: 4. Prins, H. H. 1982. Why are mosses eaten in cold environments only? Oikos 38: 374-380. Prop, J. and Vries, J. de. 1993. Impact of snow and food conditions on the reproductive performance of Barnacle Geese Branta leucopsis. Ornis Scand. 24: 110-121. Prop, J. and Vulink, T. 1992. Digestion by Barnacle Geese in the annual cycle: The interplay between retention time and food quality. Funct. Ecol. 6: 180-189. Prop, J., Eerden, M. R. van, Daan, S., Drent, R. H., Tinbergen, J. M., and Joseph, A. M. St. 1980. Ecology of the barnacle goose (Branta leucopsis) during the breeding season: Preliminary results from expeditions to Spitzbergen in 1977 and 1978. In: Proc. Norwegian-Netherlands Symp. Svalbard. Arctic Centre, Univ. of Groningen. Pullianen, E., and Eskonen, H. 1982. Chemical composition of plant matter eaten by young chicks of the Willow Grouse Lagopus lagopus in northern Finland. Ornithol. Fenn. 59: 146-148. Remsen, J. V. Jr. 1985. Community organization and ecology of birds of high elevation humid forest of the Bolivian Andes. Ornithol. Monogr. 36: 733-756. Rhoades, F. M. 1995. Nonvascular epiphytes in forest canopies: Worldwide distribution, abundance and ecological roles. In: Lowman, M. D. and Nadkarni, N. M. (eds.). Forest Canopies. Academic Press, San Diego, pp 353-408. Richardson, D. H. S. 1981. The Biology of Mosses. Blackwell, Oxford. Richardson, D. H. and Young, C. M. 1977. Lichens and vertebrates. In: Seward, M. R. D. (ed.). Lichen Ecology. Academic Press, London, pp. 121-144. Russo, N. J., Robertson, M., MacKenzie, R., Goffinet, B., and Jiménez, J. E. 2020. Evidence of targeted consumption of mosses by birds in sub-Antarctic South America. Austral Ecol. 45: 399-403.Sage, B. and Ardea. 1982. Why the Arctic dwellers gather moss. New Sci. 95: 152.

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Shaw, D. C. 2004. Vertical organization of canopy biota. In: Lowman, M. D. and Rinker, H. B. (eds.). Forest Canopies. Elsevier Academic Press, Cambridge, MA., pp. 73-101. Shi, D.-C. and Li, G.-H. 1985. A preliminary study on the Blood Pheasant diet at Baihe Reserve, Nanping, Sichuan. Zool. Res., Kuming 6(2): 137-145. Shutterstock. 2017. Common Blackbird (Turdus merula) on moss looking for food. Accessed 3 July 2017 at . Sillett, T. S. 1994. Foraging ecology of epiphyte-searching insectivorous birds in Costa Rica. Condor 96: 863-877. Solheim, B., Endal, A., and Vigstad, H. 1996. Nitrogen fixation in Arctic vegetation and soils from Svalbard, Norway. Polar Biol. 16: 35-40. Spidsø, T. K. 1980. Food selection by Willow Grouse Lagopus lagopus chicks in northern Norway. Ornithol. Scand. 11: 99105. Sugawa, S. 1960. Nutritive value of mosses as a food for domestic animals and fowls. Hikobia 2: 119-124. Wagner, David. 2013. It's about time. EugeneWeekly.com. 14 February 2013 available at 5000 species) and comprises most of the birds that use bryophytes in their nests. But then, it also includes more than half the bird species in the world (Wikipedia 2017). The order is distinguished by having three toes pointing forward and one pointing back, permitting these to be perching birds. Passerines also are altricial (hatched or born in undeveloped state and requiring care and feeding by parents). Richardson (1981) reports that a quarter of the bird species breeding in Great Britain use bryophytes in the construction of their nests. Hansell (2000) likewise reports that numerous small to medium-sized bird species use bryophytes. Large passerine birds tend to build larger nests relative to their body size when compared to small birds (Slagsvold 1989). The depth of the inner nest cup size of these birds does not relate to the size of the bird. Birds that nest off the ground in open nests have a narrow nest cup, but those with a domed nest or which build in a cavity have a broad nest cup. Birds in exposed nests are less likely to survive than those reared in nest cavities (Nice 1937, 1957). There

seem to be no data on the success of birds reared in nests made totally of mosses. Mosses and lichens alter the nest cup size, with the inner nest cup being narrower when more are used (Slagsvold 1989). Use of mosses and lichens also depends on season, with those birds nesting early in the breeding season using significantly more mosses and lichens than are used in later nests. In coniferous forests, bryophytes are often abundant. Several species of birds that breed there build nests exclusively of bryophytes. These include the Winter Wren (see below; Hejl et al. 2002), Marbled Murrelet (see Chapter 16-7; Nelson 1997), and Golden-crowned Kinglet (see Chapter 16-7; Ingold & Galati 1997). In addition, Sakai (1988) described a Hammond Flycatcher nest (see below) made with two epiphytic lichens and five bryophytes, including the epiphytic moss Isothecium sp. (Figure 11) and liverwort, Porella navicularis (Figure 17). Tyrannidae – Tyrant Flycatchers Wolf (2009) found fifteen species of Tyrannidae that use bryophytes in their nests in North America:

Chapter 16-6: Bird Nests – Passeriformes, part 1

Contopus sordidulus (Western Wood-Pewee; Figure 2) Empidonax flaviventris (Yellow-bellied Flycatcher; Figure 4) Empidonax alnorum (Alder Flycatcher; Figure 5) Empidonax minimus (Least Flycatcher; Figure 6) Empidonax difficilis (Pacific-slope Flycatcher; Figure 7-Figure 8) Empidonax hammondii (Hammond's Flycatcher; Figure 13) Empidonax occidentalis (Cordilleran Flycatcher; Figure 18) Sayornis nigricans (Black Phoebe; Figure 19) Sayornis phoebe (Eastern Phoebe; Figure 20-Figure 21) Sayornis saya (Say's Phoebe; Figure 26-Figure 27) Pitangus sulphuratus (Great Kiskadee; Figure 28) Tyrannus melancholicus (Tropical Kingbird; Figure 31) Tyrannus couchii (Couch's Kingbird; Figure 32) Tyrannus forficatus (Scissor-tailed Flycatcher; Figure 33) Pachyramphus aglaiae (Rose-throated Becard; Figure 37)

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Yellow-bellied Flycatcher (Empidonax flaviventris) In the eastern United States, Yellow-bellied Flycatcher (Empidonax flaviventris; Figure 4) nests close to mature stands of lowland coniferous forest (Harrison 1975; Hawrot & Niemi 1996). These forests often have a well-developed layer of mosses and these mosses appear to be necessary for the bird's nesting. The Yellow-bellied Flycatcher nests on the ground in a layer of mosses.

Figure 4. Empidonax flaviventris, Yellow-bellied Flycatcher. This species builds nests on a bed of mosses on the ground. Photo by Cephas, through Creative Commons.

Figure 2. Contopus sordidulus, Western Wood Pewee. Members of this species often include mosses in their nests. Photo by Tom Grey, with permission.

Olive-sided Flycatcher (Contopus cooperi) The Olive-sided Flycatchers (Contopus cooperi; Figure 3) typically hide their nests in a cluster of needles and twigs at distal ends of horizontal conifer branches (Johnsgard 2009). These may occur anywhere from 5-13 m above the ground. They use twigs, lichens, mosses, and needles to construct a cup ~12-15 cm in diameter.

Figure 3. Contopus cooperi, Olive-sided Flycatcher. Members of this species often include mosses in their nests. Photo by Jerry Oldenettel, through Creative Commons.

Figure 5. Empidonax alnorum, Alder Flycatcher. Members of this species often include mosses in their nests. Photo by Cephas, through Creative Common.

Figure 6. Empidonax minimus, Least Flycatcher. Members of this species often include mosses in their nests. Photo by MDF, through Creative Commons.

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Pacific-slope Flycatcher (Empidonax difficilis) The Pacific-slope Flycatcher (Empidonax difficilis; Figure 7-Figure 8) typically builds nests on ledges or crevices of canyon walls (Johnsgard 2009). These are often concealed by mosses or ferns. When the nest is built on trees, they are supported from below and from the rear, occurring in a crotch or on a limb that projects far from the main trunk. They contain a variety of materials, frequently including mosses (Figure 8-Figure 9).

Figure 9. Empidonax difficilis, Pacific-slope Flycatcher, nest with mosses and young bird. Photo by Don Loarie, through Creative Commons.

Figure 7. Empidonax difficilis, Pacific-slope Flycatcher, a species that uses Isothecium in their nests in Douglas fir forests of the Pacific Northwest, USA. Photo by Ron Knight, through Creative Commons.

Figure 8. Empidonax difficilis, Pacific-slope Flycatcher mossy nest with eggs. Photo from USFWS, through Creative Commons.

In the Pacific Northwest, USA, Wolf (2009) found a nest of the Pacific-slope Flycatcher (Figure 8) on a fractured piece of bark on the tree bole of Pseudotsuga menziesii (Figure 10) at ~4 m above the ground. The bird had woven strands of the moss Isothecium (Figure 11) into the rim of the nest and decorated the exterior with fragments of the lichen Sphaerophorus globosus (Figure 12). The Isothecium had been relocated from elsewhere in the forest understory.

Figure 10. Pseudotsuga menziesii bark where Pacific-slope Flycatchers (Empidonax difficilis) build their nests in crevices. Photo by Walter Siegmund, through Creative Commons

Chapter 16-6: Bird Nests – Passeriformes, part 1

Figure 11. Isothecium myosuroides, representing a genus among those used in nests of the Pacific-slope Flycatcher (Empidonax difficilis). Photo by Hermann Schachner, through Creative Commons.

Figure 12. Sphaerophorus globosus, one of the lichen materials used in nests of the Pacific-slope Flycatcher (Empidonax difficilis). Photo by Einar Timdal, through Creative Commons.

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Figure 13. Empidonax hammondii, Hammond's Flycatcher. Members of this species often include mosses in their nests. Photo by Pablo Leautaud, through Creative Commons.

Figure 14. Dendroalsia abietina, a nest component of the Hammond's Flycatcher in the Pacific Northwest. Photo by James Maughn, through Creative Commons.

Hammond's Flycatcher (Empidonax hammondii) The Hammond's Flycatcher (Empidonax hammondii; Figure 13) has a nest that is distinctly different from that of the Pacific Slope Flycatcher (Empidonax difficilis; Figure 7-Figure 9) (Sakai 1988). The Hammond's Flycatcher nest is taller, more tightly woven, and mimics the surrounding substrate. The outer bowl of the only retrieved nest was made with mostly white scale lichens, mosses Dendroalsia abietina (Figure 14), Homalothecium nuttallii (Figure 15), Isothecium sp. (Figure 11), Alsia sp. (Figure 16), and the leafy liverwort Porella navicularis (Figure 17). By comparison, in the 22 Pacific-slope Flycatcher nests, the material was mostly mosses. They often lacked the camouflage effect because they used the same materials on all substrates. The nests were held together with spider webs that were also used to secure the nests to the substrate.

Figure 15. Homalothecium nuttallii, a species used in nests of the Hammond's Flycatcher in the Pacific Northwest. Photo by Doug Murphy, through Creative Commons.

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Chapter 16-6: Bird Nests – Passeriformes, part 1

Figure 19. Sayornis nigricans, Black Phoebe. Members of this species often include mosses in their nests. Photo by Tom Grey, with permission. Figure 16. Alsia californica, member of a genus used in nests of the Hammond's Flycatcher in parts of North America. Photo by John Game, through Creative Commons.

Figure 17. Porella navicularis, a leafy liverwort used in nests of the Hammond's Flycatcher in the Pacific Northwest. Photo by Matt Goff , with permission.

Eastern Phoebe (Sayornis phoebe) I picked up my copy of "A Complete Field Guide to Nests in the United States" with eager anticipation. I quickly scanned the keys that depended on nesting location and materials and found several that mentioned mosses or peatlands. As I looked up each appropriate item in the key, I soon discovered only one bird was cited as a bryophyte user, the Eastern Phoebe – Sayornis phoebe (Figure 20) (Headstromn 1970). The Eastern Phoebe builds a cupshaped nest (Figure 21) lined with mud and fibrous plant material. It uses mosses as a binding material with mud in the inner cup (Breil & Moyle 1976). It also uses mosses to line the cup. The outermost layer is also covered with moss (Headstromn 1970). Bent (1963) provided interesting bryological information. In a single nest, Mnium stellare (Figure 22), Funaria sp. (Figure 23), Polytrichum sp. (Figure 24), Hypnum "cristatum," and Climacium dendroides (Figure 25) were used as construction materials.

Figure 18. Empidonax occidentalis, Cordilleran Flycatcher. Members of this species often include mosses in their nests. Photo from Amado Demesa, through Creative Commons.

Figure 20. Sayornis phoebe, Eastern Phoebe, a bird that can be identified by the mosses in its nest. Photo by John Benson, through Creative Commons.

Chapter 16-6: Bird Nests – Passeriformes, part 1

Figure 21. Sayornis phoebe, Eastern Phoebe, nest. Photo by Bernard Goffinet, through Creative Commons.

Figure 22. Mnium stellare, a moss used in the Eastern Phoebe (Sayornis phoebe) nests. Photo by Hermann Schachner, through Creative Commons.

Figure 23. Funaria hygrometrica with immature capsules, a species used in nests of the Eastern Phoebe. Photo by Hermann Schachner, through Creative Commons.

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Figure 24. Polytrichum commune, member of a genus used in construction of nests of the Eastern Phoebe. Photo by Hermann Schachner, through Creative Commons.

Figure 25. Climacium dendroides, a moss used in nests of the Eastern Phoebe. Photo by Stan Phillips, through public domain.

Figure 26. Sayornis saya, Say's Phoebe. Members of this species often include mosses in their nests. Photo by Tom Grey, with permission.

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Chapter 16-6: Bird Nests – Passeriformes, part 1

Figure 27. Sayornis saya, Say's Phoebe, nest with young. Photo by Tom Grey, with permission. Figure 30. Tyrannus tyrannus, Eastern Kingbird, nest with eggs. Photo by Anc516, through Creative Commons.

Figure 28. Pitangus sulphuratus, Great Kiskadee. Members of this species often include mosses in their nests.. Photo by Tom Grey, with permission.

Figure 31. Tyrannus melancholicus, Tropical Kingbird. Members of this species often include mosses in their nests.. Photo by Tom Grey, with permission.

Eastern Kingbird (Tyrannus tyrannus) The Eastern Kingbird (Tyrannus tyrannus; Figure 29) of the Great Plains typically lives in forests where the canopy level is uneven, providing high points for observation and foraging (Johnsgard 2009). The female picks the nest site and builds the nest (Figure 30). She places it on outer branches of shrubs or small trees and often incorporates mosses in the construction.

Figure 29. Tyrannus tyrannus, Eastern Kingbird. Members of this species often include mosses in their nests. Photo by MDF, through Creative Commons.

Figure 32. Tyrannus couchii, Couch's Kingbird. Members of this species often include mosses in their nests. Photo by Ruben, through Creative Commons.

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Figure 33. Tyrannus forficatus, Scissor-tailed Flycatcher. Members of this species often include mosses in their nests.. Photo by Tom Grey, with permission.

Yellow-bellied Chat-tyrant (Ochthoeca diadema) Miller and Greeney (2008) described the nest of the Yellow-bellied Chat-tyrant (Ochthoeca diadema; Figure 34). They found a partially domed cup built into a vertical mat of mosses that hung from a horizontal vine. The cup was thick and composed of bryophytes with a sparse lining of feathers. The dome covered only about one-third of the cup. Closer examination revealed that the nest was actually build into the vertical sheet of mosses.

Figure 34. Ochthoeca diadema, Yellow-bellied Chat Tyrant. Members of this species sometimes build their nests into vertical hanging mats of mosses. Photo by Andres Cuervo, through Creative Commons.

Figure 35. Ochthoeca rufipectoralis, Rufous-breasted Chat Tyrant. Members of this species often include mosses in their nests. Photo by Dick Cook, through Creative Commons.

Figure 36. Ochthoeca cinnamomeiventris. Members of this species place mossy cups on ledges. Photo by Ken-ichi Ueda, through Creative Commons.

Crowned Chat-tyrant (Ochthoeca frontalis) Miller and Greeney (2008) found the Crowned Chattyrant (Ochthoeca frontalis) where it built its nest into a clump of mosses that was hanging 50 cm below a horizontal tree trunk (Miller & Greeney 2008). This provided good concealment by vegetation on the upper side. The nest was a partial dome made of mosses built into growing mosses and ferns. Other species, such as Rufous-breasted Chat (Ochthoeca rufipectoralis; Figure 35) and Slaty-backed Chat-tyrant (O. cinnamomeiventris; Figure 36) also place their mossy cups on ledges (Hilty & Brown 1986; Greeney 2007).

Figure 37. Pachyramphus aglaiae, Rose-throated Becard. Members of this species often include mosses in their nests. Photo by Dominic Sherony, through Creative Commons.

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Chapter 16-6: Bird Nests – Passeriformes, part 1

Laniidae – Shrikes Wolf (2009) found two species of Laniidae that use bryophytes in their nests in North America: Lanius ludovicianus (Loggerhead Shrike; Figure 38) Lanius excubitor (Northern Shrike; Figure 39)

Figure 40. Vireo griseus, White-eyed Vireo. Members of this species often include bryophytes in their nests. Photo by Andy Reago and Chrissy McClarren, through Creative Commons.

Figure 38. Lanius ludovicianus, Loggerhead Shrike. Members of this species often include mosses in their nests. Photo by Tom Grey, with permission.

Figure 41. Vireo cassinii, Cassin's Vireo. Members of this species often include mosses in their nests. Photo by Tom Grey, with permission.

Figure 39. Lanius excubitor, Northern Shrike. Members of this species often include mosses in their nests. Photo by Smudge 9000, with permission.

Vireonidae – Typical Vireos Wolf (2009) found three species of Corvidae that use bryophytes in their nests in North America: Vireo griseus (White-eyed Vireo; Figure 40) Vireo cassinii (Cassin's Vireo; Figure 41-Figure 42) Vireo huttoni (Hutton’s Vireo; Figure 43)

Figure 42. Vireo cassinii, Cassin's Vireo, nest with female. Members of this species often include mosses in their nests. Photo by Tom Grey, with permission.

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Monarchidae – Monarch Flycatchers The Rarotonga Flycatcher (Pomarea dimidiata; Figure 46), an endangered species in the Cook Islands of Polynesia, makes a nest entirely from mosses (Figure 46Figure 47), mostly Meteoriaceae (Figure 48) (John Game, Bryonet 22 June 2016).

Figure 43. Vireo huttoni, Hutton's Vireo. Members of this species often include mosses in their nests. Photo by Tom Grey, with permission.

Rhipiduridae – Fantails The Grey Fantail (Rhipidura albiscapa) in Tasmania builds a tidy nest of grass, moss sporophytes, bark, other plant fibers, ad spider webs (Lloyd 2013). The webs are used to attach the nest to a branch. The moss sporophytes are used to line the cup of the nest. These nests are built by the males and females in the understorey shrubs and small trees and both birds contribute to feeding.

Figure 46. Pomarea dimidiata, Rarotonga Flycatcher, at mossy nest. Photo by G. McCormack © CINHP , with online permission.

Figure 47. Pomarea dimidiata, Rarotonga Flycatcher, on nest of mosses. Photo by G. McCormack © CINHP , with online permission. Figure 44. Rhipidura albiscapa (Grey Fantail), a species that lines its nest with moss sporophytes. Photo by Patrick Kavanagh, through Creative Commons.

Figure 48. Weymouthia mollis, member of Meteoraceae that is common in bird nests. Photo by Clive Shirley, Hidden Forest , with permission. Figure 45. Rhipidura albiscapa (Grey Fantail) nest and nestlings. Photo by Benjamint444, through Creative Commons.

Myiagra cyanoleuca (Migratory Satin Flycatcher; Figure 49) builds a nest on a dead horizontal branch 5-25 m

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Chapter 16-6: Bird Nests – Passeriformes, part 1

above ground (Lloyd 2013). It uses bark strips and moss tightly bound with spider webs, making it neat and well disguised.

Figure 51. Cyanocitta stelleri, Steller's Jay. Members of this species often include mosses in their nests. Photo by Tom Grey, with permission.

Figure 49. Myiagra cyanoleuca (Satin Flycatcher) male, a species that includes mosses in its nests. Aviceda at English Wikipedia, though Creative Commons.

Corvidae – Jays, Magpies, & Crows Wolf (2009) found nine species of Corvidae that use bryophytes in their nests in North America: Perisoreus canadensis (Gray Jay; Figure 50) Cyanocitta stelleri (Steller’s Jay; Figure 51) Cyanocitta cristata (Blue Jay; Figure 52) Cyanocorax yncas (Green Jay; Figure 53) Aphelocoma californica (California Scrub-jay; Figure 54) Gymnorhinus cyanocephalus (Pinyon Jay; Figure 55) Nucifraga columbiana (Clark’s Nutcracker; Figure 56) Corvus brachyrhynchos (American Crow; Figure 57) Corvus caurinus (Northwestern Crow; Figure 58) Corvus corax (Common Raven; Figure 59)

Figure 50. Perisoreus canadensis, Gray Jay. Members of this species often include mosses in their nests. Photo by Walter Siegmund, through Creative Commons.

Figure 52. Cyanocitta cristata, Blue Jay. Members of this species often include mosses in their nests. Photo by Tom Grey, with permission.

Figure 53. Cyanocorax yncas, Green Jay. Members of this species often include mosses in their nests. Photo by Tom Grey, with permission.

Chapter 16-6: Bird Nests – Passeriformes, part 1

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Figure 54. Aphelocoma californica, California Scrub-jay. Members of this species often include mosses in their nests. Photo by Tom Grey, with permission. Figure 57. Corvus brachyrhynchos, American Crow. Members of this species often include mosses in their nests. Photo by Tom Grey, with permission.

Figure 55. Gymnorhinus cyanocephalus, Pinyon Jay. Members of this species often include mosses in their nests. Photo by James St. John, through Creative Commons.

Figure 56. Nucifraga columbiana, Clark's Nutcracker. Members of this species often include mosses in their nests. Photo by Tom Grey, with permission.

Figure 58. Corvus caurinus, Northwestern Crow. Members of this species often include mosses in their nests. Photo by T Greyfox, through Creative Commons.

Common Raven (Corvus corax) The Raven (Corvus corax; Figure 59) uses mosses to line its nest (Giannetta 2000).

Figure 59. Corvus corax, Raven. Members of this species often include mosses in their nests. Photo by Dick Daniels, through Creative Commons.

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Chapter 16-6: Bird Nests – Passeriformes, part 1

Hirundinidae – Swallows Wolf (2009) found only two species of Hirundinidae that use bryophytes in their nests in North America: Tachycineta bicolor (Tree Swallow; Figure 60-Figure 61) Stelgidopteryx serripennis (Northern Rough-winged Swallow; Figure 62)

Tree Swallow (Tachycineta bicolor) Tree Swallows (Tachycineta bicolor; Figure 60) are known to construct a basket nest (Figure 61) of sticks with an "upholstery" of moss, grass, and animal fur (Heinrich 2000). Heinrich assumed these to provide insulation and to cushion the eggs.

Figure 62. Stelgidopteryx serripennis, Northern Roughwinged Swallow. Members of this species often include mosses in their nests. Photo by Tom Grey, with permission.

Paridae – True Tits Wesołowski (unpublished data) found that the tits typically gathered moss for their nests in the immediate vicinity of the nest cavity, but they also would travel up to 80 m to gather nesting materials. Wolf (2009) found eight species of Paridae that use bryophytes in their nests in North America: Figure 60. Tachycineta bicolor, Tree Swallow, male. Members of this species use bryophytes in their treehole nests. Photo by Tom Grey, with permission.

Figure 61. Tachycineta bicolor, tree swallow, in a nest where bryophytes were used. Photo through public domain.

Poecile atricapillus (Black-capped Chickadee; Figure 74) Poecile gambeli (Mountain Chickadee; Figure 89) Poecile rufescens (Chestnut-backed Chickadee; Figure 90) Poecile hudsonicus (Boreal Chickadee; Figure 91) Poecile cinctus (Gray-headed Chickadee; Figure 92) Baeolophus inornatus (Oak Titmouse; Figure 93) Baeolophus ridgwayi (Juniper Titmouse; Figure 94) Baeolophus bicolor (Tufted Titmouse; Figure 95) Wesołowski and Wierzcholska (2018) compared the nesting materials used by three species of tit (Figure 63) and demonstrated that they were selective. Furthermore, the selections differed among the species. They avoided the abundant Brachythecium rutabulum (Figure 64), and Plagiothecium nemorale (Figure 65) and almost never used Anomodon longifolius (Figure 66) or Brachythecium oedipodium (Figure 67). Of the 54 available species, 21 were never used. Most plots associated with the nests had an average of 10.2-11.6 moss species/plot. The liverwort Metzgeria furcata (Figure 68) was used exclusively by Marsh Tits, and in greater proportion than in the environment. Brachythecium salebrosum was used only by Blue Tits, who also used large quantities of two forms of Hypnum cupressiforme (Figure 103). Great Tits underused Hypnum cupressiforme forms but used Anomodon viticulosus (Figure 69), and possibly also Pleurozium schreberi (Figure 70) in greater proportion than their

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availability. Wesołowski and Wierzcholska found no difference in water uptake between used and unused mosses. The Great Tits used mosses (Anomodon viticulosus, Isothecium alopecuroides (Figure 71), Pleurozium schreberi) with stems twice as thick as those used by the Marsh Tits [Hypnum cupressiforme mod. filiforme (Figure 72), Neckera complanata (Figure 73)].

Figure 65. Plagiothecium nemorale, an abundant moss that is avoided by tits as a nesting material. Photo by Michael Luth, with permission.

Figure 66. Anomodon longifolius, an abundant moss that is rarely used by tits for their nests. Photo by Hermann Schachner, through Creative Commons. Figure 63. Moss choice in nests of three species of tits in Poland. The percent represents to the percent of volume of mosses in the moss layer of nests that had mosses. Small squares represent the medians, boxes indicate 25-75% quartiles, and whiskers show the ranges. Numbers in parentheses are sample sizes. Modified from Wesołowski & Wierzcholska 2018.

Figure 64. Brachythecium rutabulum, an abundant moss avoided by tits. Photo by Janice Glime.

Figure 67. Brachythecium oedipodium, an abundant moss that is rarely used by tits for their nests. Photo by Michael Luth, with permission.

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Chapter 16-6: Bird Nests – Passeriformes, part 1

Figure 68. Metzgeria furcata, a liverwort that often occurs in tit nests, but in small quantity. Photo by Michael Luth, with permission.

Figure 69. Anomodon viticulosus, a preferred moss for nests by Great Tits. Photo by Janice Glime.

Figure 71. Isothecium alopecuroides with capsules, mosses with thick stems that preferred by Great Tits for nest materials. Photo by David T. Holyoak, with permission.

Figure 72. Hypnum cupressiforme mod. filiforme, a moss with thin stems and that is used for nest materials by Marsh Tits. Photo by Jan-Peter Frahm, with permission.

Figure 73. Neckera complanata, a moss with thin stems and that is used for nest materials by Marsh Tits. Photo by Michael Lüth, with permission. Figure 70. Pleurozium schreberi, a preferred moss for nests of Great Tits. Photo by Janice Glime.

But why did these birds travel as much as 80 m to gather some species when unused ones were much closer? When Wesołowski and Wierzcholska (2018) used human

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plucking of the mosses used in nests and compared them to plucking of the unused species, they found that the used species yielded larger (heavier) bundles of moss and contained longer shoots than of those mosses that were ignored by the birds. This suggests that there is an energy benefit when using the selected species. Black-capped Chickadee (Poecile atricapillus) Allen (2017) observed a Black-capped Chickadee (Poecile atricapillus; Figure 74) busily gathering dry moss for its nest, then flying to the nestbox. The stream had lots of moss, but the bird ignored these, preferring the dry patch next to the stream. The Robin, on the other hand, preferred the wet moss for its open, mud-lined nest.

Figure 76. Poecile carolinensis, Carolina Chickadee, with nesting materials. Photo by Tom Grey, with permission.

Figure 74. Poecile atricapillus, Black-capped Chickadee. Members of this species gather dry mosses near a stream for their nests. Photo by Tattooed Dreamer, through Creative Commons.

Carolina Chickadee (Poecile carolinensis) Erichsen (1919) describes the appearance of "down" on the cinnamon and royal ferns as a signal that the Carolina Chickadee (Poecile carolinensis; Figure 75) will begin its nest building (Figure 76). The Carolina Chickadee often begins this nest (Figure 77) by placing a thick mat of green moss (often Hypnum; Figure 78) from the tree trunks into the nesting cavity (Figure 77). This always occurs first, followed by the soft down of the ferns.

Figure 75. Poecile carolinensis, Carolina Chickadee. Members of this species often include mosses in their nests. Photo by Tom Grey, with permission.

Figure 77. Poecile carolinensis, Carolina Chickadee, nest. Photo courtesy of Diane Lucas.

Figure 78. Hypnum imponens, a common species in a genus used for nests of the Carolina Chickadee. Photo by Janice Glime.

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Andreas (2010) observed nests of two Carolina Chickadees (Poecile carolinensis; Figure 75). These included ten mosses and two liverworts. The dominant species were the pleurocarpous moss Platygyrium repens (Figure 79) and the leafy liverwort Frullania eboracensis (Figure 80) plus a few others, which comprised 55% of the nesting material by volume. In another year, the bryophytes comprised 70.4% of the nest material. The selection of bryophytes was not in proportion to their abundance and all species used were epiphytic on bark. Andreas suggested that they may select Frullania eboracensis for its chemical properties, possibly protecting them from mites (Figure 111). Only corticolous (growing on tree bark) bryophytes were used, with the exception of a single piece of Bryoandersonia illecebra (Figure 81) in one nest. But even clumps of acrocarpous (mostly upright with archegonia and capsules forming at tip of stem) mosses were removed from the tree trunks as tiny tufts for nest usage, including Orthotrichum ohioense and Dicranum montanum (Figure 82). Other corticolous bryophytes, including Anomodon attenuatus (Figure 83), Brachythecium laetum (Figure 84), Clasmatodon parvulus (Figure 85), Hypnum pallescens (Figure 86), and Ulota crispa (Figure 87), were ignored.

Figure 81. Bryoandersonia illecebra, the only non-epiphytic moss used in a Carolina Chickadee nest. Photo by Bob Klips, with permission.

Figure 82. Dicranum montanum, an acrocarpous moss used in the nest of a Carolina Chickadee. Photo by Hermann Schachner, through Creative Commons. Figure 79. Platygyrium repens with bulbils, a moss used in nests of Carolina Chickadees. Photo by Hermann Schachner, through Creative Commons.

Figure 80. Frullania eboracensis, a leafy liverwort used in nests of Carolina Chickadees. Photo from Dale A. Zimmerman Herbarium, Western New Mexico University, with permission.

Figure 83. Anomodon attenuatus with capsules, an epiphytic moss that was ignored when the Carolina Chickadee built its nest. Photo by Bob Klips, with permission.

Chapter 16-6: Bird Nests – Passeriformes, part 1

Figure 84. Brachythecium laetum, an epiphytic moss that was ignored when the Carolina Chickadee built its nest. Photo by Bob Klips, with permission.

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Figure 87. Ulota crispa, an epiphytic moss that was ignored when a Carolina Chickadee built its nest. Photo by Michael Lüth, with permission.

In Cashiers, NC, a Carolina Chickadee (Poecile carolinensis; Figure 75) used Thuidium delicatulum (Figure 88) in its nest in an English Boxwood shrub (Annie Martin, Bryonet 1 June 2010).

Figure 88. Thuidium delicatulum, a ground moss used in the nest of a Carolina Chickadee. Photo by Janice Glime. Figure 85. Clasmatodon parvulus, an epiphytic moss that was ignored when a Carolina Chickadee built its nest. Photo by A. Newman, through Creative Commons.

Figure 86. Hypnum pallescens, an epiphytic moss that was ignored when a Carolina Chickadee built its nest. Photo by Michael Lüth, with permission.

Figure 89. Poecile gambeli, Mountain Chickadee. Members of this species often include mosses in their nests. Photo by Tom Grey, with permission.

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Figure 93. Baeolophus inornatus, Oak Titmouse, with its nest in the large hole at the bottom left. Members of this species include bryophytes in their nests. Photo by Tom Grey, with permission. Figure 90. Poecile rufescens, Chestnut-backed Chickadee. Members of this species often include mosses in their nests. Photo by Tom Grey, with permission.

Figure 91. Poecile hudsonicus, Boreal Chickadee. Members of this species often include mosses in their nests. Photo by David Mitchell, through Creative Commons.

Figure 92. Poecile cinctus, Grey-headed Chickadee. Members of this species often include mosses in their nests. Photo by Jargal Lamjav, through Creative Commons.

Figure 94. Baeolophus ridgwayi, Juniper Titmouse. Members of this species often include mosses in their nests. Photo by Tom Grey, with permission.

Figure 95. Baeolophus bicolor, Tufted Titmouse. Members of this species often include mosses in their nests. Photo by Tom Grey, with permission.

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Varied Tit (Sittiparus varius) The Varied Tit (Sittiparus varius; Figure 96) lives in coniferous forests, mixed forests, and bamboo in eastern Japan, Korea, and some parts of northeastern China and extreme southeastern Russia (southern Kurile Islands). It is one of the birds that uses bryophytes for nesting material (Sakai 2007).

Figure 97. Parus major, Great Tit,. Members of this species often include bryophytes in their nests. Photo by Paul Gulliver, through Creative Commons.

Figure 96. Sittiparus varius, Varied Tit. Members of this species often include mosses in their nests. Photo by Alpsdake, through Creative Commons. Figure 98. Parus major, Great Tit, nest with bryophytes and eggs. Photo by Oh Wei, through Creative Commons.

Blue Tit (Cyanistes caeruleus), Great Tit (Parus major), and Japanese Tit (Parus minor) The Great Tit (Parus major; Figure 97-Figure 98) and the Blue Tit (Cyanistes caeruleus; Figure 99-Figure 101) both use mosses to build their nests (Figure 98) (Hribek 1985). Likewise, Gustavo Tomás and Andrew Spink (pers. comm. 2010) have collected mosses from a large number of Blue Tit (Cyanistes caeruleus) and Coal Tit (Periparus ater; Figure 102) nests in the Netherlands. The most common species in the nest is the locally common Hypnum cupressiforme (Figure 103). But other locally common species are not common in the nests, suggesting a preference. It appears that different species may be used in different parts of the nest, but so far there is no quantitative analysis available to support this. Figure 108 demonstrates the use of a Hypnum species (with Thuidium) in the nest of an unknown bird in Pennsylvania, USA.

Figure 99. Cyanistes caeruleus, Eurasian Blue Tit,. Members of this species build their nests with mosses. Photo by Francis C. Franklin, through Creative Commons.

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Chapter 16-6: Bird Nests – Passeriformes, part 1

Figure 100. Cyanistes caeruleus, Blue Tit, mossy nest and eggs. Photo by Notts Ex Miner, through Creative Commons.

Figure 103. Hypnum cupressiforme, a preferred moss in the nests of Blue Tits and Coal Tits. Photo by Michael Lüth, with permission.

Although the population may use a wide variety of mosses, a few species of bryophytes typically comprise the nest. For example, the Japanese Tit, Parus minor, used 21 species of bryophytes in the nests, but among 91% of the 47 nests, more than 50% of the volume was comprised of only three bryophyte species (Hamao et al. 2016). In this case, the preference seems to relate to a potential food source. The Japanese Tits preferred pleurocarpous mosses. In thse nests, seven species of moths emerged from the nesting material and were more frequent in nests with successful fledgine than in failed nests.

Figure 101. Cyanistes caeruleus, Blue Tit, nest with moss and nestlings. Photo by Notts Ex Miner, through Creative Commons.

Figure 104. Parus minor, Japanese Tit, a species that seems to be selctive in choice of mosses for its nests. Photo by Hyun-tae Kim, through Creative Commons.

Figure 102. Periparus ater, Coal Tit. Members of this species often include mosses in their nests, preferring Hypnum cupressiforme. Photo by Aviceda, through Creative Commons.

In the Czech Republic, Hříbek (1985) found that Blue Tits (Figure 99-Figure 101) used mostly softer species (Hypnum cupressiforme (Figure 103), Leptodictyum riparium (Figure 105), whereas the Great Tits used mostly the large-stemmed mosses such as Calliergonella cuspidata (Figure 106) and Rhytidiadelphus squarrosus (Figure 107).

Chapter 16-6: Bird Nests – Passeriformes, part 1

Figure 105. Leptodictyum riparium, a favorite nesting material of Blue Tits in the Czech Republic, with capsule. Photo by Michael Lüth, with permission.

Figure 106. Calliergonella cuspidata, one of the nesting materials of Great Tits in the Czech Republic. Photo by David T. Holyoak, with permission.

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The researchers set out to support this hypothesis with the Great Tit, a species that has a wide range of habitats, using populations in four different Mediterranean habitats. Interestingly, the clutch size decreased as moss mass increased in the four sites. However, hatching success increased as the moss mass increased in one site. And in all the habitats, the nestling condition was poorer in nests with a greater proportion of sticks and feathers. Mainwaring et al. (2012) reported that the nests of Blue Tits (Cyanistes caeruleus) and Great Tits (Parus major; Figure 97-Figure 98) in Great Britain consist of a "pad of moss mixed with dry grass and other plant material placed at the base of the nest box" (Figure 109) (Cramp & Perrins 1993; Mainwaring et al. 2008; Mainwaring & Hartley 2008, 2009; Britt & Deeming 2011). The nest cup is lined with fine dry grass, hair, wool and feathers. In this case, it appears that the mosses may be used to regulate the temperature and insulate the eggs and young birds. When temperatures increase, the female reduces the amount of lining material.

Figure 108. Hypnum and Thuidium in unidentified nest. Photo courtesy of Jeri Peck.

Figure 107. Rhytidiadelphus squarrosus, one of the nesting materials of Great Tits in the Czech Republic. Photo by Michael Lüth, with permission.

Álvarez et al. (2013) asserted that the properties and structure of a nest affect breeding performance. This drives the selection of behavior that produces nests characteristic of the species, including the appropriate nesting materials. Where preferred materials are low, birds select alternative materials, often at the cost of reduced breeding success.

Figure 109. Parus major, Great Tit, with eggs in nest on mosses. Photo by Notts Ex Miner, through Creative Commons.

When Great Tits (Parus major; Figure 97) built a second nest in nest boxes after rearing their first brood, they still used mosses in the nest, but there was no lining or inner layer – or any eggs (Slagsvold 1984).

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Chapter 16-6: Bird Nests – Passeriformes, part 1

The Corsican Blue Tit (Cyanistes caeruleus ogliastrae; Figure 110) includes 1-5 aromatic herbs in its nest (Lambrechts & Dos Santos 2000). Herbs are included in a number of kinds of bird nests, and researchers have suggested that they may serve an anti-parasite function (Figure 111) (Wimberger 1984; Bucher 1988; Cowie & Hinsley 1988; Clark 1991; Banbura et al. 1995). Using an herb removal experiment when the young hatched, these researchers found that the parents brought fresh aromatic greens to the nest. They proposed the Potpourri hypothesis that included at least seven functional causes for materials used in the nests. When the Blue Tits breed in cavities, they use predominately mosses, but also include other materials, including fresh herbaceous leaves. They suggested that mosses may optimize the microclimate in the nest cavity. The aromatic herbs are likely to serve an anti-parasitic function.

2004). But more recently it appears that it should be classified in the Paridae with the Chickadees. These birds are common in forests and woody suburbs of Europe and North America, but it appears that their ancestors lived on the dry, treeless Tibetan plateau. They nest in cavities where they build nests of grasses and mosses. Like Jays, they rarely fly, but they do not run like the Jays; rather, they hop.

Figure 112. Pseudopodoces humilis, Ground Tit. Members of this species build nests of grasses and mosses. Photo by David Blank, through Creative Commons.

Pipridae – Manakins, Piprites Black-capped Piprites (Piprites pileata) Only one example in this family has emerged. The Black-capped Piprites (Piprites pileata; Figure 113) builds a spherical nest made of mosses (Cocckle et al. 2008). Figure 110. Cyanistes caeruleus ogliastrae, Corsican Blue Tit. Members of this species often include mosses in their nests. Photo by Valter Jacinto, through Creative Commons.

Figure 111. Cyanistes caeruleus, Eurasian Blue Tit, with mite infestation causing balding. Photo by Michael Palmer, through Creative Commons.

Ground Tit (Pseudopodoces humilis) Ground Tit, also known as Hume's Jay, (Pseudopodoces humilis; Figure 112) has been considered the smallest member of the Jay and Crow family (Lipske

Figure 113. Piprites pileata, Black-capped Piprites. Members of this species often build their nests of mosses. Photo by Bruno Lima, through Creative Commons.

Aegithalidae – Long-tailed Tits Wolf (2009) found one species of Aegithalidae whose members use bryophytes in their nests (Figure 114) in North America: Psaltriparus minimus (Bushtit; Figure 115).

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Figure 114. Psaltriparus minimus, Bushtit, at mossy nest. Photo by Walter Siegmund, through Creative Commons. Figure 117. Aegithalos caudatus, Long-tailed Tit juvenile. Photo by Charles J. Sharp, through Creative Commons.

Figure 115. Psaltriparus minimus, Bushtit, pulling on nest materials. Photo by Mikul, through Creative Commons.

Long-Tailed Tit (Aegithalos caudatus) The Long-tailed Tit (Aegithalos caudatus; Figure 116Figure 117) has been separated from other tits and has different feeding and nesting (Figure 118) habits from them. These are not seed-eaters, eating mostly insects from bark crevices and buds. The families stay together, so that a flock will contain only related birds. Relatives that have lost their family members will join the flock. Nests may be tended by 1-8 adults. The female sits on the eggs and the male brings the food. Once the dozen or more babies hatch, helper adults gather food to feed them.

Figure 118. Aegithalos caudatus, Long-tailed Tit, building her nest in a hedgerow. Photo by Gail Hampshire, through Creative Commons.

The nests are bag-shaped and woven from mosses, bound with spider webs (Burton 1996). The birds cover the outside of the nest with lichens, sometimes substituting plastic and newspaper in areas of human habitation. This nest is insulated on the inside with feathers. The tits may accumulate ~1130 km of travel to gather nest materials. Hansell (2002) reported a nest with 5000-6000 pieces of material, including short-leaved mosses and cocoons intertangled, creating a Velcro effect with a few hundred sprigs of mosses. Sittidae – Nuthatches

Figure 116. Aegithalos caudatus, Long-tailed Tit, a species whose members build nests with mosses. Photo by drplokta, through Creative Commons.

Wolf (2009) found two species of Sittidae that use bryophytes in their nests in North America: Sitta carolinensis (White-Breasted Nuthatch; Figure 119) Sitta pygmaea (Pygmy Nuthatch; Figure 121)

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Certhiidae – Holarctic Treecreepers Wolf (2009) found one species of Certhidae whose members use bryophytes in their nests in North America: Certhia americana (Brown Creeper; Figure 122-Figure 123).

Figure 119. Sitta carolinensis, White-breasted Nuthatch. Members of this species often include bryophytes in their nests. Photo by Tom Grey, with permission.

Red-Breasted Nuthatch (Sitta canadensis) The Red-breasted Nuthatch (Sitta canadensis; Figure 120) builds its nest in tree holes, generally about 2.5 cm in diameter (Heinrich 2009; Moss Musings 2017). Inside the hole it lines the nest with mosses, down, and fibers. In fact, its nest can be recognized from those of woodpeckers because they never line their nests.

Figure 122. Certhia americana, Brown Creeper, with a beak full of dinner. Photo by Alan and Elaine Wilson, through Creative Commons.

Figure 120. Sitta canadensis, Red-breasted Nuthatch, outside the mossy nest in the treehole. Photo by Cephas, through Creative Commons.

Figure 123. Certhia americana, Brown Creeper, a bird that uses mosses to construct its nests. Photo by Badjoby, through Creative Commons.

Troglodytidae – Wrens Wolf (2009) found five species of Troglodytidae that use bryophytes in their nests in North America:

Figure 121. Sitta pygmaea, Pygmy Nuthatch, at tree hole. Members of this species often include mosses in their nests. Photo by Tom Grey, with permission

Salpinctes obsoletus (Rock Wren; Figure 124) Catherpes mexicanus (Canyon Wren; Figure 125) Thryothorus ludovicianus (Carolina Wren; Figure 126-Figure 127) Thryomanes bewickii (Bewick’s Wren; Figure 128) Troglodytes pacificus (Pacific Winter Wren; Figure 131-Figure 133)

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Figure 124. Salpinctes obsoletus, Rock Wren. Members of this species often include mosses in their nests. Photo by Tom Grey, with permission.

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Figure 126. Thryothorus ludovicianus, Carolina Wren. Members of this species often include mosses in their nests and nest linings. Photo by Ken Thomas, through public domain.

Figure 127. Thryothorus ludovicianus, Carolina Wren, nest with a considerable proportion of mosses, and nestlings. Photo by Marvin, through Creative Commons. Figure 125. Catherpes mexicanus, Canyon Wren. Members of this species often include mosses in their nests. Photo by Tom Grey, with permission.

Carolina Wren (Thryothorus ludovicianus) The tiny Carolina Wren (Thryothorus ludovicianus; Figure 126) is revered in places like Virginia because of its penchant for eating lots of insects (Harrison 2003). They nest mostly in nooks and crannies, so nest boxes are especially suitable for them. Their nests (Figure 127) often contain mosses, along with leaves, twigs, rootlets, weed stalks, and even cast-off snake skins. Both males and females are the nest builders, but it is she who lines the nest with feathers, hair, fine grass, and moss. These prolific breeders will typically lay a second set of eggs as soon as the young birds leave the nest and may even have a third set.

Figure 128. Thryomanes bewickii, Bewick's Wren. Members of this species often include mosses in their nests. Photo by Tom Grey, with permission.

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Pacific Wren (Troglodytes pacificus) and Winter Wren (T. hiemalis) The Winter Wren has been divided into two species, the Pacific Wren (Troglodytes pacificus; Figure 129) and the Winter Wren (Troglodytes hiemalis; Figure 130), the eastern species (Toews & Irwin 2008). Where their breeding ranges overlapped, the two species were distinguishable by their songs and lack of cross mating. This evidence was supported by DNA analysis.

Figure 130. Troglodytes hiemalis, Winter Wren. Members of this species often include mosses in their nests. Photo by Paul Stein, through Creative Commons.

Eurasian Wren (Troglodytes troglodytes) Figure 129. Troglodytes pacificus, Pacific Wren. Members of this species often include mosses in their nests. Photo by Tom Talbott, through Creative Commons.

The Pacific Wren (Troglodytes pacificus; Figure 129) breeds in the coniferous forests of the Pacific Northwest and constructs a nest almost entirely of mosses (Hejl et al. 2002). These wrens protect their nests with a dome and small side entrance (Heinrich 2009). The winter wren places green mosses and small evergreen twigs on the outside. Some birds place their nests in hanging mosses near the ground, but more commonly they place them on tip-up mounds formed by roots of fallen trees. The Pacific Wren builds a round nest of grass, moss, lichens, or leaves that it stuffs into a hole in a wall, crack in a rock, corner of a building, or tree trunk, but can also put it in bushes or overhanging boughs (Wikipedia 2010).

Nests of the Eurasian Wren (Troglodytes troglodytes; Figure 131) can make its nest almost entirely of bryophytes (Figure 132). The Japanese variety (Troglodytes troglodytes fumigatus) likewise uses mosses (Figure 133).

Eastern Winter Wren (Troglodytes hiemalis) Piers (1897) reported two Winter Wren (Troglodytes hiemalis; Figure 130) nests in Nova Scotia, Canada, built in moss that was constantly saturated by water trickling from the bank above. Piers suspected that the second nest was a later one built by the same pair as the first.

Figure 131. Troglodytes troglodytes, Eurasian Wren, a bryophyte nest builder. Photo by Dibyendu Ash, through Creative Commons.

Chapter 16-6: Bird Nests – Passeriformes, part 1

Figure 132. Troglodytes troglodytes, Eurasian Wren, feeding young in nest of mosses and other materials. Photo by Sonja Kübelbeck, through Creative Commons.

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Figure 134. Cinclus mexicanus, American Dipper, on mosses on the streambank. Photo by Stephen Shunk, through Creative Commons.

Figure 135. Cinclus mexicanus, American Dipper, gathering moss for its nest. Photo by Frank D. Lospalluto, through Creative Commons.

Figure 133. Troglodytes troglodytes fumigatus, Japanese Winter Wren, shown here gathering mosses for its nest. Photo by Alpsdake, through Creative Commons.

Cinclidae – Dippers Wolf (2009) found one species of Cinclidae whose members use bryophytes in their nests in North America: Cinclus mexicanus (American Dipper; Figure 134-Figure 135), also known as the Water Ouzel.

The American Dipper (Figure 134-Figure 135) is the only aquatic songbird in North America (Rosentreter 2014). It is a year-round resident, maintaining its streamside territorial defense year-round. It is known for its diving ability, down to nearly 7 m below the surface, and lives along unpolluted streams with riffles, cascades, and waterfalls. It makes a ball-shaped nest with a side entrance, placed on a cliff face, in a crevice, or under a bridge abutment, positions that help it to avoid predators. The outer shell of this nest is moss with its inner chamber made of pine needles. It uses stream mosses that it dives to obtain, hence they are dripping wet. These are woven into the nest, still wet, and as they dry they tighten the weave and help to affix the nest to its vertical substrate. I have seen the nest of an American Dipper (Figure 134-Figure 135) in Colorado with the busy expectant mother diving into the water to collect Platyhypnidium

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riparioides (Figure 136) for the construction. The nest (Figure 137), wedged under the cliff behind a waterfall, appeared to be made entirely of mosses. Dan Norris (Bryonet 22 November 1995) reports that this bird is indeed selective, using mosses with a different frequency from that found in their habitat.

Figure 136. Platyhypnidium riparioides, a common moss used in nests of the American Dipper (Cinclus mexicanus). Photo by Stan Phillips, through public domain. Figure 138. Scouleria marginata, a common component of the American Dipper nests. Photo by Martin Hutten, with permission.

Figure 137. Cinclus mexicanus, American Dipper, nest of Hygrohypnum and Hygroamblystegium. Photo by Janice Glime.

Terry McIntosh (Bryonet 2 June 2010) identified mosses in Dipper (Cinclus mexicanus; Figure 134-Figure 135) nests from northern Idaho. To his surprise, he found only one species, Scouleria marginata (Figure 138), a somewhat rare moss, despite the much greater abundance of S. aquatica (Figure 139). He attributed this selection to the stronger plants of S. marginata. By contrast, Ellen Anderson (Bryonet 2 June 1010) found 30 moss species and 5 liverwort species (plus a few unknowns) in 7 dipper nests in the area around Juneau, Alaska, USA. Most of the nests had only traces of mosses, but nevertheless had quite a few species, numbering 1, 7, 10, 11, 13, 14, and 16 (plus 5 unknowns).

Figure 139. Scouleria aquatica, a common moss that is ignored as nesting material for the American Dipper when S. marginata is present. Photo by Matt Goff, with permission.

Roger Rosentreter (pers. comm. 20 January 2014) observed numerous American Dipper (Cinclus mexicanus; Figure 134-Figure 135) nests on the Payette River, Idaho, USA, reaching up to 2 nests per kilometer. In this case, the nests were composed primarily of the aquatic moss Scouleria aquatica (Figure 139), an abundant moss in the river.

Chapter 16-6: Bird Nests – Passeriformes, part 1

Brown Dipper (Cinclus pallasii) The Brown Dipper, also known as the Pallas Dipper, (Cinclus pallasii; Figure 140), is an Asian dipper that uses mosses in its nests (Nishimura et al. 1980).

Figure 140. Cinclus pallasii pallasii, Brown Dipper, a bird that uses aquatic bryophytes in its nests Photo by Alpsdake, through Creative Commons.

Summary The Passeriformes is the largest order of birds and contains the majority of birds that use bryophytes in their nests. Nevertheless, they seem to be a small proportion of the total species in the order. In this first part, the members using bryophytes include Tyrant Flycatchers, shrikes, Vireos, Jays and Crows, Swallows, Tits, Piprites, Nuthatches, and Wrens. Among these, the American Dipper is an aquatic bird that often dives for mosses to build its nest. Their selective choices may be energy savings by being able to gather larger bryophyte materials, providing nest-inhabiting food organisms, and in some cases possibly providing more constant moisture.

Acknowledgments Thank you to Brian Dykstra for sending me the wonderful thesis on birds and epiphytes by Adrian Wolf, as well as other references and personal observations. David Dumond shared the references he got from Bryonet. Many photographers have provided permission or put their images in Creative Commons, for which I am deeply appreciative. Thank you to Tom Grey and Janet Marr for a critical reading of the manuscript. And thank you to

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Bernard Goffinet for his continued support in sending me images and interesting publications.

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Glime, J. M. 2017. Bird Nests – Passeriformes, part 2. Chapt. 16-7. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. eBook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 19 July 2020 and available at .

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CHAPTER 16-7 BIRD NESTS – PASSERIFORMES, PART 2 TABLE OF CONTENTS Passeriformes (cont.) Grallariidae ................................................................................................................................................ 16-7-2 Regulidae – Kinglets .................................................................................................................................. 16-7-2 Sylviidae – Old-World Warblers and Gnatcatchers ................................................................................... 16-7-3 Turdidae – Thrushes................................................................................................................................... 16-7-3 Muscicapidae – Old World Flycatchers ..................................................................................................... 16-7-9 Petroicidae – Australian Robins ............................................................................................................... 16-7-10 Sturnidae – Starlings etc .......................................................................................................................... 16-7-10 Motacillidae – Wagtails and Pipits .......................................................................................................... 16-7-11 Bombycillidae – Waxwings ..................................................................................................................... 16-7-14 Peucedramidae – Olive Warbler .............................................................................................................. 16-7-14 Parulidae – Wood Warblers etc................................................................................................................ 16-7-15 Furnariidae – Neotropical Ovenbirds ....................................................................................................... 16-7-21 Thraupidae – Tanagers and Honeycreepers ............................................................................................. 16-7-24 Emberizidae – Emberizines...................................................................................................................... 16-7-24 Icteridae – Blackbirds, Orioles, etc. ......................................................................................................... 16-7-30 Fringillidae – Fringilline Finches ............................................................................................................. 16-7-31 Leiothrichidae – Laughing Thrushes........................................................................................................ 16-7-35 Ptilonorhynchidae – Bower Birds ............................................................................................................ 16-7-35 Acanthizidae – Scrubwrens, Thornbills, and Gerygones ........................................................................ 16-7-36 Rhinocryptidae – Tapaculos..................................................................................................................... 16-7-37 Callaeatidae – New Zealand Wattlebirds ................................................................................................. 16-7-37 Zosteropidae – White-eyes....................................................................................................................... 16-7-38 Effect of Cavity-nesting Birds on Bryophyte Communities............................................................................ 16-7-38 Edible Nests .................................................................................................................................................... 16-7-41 Summary ......................................................................................................................................................... 16-7-41 Acknowledgments ........................................................................................................................................... 16-7-41 Literature Cited ............................................................................................................................................... 16-7-41

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Chapter 16-7: Bird Nests – Passeriformes, part 2

CHAPTER 16-7 BIRD NESTS – PASSERIFORMES, PART 2

Figure 1. Grallaricula peruviana is a rare bird, shown here with bryophytes in its nest. Photo by Harold Greeney, through Creative Commons

Grallariidae The Peruvian Antpitta (Grallaricula peruviana) is a rare species that uses bryophytes in its nest, as seen in Figure 1. Regulidae – Kinglets Wolf (2009) found two species of Regulidae that use bryophytes in their nests in North America: Regulus satrapa (Golden-Crowned Kinglet; Figure 2) Regulus calendula (Ruby-Crowned Kinglet; Figure 4) The Golden-crowned Kinglet (Regulus satrapa; Figure 2) breeds in the coniferous forests (Figure 3) of the Pacific Northwest and constructs a nest almost entirely of mosses (Ingold & Galati 1997).

Figure 2. Regulus satrapa, Golden-crowned Kinglet. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

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Turdidae – Thrushes Wolf (2009) found thirteen species of Turdidae that use bryophytes in their nests in North America: Luscinia svecica (Bluethroat; Figure 6) Oenanthe oenanthe (Northern Wheatear; Figure 7) Sialia mexicana (Western Bluebird; Figure 8) Myadestes townsendi (Townsend’s Solitaire; Figure 9) Catharus fuscescens (Veery; Figure 11) Catharus minimus (Gray-Cheeked Thrush; Figure 12) Catharus bicknelli (Bicknell’s Thrush; Figure 13) Catharus ustulatus (Swainson’s Thrush; Figure 14) Catharus guttatus (Hermit Thrush; Figure 15-Figure 16) Turdus pilaris (Fieldfare; Figure 18-Figure 19) Turdus iliacus (Redwing; Figure 20) Turdus migratorius (American Robin; Figure 21-Figure 22) Ixoreus naevius (Varied Thrush; Figure 38) Figure 3. Conifer forest, Garibaldi National Park, BC, home to the Golden-crowned Kinglet, Regulus satrapa. Photo by The Simkin, through public domain.

Figure 4. Regulus calendula, Ruby-crowned Kinglet. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Sylviidae – Old-World Warblers & Gnatcatchers Wolf (2009) found one species of Sylviidae that use bryophytes in their nests in North America: Phylloscopus borealis (Arctic Warbler; Figure 5).

Figure 5. Phylloscopus borealis, Arctic Warbler. Members of this species use bryophytes in their nests. Photo by Osado, through Creative Commons.

Figure 6. Luscinia svecica, Bluethroat. Members of this species use bryophytes in their nests. Photo by Andreas Trepte, through Creative Commons.

Figure 7. Oenanthe oenanthe, Northern Wheatear. Members of this species use bryophytes in their nests. Photo by Craig Nash, through Creative Commons.

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Chapter 16-7: Bird Nests – Passeriformes, part 2

Figure 8. Sialia mexicana, Western Bluebirds. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 9. Myadestes townsendi, Townsend's Solitaire. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 10. Myadestes palmeri, Puaiohi, nest in a mossy cavity. Photo by Lucas Behnke, with permission.

Figure 11. Catharus fuscescens, Veery. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 12. Catharus minimus, Gray-cheeked Thrush. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 13. Catharus bicknelli, Bicknell's Thrush, on mossy nest. Photo by Kent McFarland, through Creative Commons.

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grasses. It is not lined with mosses, but rather with conifer needles, rootlets, and plant fibers.

Figure 14. Catharus ustulatus, Swainson's Thrush. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission. Figure 17. Bird nest in Coast Range of the Pacific Northwest, USA, with mosses still growing. Photo by JeriLynn Peck.

Figure 15. Catharus guttatus, Hermit Thrush. Members of this species use bryophytes in their nests. Photo by Cephas, through Creative Commons.

Figure 18. Turdus pilaris, Fieldfare. Members of this species use bryophytes in their nests. Photo by Allan Drewitt, through Creative Commons.

Figure 16. Catharus guttatus, Hermit Thrush nest and hatchlings. Photo by Per ver Donk, with permission.

Hermit Thrush (Catharus guttatus) Once again, the female is the sole nest-builder in the Hermit Thrush (Catharus guttatus; Figure 15-Figure 16) (Cornell Lab of Ornithology). Her bulky handiwork includes mosses in addition to twigs, bark strips, ferns, and

Figure 19. Turdus pilaris, Fieldfare, nest, showing occasional mosses mixed with grasses in the nest. Photo by Andreas Trepte, through Creative Commons.

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Chapter 16-7: Bird Nests – Passeriformes, part 2

Figure 20. Turdus iliacus, Redwing. Members of this species use bryophytes in their nests. Photo by Ómar Runólfsson, through Creative Commons.

American Robin (Turdus migratorius) The American Robin (Turdus migratorius; Figure 21) uses mosses as a binding material with mud in the inner cup of the nest (Figure 22-Figure 23) (Breil & Moyle 1976). It also uses mosses to line the cup. It seems to have a preference for Thuidium delicatulum (Figure 24), Plagiomnium cuspidatum (Figure 25), Brachythecium acuminatum (Figure 26), B. salebrosum (Figure 27), and Amblystegium varium (Figure 28).

Figure 23. Turdus migratorius, American Robin, nest and young. Photo by Tom Grey, with permission.

Figure 24. Thuidium delicatulum, a moss used as a mud binder to line the Robin's nest. Photo by Janice Glime.

Figure 21. Turdus migratorius, American Robin. Members of this species sometimes use mosses as a binder for the mud linings of their nests. Photo by Tom Grey, with permission.

Figure 22. Turdus migratorius, American Robin, on nest. Photo by Jane and Phil, through Creative Commons.

Figure 25. Plagiomnium cuspidatum, a moss used as a mud binder to line the Robin's nest. Photo by Hermann Schachner, through Creative Commons.

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Other members of the genus, such as the Yellowlegged Thrush (Turdus flavipes; Figure 29-Figure 30), place bryophytes on the outside of the nest.

Figure 26. Brachythecium acuminatum, a moss used as a mud binder to line the Robin's nest. Photo by Charles T. Bryson, through Creative Commons.

Figure 29. Turdus flavipes, Yellow-legged Thrush. Members of this species use mosses on the outsides of their nests. Photo by David R. Santiago, through Creative Commons.

Figure 27. Brachythecium salebrosum with capsules, a moss used as a mud binder to line the Robin's nest. Photo by Michael Lüth, with permission.

Figure 30. Turdus flavipes, Yellow-legged Thrush, nest with eggs and bryophytes. Photo by David R. Santiago, through Creative Commons.

Chinese Thrush (Turdus mupinensis)

Figure 28. Amblystegium varium, a moss used as a mud binder to line the Robin's nest. Photo by J. C. Schou, through Creative Commons.

In a Chinese study (Zhao et al. 2005), nests of the Chinese Thrush (Turdus mupinensis; Figure 31) were collected from Xiaolongmen Nature Reserve of Beijing. Nests exhibited seven bryophyte species: Anomodon sp., A. minor (Figure 32), Entodon sp. (Figure 33), Lindbergia sinensis (see Figure 34), Brachythecium sp. (see Figure 27), Herpetineuron sp. (Figure 35), Plagiomnium sp. (see Figure 25), and Myuroclada maximowiczii (Figure 36). Anomodon minor was one of the major nest components.

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Chapter 16-7: Bird Nests – Passeriformes, part 2

Figure 34. Lindbergia koelzii with capsules, member of a genus used in nests of the Chinese Thrush, Turdus mupinensis. Photo by Michael Lüth, with permission.

Figure 31. Turdus mupinensis, Chinese Thrush. Members of this species use mosses in their nests in China. Photo by Charles Lam, through Creative Commons.

Figure 35. Herpetineuron toccoae, member of a genus used in nests of the Chinese Thrush, Turdus mupinensis. Photo by Li Zhang, with permission.

Figure 32. Anomodon minor, a species that is used in nests of the Chinese Thrush. Photo by Michael Lüth, with permission.

Figure 36. Myuroclada maximoviczii, a species that is used in nests of the Chinese Thrush. Photo by Janice Glime

Blackbird (Turdus merula)

Figure 33. Entodon concinnus, in a genus that is used in nests of the Chinese Thrush. Photo by Hermann Schachner, through Creative Commons.

The Common Blackbird (Turdus merula; Figure 37) makes a bulky cup in its nest, using dry grasses, twigs, stalks, and yes, mosses (Snow 1958). These are plastered with mud or muddy leaves and lined with fine grass, thin dead stems, or rootlets. Mainwaring et al. (2014) found that as spring temperatures increased in the lower latitudes, the quantity of mosses used in the nests decreased, suggesting that mosses may be needed for insulation at cooler temperatures (Mainwaring et al. 2012).

Chapter 16-7: Bird Nests – Passeriformes, part 2

Figure 37. Turdus merula, Common Blackbird, nesting. Members of this species use bryophytes in their nests. Photo by J. J. Harrison, through Creative Commons.

Nest size of birds is limited on the upper end by becoming more conspicuous and requiring more energy to prepare (Møller 1990). On the small end, it loses insulating ability, stability, and protection to prevent nestlings from falling out of the nest. Møller manipulated nest size of the Blackbird (Turdus merula; Figure 37), a species that makes an open-cup woodland nest. When nests were exchanged for smaller or larger nests, there was no effect on nest egg predation by the exchange itself, but larger nests experienced more predation. But real nests that experienced predation were not significantly larger than successful nests. Møller suggested that nest size in nature is dependent on nest site.

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Figure 39. Ficedula narcissina, a Chinese species. Members of this species use bryophytes in their nests. Photo by Alpsdake, through Creative Commons.

Figure 40. Cyanoptila cyanomelana, Blue-and-white Flycatcher male, a species that uses bryophytes to make nests. Photo by Alpsdake, through Creative Commons.

Figure 38. Ixoreus naevius, Varied Thrush. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Muscicapidae – Old World Flycatchers In the same Chinese study (Zhao et al. 2005), nests of three members of this family [Narcissus Flycatcher (Ficedula narcissina; Figure 39), Blue-and-white Flycatcher (Cyanoptila cyanomelana; Figure 40-Figure 41), Daurian Redstart (Phoenicurus auroreus; Figure 42)] were collected from Xiaolongmen Nature Reserve of Beijing. These nests, like those of the Chinese Thrush, exhibited the same seven bryophyte species, with the moss Anomodon minor (Figure 32) as the main component of nests of all three bird species.

Figure 41. Cyanoptila cyanomelana, Blue-and-white Flycatcher male. Members of this species make their nests with bryophytes. Photo by Alpsdake, through Creative Commons.

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Chapter 16-7: Bird Nests – Passeriformes, part 2

intertwined moss branches. The nest was attached to a branch by numerous strands that were wrapped around the main branch and a smaller branch.

Figure 42. Phoenicurus auroreus, Daurian Redstart male. Members of this species use bryophytes in their nests. Photo by Alpsdake, through Creative Commons.

Petroicidae – Australian Robins Australian Pink Robin (Petroica rodinogaster) The Australian Pink Robin (Petroica rodinogaster; Figure 43) includes both lichens and mosses in its nest (Figure 44) (Newman & Bratt 1976). Figure 44. The tiny Australian Pink Robin’s nest woven from Thuidiopsis sparsa (Figure 45), with Emma´s index finger for 'scale.' Photo courtesy of Emma Pharo and David Meagher.

Figure 43. Petroica rodinogaster, Australian Pink Robin. Members of this species build their nests of mosses, especially Thuidiopsis sparsa. Photo by J. J. Harrison, through Creative Commons.

Pharo and Meagher (2011) reported finding a Pink Robin's nest that was made almost entirely from mosses. It was located in a mountain ash forest in Victoria, Australia, in an area that had been lightly burned two years earlier. The nest was "extraordinarily tiny on a branch of Olearia agrophylla." The nest was woven exclusively from Thuidiopsis sparsa (Figure 45) except for a few strands of grass. It is interesting that the moss was not even growing at the site. Therefore, the birds deliberately hunted that moss. The nest has a loose weave, but was strong, with

Figure 45. Thuidiopsis sparsa, a moss used to make the nest of the Australian Pink Robin (Petroica rodinogaster). Photo through Creative Commons.

Sturnidae – Starlings, etc. Wolf (2009) found one species of Sturnidae whose members use bryophytes in their nests in North America: European Starling (Sturnus vulgaris; Figure 46-Figure 47).

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Figure 48. Daucus carota leaves, a species included in nests of the European Starling, presumably to reduce parasite infections. Photo by BioImages, through Creative Commons. Figure 46. Sturnus vulgaris, European Starling, the only member of this family that uses mosses in its nest in North America. Photo by Ingrid Taylar, through Creative Commons.

Figure 47. Sturnus vulgaris, European Starling, at nest. Photo by Gynti 46, through Creative Commons.

The European Starling "prefers" to use the wild carrot Daucus carota (Figure 48) or the fleabane Erigeron philadelphicus (Figure 49) in its nest, both of which have known abilities to suppress parasitic mites in nests (Clark & Mason 1985). We can only wonder if the bryophytes might serve a protective role against mites and other parasites in forested sites.

Figure 49. Erigeron philadelphicus, a species included in nests of the European Starling, presumably to reduce parasite infections. Photo by Fritzflohr Reynolds, through Creative Commons.

Motacillidae – Wagtails & Pipits Wolf (2009) found one species of Motacillidae whose members use bryophytes in their nests in North America: Motacilla alba (White Wagtail; Figure 50-Figure 51) Anthus cervinus (Red-throated Pipit; Figure 54) Anthus rubescens (American Pipit; Figure 55)

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Chapter 16-7: Bird Nests – Passeriformes, part 2

White Wagtail (Motacilla alba) Des Callaghan (Bryonet 23 June 2016) reported that while in the wonderful north of Finland one summer, a fine place for Splachnaceae, he noticed an intriguing association between Splachnum vasculosum (Figure 52Figure 53) and the insectivorous passerine bird Motacilla alba (Figure 50). Could the Wagtails be attracted by the odor? Are the mosses a food source? Or do the S. vasculosum and Motacilla alba simply like the same habitat? Callaghan recorded this interesting habitat . Mosses are included in nests (Figure 51) of this wagtail species (Bouglouan 2016).

Figure 53. Splachnum vasculosum with capsules and males. Photo by Dick Haaksma, with permission.

Figure 50. Motacilla alba alba, White Wagtail. Members of this species use bryophytes in their nests. Photo by Luis Garcia, through Creative Commons.

Figure 54. Anthus cervinus, Red-throated Pipit. Members of this species use bryophytes in their nests. Photo by Tom Grey with permission.

Figure 51. Motacilla alba, White Wagtail, nest with eggs, a nest that often includes bryophytes. Photo by Walcoford, through Creative Commons.

Figure 52. Splachnum vasculosum colony, a preferred perch for White Wagtail (Motacilla alba). Photo by Des Callaghan, with permission.

Figure 55. Anthus rubescens, American Pipit, with insect. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

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Small Kauai Thrush (Myadestes palmeri) The Small Kauai Thrush or Puaiohi (Myadestes palmeri; Figure 56), a small Hawaiian endemic, builds a cavity nest (Figure 57) along a stream bank comprised mostly of bryophytes and tiny ferns, with a weave of fine grass (Kepler & Kepler 1983). The bryophytes trail out of the cavity mouth from the base of the nest, providing an opportunity for these bryophytes to attach and grow on the stream bank. Included bryophytes were the mosses Dicranum speirophyllum (Figure 58) and Campylopus sp. (Figure 59) and the liverworts Bazzania sp. (Figure 60) and Lepidozia sp. (Figure 61). Figure 58. Dicranum speirophyllum, a moss used in the Puaiohi (Myadestes palmeri) nest. Photo by John Game, through Creative Commons.

Figure 59. Campylopus umbellatus, a moss representing a genus used in the Puaiohi (Myadestes palmeri) nest. Photo by Michael Lüth, with permission. Figure 56. Myadestes palmeri, Small Kauai Thrush. Members of this species use bryophytes in their nests. Photo by Eike Wulfmeyer, through Creative Commons.

Figure 57. Myadestes palmeri, Puaiohi, nest with mosses in a cavity. Photo by Lucas Behnke, with permission.

Figure 60. Bazzania sp., a leafy liverwort representing a genus used in the Puaiohi (Myadestes palmeri) nest. Photo by Ondřej Zicha, through Creative Commons.

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Chapter 16-7: Bird Nests – Passeriformes, part 2

Figure 63. Bombycilla cedrorum, Cedar Waxwing. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission. Figure 61. Lepidozia sp., a leafy liverwort representing a genus used in the Puaiohi (Myadestes palmeri) nest. Photo by Ken-ichi Uedo, through Creative Commons.

Bombycillidae – Waxwings Wolf (2009) found two species of Bombycillidae that use bryophytes in their nests in North America: Bombycilla garrulus (Bohemian Waxwing; Figure 62) Bombycilla cedrorum (Cedar Waxwing; Figure 63-Figure 64)

Figure 64. Bombycilla cedrorum, Cedar Waxwing, nest with moss & eggs. Photo by Rich Mooney, through Creative Commons.

Peucedramidae – Olive Warbler Wolf (2009) found one species of Peucedramidae that uses bryophytes in their nests in North America: Peucedramus taeniatus (Olive Warbler; Figure 65).

Figure 62. Bombycilla garrulus, Bohemian Wax Wing. Members of this species use bryophytes in their nests. Photo by Randen Pederson, through Creative Commons.

Cedar Waxwing (Bombycilla cedrorum) The Cedar Waxwing (Bombycilla cedrorum; Figure 63) nests in edge habitat, using small evergreens and deciduous trees to hold its nests (Figure 64) (Heinrich 2009). The nest is somewhat similar to that of a Robin in size and rough appearance, but it has no mud lining. The outside typically is decorated with lichens and mosses, probably providing camouflage.

Figure 65. Peucedramus taeniatus, Olive Warbler. Members of this species use bryophytes in their nests. Photo by Ron Knight, through Creative Commons.

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Parulidae – Wood Warblers, etc. Wolf (2009) found 27 species of Parulidae that use bryophytes in their nests in North America: Oreothlypis ruficapilla (Nashville Warbler; Figure 67) Oreothlypis celata (Orange-crowned Warbler; Figure 66, Figure 68) Oreothlypis virginiae (Virginia's Warbler; Figure 69) Dendroica coronata (Yellow-rumped Warbler; Figure 70) Setophaga pitiayumi (Tropical Parula; Figure 71) Setophaga magnolia (Magnolia Warbler; Figure 72) Setophaga tigrina (Cape May Warbler; Figure 73) Setophaga caerulescens (Black-throated Blue Warbler; Figure 74-Figure 75) Setophaga nigrescens (Black-throated Gray Warbler; Figure 76) Setophaga virens (Black-throated Green Warbler; Figure 77) Setophaga townsendi (Townsend’s Warbler; Figure 78) Setophaga occidentalis (Hermit Warbler; Figure 79) Setophaga kirtlandii (Kirtland’s Warbler; Figure 80) Setophaga striata (Blackpoll Warbler; Figure 81) Setophaga cerulea (Cerulean Warbler; Figure 82) Setophaga ruticilla (American Redstart; Figure 83) Setophaga citrina (Hooded Warbler; Figure 84-Figure 85) Protonotaria citrea (Prothonotary Warbler; Figure 86) Helmitheros vermivorum (Worm-eating Warbler; Figure 88) Limnothlypis swainsonii (Swainson’s Warbler; Figure 90) Seiurus aurocapilla (Ovenbird; Figure 91-Figure 92) Parkesia noveboracensis (Northern Waterthrush; Figure 97) Parkesia motacilla (Louisiana Waterthrush; Figure 98) Oporornis agilis (Connecticut Warbler; Figure 99) Geothlypis trichas (Common Yellowthroat; Figure 100) Cardellina pusilla (Wilson’s Warbler; Figure 101) Cardellina canadensis (Canada Warbler; Figure 102)

Figure 68. Oreothlypis celata, Orange-crowned Warbler. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 69. Oreothlypis virginiae, Virginia's Warbler. Members of this species use bryophytes in their nests. Photo by Jerry Oldenettel, through Creative Commons. Figure 66. Oreothlypis celata, Orange-crowned Warbler. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 67. Oreothlypis ruficapilla, Nashville Warbler. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 70. Dendroica coronata, Yellow-rumped Warbler. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

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Chapter 16-7: Bird Nests – Passeriformes, part 2

Figure 71. Setophaga pitiayumi, Tropical Parula. Members of this species use bryophytes in their nests. Photo by Dario Sanchez, through Creative Commons.

Figure 74. Setophaga caerulescens, Black-throated Blue Warbler. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 72. Setophaga magnolia, Magnolia Warbler. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 75. Setophaga caerulescens, Black-Throated Blue Warbler, feeding young in nest. Members of this species use bryophytes in their nests. Photo by USFWS, through public domain.

Figure 73. Setophaga tigrina, Cape May Warbler. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 76. Setophaga nigrescens, Black-throated Gray Warbler. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

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Kirtland's Warbler (Setophaga kirtlandii) In Michigan the Kirtland's Warbler (Setophaga kirtlandii; Figure 80) harvests moss sporophytes (Brian Dykstra, pers. comm. 10 December 2011).

Figure 77. Setophaga virens, Black-throated Green Warbler. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Townsend's Warbler (Setophaga townsendi) Some birds have very specific uses for the bryophytes. The Townsend's Warbler (Setophaga townsendi; Figure 78) lines its nest with the setae (stalks of moss capsules) of mosses (and hair) (Baicich & Harrison 2005). Figure 80. Setophaga kirtlandii, Kirtland's Warbler, in Jack pine. Members of this species harvest moss sporophytes, presumably for their nests. Photo by Ron Austing, through Creative Commons.

Figure 78. Setophaga townsendi, Townsend's Warbler. Members of this species use bryophytes in their nests. Photo by Jerry Oldenettel, through Creative Commons. Figure 81. Setophaga striata, Blackpoll Warbler. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 79. Setophaga occidentalis, Hermit Warbler. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 82. Setophaga cerulea, Cerulean Warbler. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

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Chapter 16-7: Bird Nests – Passeriformes, part 2

Prothonotary Warbler (Protonotaria citrea)

Figure 83. Setophaga ruticilla, American Redstart. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 84. Setophaga citrina, Hooded Warbler. Members of this species use bryophytes in their nests. Photo by Mary Elliott, through Creative Commons.

The Prothonotary Warbler (Protonotaria citrea; Figure 86) nests in abandoned holes made by woodpeckers. Although it sometimes uses few mosses in the actual nest, it does build it on a bed of bryophytes, both mosses and liverworts (Bent 1953; Petit 1989; Blem & Blem 1992, 1994). When building in a nest box, the mosses go in first to form the bed. Then the nest is built on top of them. The bryophytes remain moist, but the cup is not. Blem and Blem found that 75-80% of the dry mass of the nests they studied is composed of mosses and liverworts. They identified five species of mosses and two liverworts (Table 1), with the moss Anomodon attenuatus (Figure 87) predominating. They suggested that the bryophytes maintain the needed environment within the nest cavity (e.g. Mertens 1977a, b). In addition to ameliorating the moisture, bryophytes may serve to reduce pathogens and parasites (Clark & Mason 1985). I have seen several pictures of these nests, but unfortunately I could not find the name of the photographer on those sites.

Figure 86. Protonotaria citrea, Prothonotary Warbler, a species that uses a bed of bryophytes under its nest. Photo by David Inman, through Creative Commons.

Table 1. Occurrence of bryophytes in Prothonotary Warbler (Protonotaria citrea) nests in nest boxes along the James River, Virginia, USA. From Blem & Blem 1994.

Figure 85. Setophaga citrina, Hooded Warbler. Members of this species use bryophytes in their nests. Photo by USFSW, through public domain.

Species Mosses Anomodon attenuatus Haplocladium microphyllum Amblystegium varium Plagiomnium cuspidatum Thuidium delicatulum Liverworts Porella platyphylla Frullania eboracensis

Percent occurrence Mid- BotTop dle tom Total 97.3 96.4 91.4 95.0 20.6 13.4 21.0 18.3 6.7 7.6 1.3 5.2 2.7 1.3 3.1 2.4 0.4 1.3 0.0 0.6 21.9 27.3 32.1 27.1 0.4 0.8 0.8 0.7

Chapter 16-7: Bird Nests – Passeriformes, part 2

Figure 87. Anomodon attenuatus with capsules, the primary bryophyte used in the nest of the Prothonotary Warbler. Photo by Bob Klips, with permission.

Worm-eating Warbler (Helmitheros vermivorum) The Worm-eating Warbler (Helmitheros vermivorum; Figure 88) uses stems of Polytrichum in its nest (Figure 89) (Baicich & Harrison 2005).

Figure 88. Helmitheros vermivorum, Worm-eating Warbler. Members of this species use bryophytes in their nests. Photo by Jerry Oldenettel, through Creative Commons.

Figure 89. Polytrichum commune, a moss in a genus used in nests of Helmitheros vermivorum, Worm-eating Warblers. Photo by Hermann Schachner, through Creative Commons.

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Figure 90. Limnothlypis swainsonii, Swainson's Warbler. Members of this species use bryophytes in their nests. Photo by Carol Foil, through Creative Commons.

Ovenbird (Seiurus aurocapilla) The seclusive Ovenbird (Seiurus aurocapilla; Figure 91-Figure 92) may be dependent on mosses in its environment. Apfelbaum and Haney (1981) reported the disappearance of the Ovenbird from a severely burned Jack pine (Pinus banksiana; Figure 93-Figure 95) forest in the Great Lakes area. In that fire, ~80% of the feather moss (Figure 96) communities suffered severe loss due to the fire. But other factors related to the fire may have caused them to disappear.

Figure 91. Seiurus aurocapilla, Ovenbird, a ground nester that may be dependent on mosses in its habitat. Photo by Tom Grey, with permission.

Figure 92. Seiurus aurocapilla, Ovenbird, nest and nestlings. Photo by Fredlyfish4, through Creative Commons.

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Chapter 16-7: Bird Nests – Passeriformes, part 2

Figure 93. Jack pine (Pinus banksiana) healthy forest. Photo by M. Ricon, through Creative Commons.

Figure 97. Parkesia noveboracensis, Northern Waterthrush. Some members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 94. Pinus banksiana after fire in Baraga, Michigan, USA. Photo by Janice Glime.

Figure 98. Parkesia motacilla, Louisiana Waterthrush. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 95. Burned moss in Jack pine forest, Baraga, MI. Photo by Janice Glime.

Figure 96. Pleurozium schreberi, a feather moss that covers vast areas of ground in conifer forests. Photo by Sture Hermansson, with online permission.

Figure 99. Oporornis agilis, Connecticut Warbler. Members of this species use bryophytes in their nests. Photo from connecticut-warbler-audubon-field-guide, free stock photos.

Chapter 16-7: Bird Nests – Passeriformes, part 2

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of mosses (Figure 104-Figure 106). This nest may be suspended from structures such as logs.

Figure 100. Geothlypis trichas, Common Yellowthroat. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 103. Premnoplex brunnescens, Spotted Barbtail. Members of this species build domed nests of bryophytes. Photo by Murray Cooper, through Creative Commons.

Figure 101. Cardellina pusilla, Wilson's Warbler. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 104. Premnoplex brunnescens, Spotted Barbtail, nest of bryophytes. Photo by Juan Ignacio Areta, through Creative Commons.

Figure 102. Cardellina canadensis, Canada Warbler. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Furnariidae – Neotropical Ovenbirds In the Neotropical ovenbirds (Furnariidae) moss use in nesting materials seems to have at least somewhat followed evolutionary lines (Zyskowski & Prum 1999). Premnoplex brunnescens (Figure 103) builds a domed nest

Figure 105. Premnoplex brunnescens, Spotted Barbtail, nest of bryophytes. Photo by Harold Greeney, through Creative Commons.

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Chapter 16-7: Bird Nests – Passeriformes, part 2

Figure 106. Premnoplex brunnescens, Spotted Barbtail, nest of bryophytes. Photo by Gustavo Londoño, through Creative Commons.

In the Neotropical Cranioleuca albiceps group (see Figure 107), Margarornis (Figure 108-Figure 109), Premnoplex brunnescens (Figure 103-Figure 106), Siptornis (Figure 110), and Plain Softtail, (Phacellodomus fusciceps; see Figure 111), a "pensile" nest (Figure 109) is constructed (Zyskowski & Prum 1999). This is a large nest with a small brood chamber that is entered from below. It is constructed from top down by draping long strands of green mosses or strips of other plant material. The nest hangs down from a log or rocky overhang and in Premnoplex brunnescens it may also hang from vines. Asthenes (Figure 112) species construct an ovoid nest (Figure 113) using fresh Sphagnum (Figure 114). An outer shell of herbaceous stems loosely surrounds it.

Figure 108. Margarornis rubiginosus, Ruddy Treerunner. Members of this species make nests among bryophytes. Photo by Carmelo López Abad, through Creative Commons.

Figure 109. Margarornis squamiger, Pearled Treerunner, pensile nest imbedded in bryophytes and rootlets with an entrance at the bottom. Photo by Harold Greeney, through Creative Commons.

Figure 107. Cranioleuca pallida, Pallid Spinetail, in Brazil. Members of the Cranioleuca albipes group build pensile nests that incorporate bryophytes. Photo by Ciro Albano, through Creative Commons.

Figure 110. Siptornis striaticollis, Spectacled Prickletail, nest. Photo by Harold Greeney, through Creative Commons.

Chapter 16-7: Bird Nests – Passeriformes, part 2

Figure 111. Phacellodomus ruber, Greater Thornbird. Members of this species construct their nests using mosses and other plant material. Photo by Cláudio Dias Timm, through Creative Commons.

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Figure 114. Sphagnum austinii, member of a genus used in nests of Asthenes. Stan Phillips, through public domain

White-browed Spinetail (Hellmayrea gularis) In the Andean cloud forests, the White-browed Spinetail (Hellmayrea gularis; Figure 115) nests (Figure 116) were embedded in hanging masses of epiphytic mosses, but rather than being pendulous, the nests were supported from below or from the sides by stems (Greeney & Zyskowski 2008). These nests were ball-shaped with a side entrance. The exterior consisted of green moss, whereas the internal side consisted of dry bamboo leaves. The nest was lined with soft materials, either Tillandsia seed down (Figure 117) or tree-fern scales (Figure 118).

Figure 112. Asthenes anthoides, Austral Canastero, Members of Asthenes incorporate bryophytes in their nests. Photo by Collaerts brothers, through Creative Commons.

Figure 113. Asthenes flammulata, Many-striped Canastero nest in Ecuador. Photo by Harold Greeney, through Creative Commons.

Figure 115. Hellmayrea gularis, White-browed Spinetail, bringing grub to nest. Photo by Murray Cooper, through Creative Commons.

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Chapter 16-7: Bird Nests – Passeriformes, part 2

Thraupidae – Tanagers & Honeycreepers Wolf (2009) found one species of Thraupidae that use bryophytes in their nests in North America: Piranga ludoviciana (Western Tanager; Figure 119).

Figure 119. Piranga ludoviciana, Western Tanager. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission. Figure 116. Hellmayrea gularis, White-browed Spinetail, nest embedded in mosses. Photo by Harry Greeney, through Creative Commons.

Yellow-bellied Dacnis (Dacnis flaviventer) The Yellow-bellied Dacnis (Dacnis flaviventer; Figure 120) is a bird of the high canopy and nests in this genus are largely unknown. Sheldon and Greeney (2008) were fortunate enough to find one nest and describe it. Although most of the nest is made of ferns, mosses comprise the sparse lining of the cup, woven with rootlets and dried grasses in a circular fashion.

Figure 117. Tillandsia schiedeana; the down (coma) of seeds in this genus are used in the nests of the White-browed Spinetail (Hellmayrea gularis). Photo by Roger Culos, through Creative Commons.

Figure 120. Dacnis flaviventer, Yellow-bellied Dacnis male. Members of this species line their nests with mosses. Photo by Patty McGann, through Creative Commons.

Emberizidae – Emberizines Wolf (2009) found thirteen species of Emberizidae that use bryophytes in their nests in North America:

Figure 118. Hairy tree fern frond showing scales and hairs used in nests of the White-browed Spinetail, Hellmayrea gularis. Photo by Janna Schreier , with permission.

Spizella arborea (American Tree Sparrow; Figure 121-Figure 122) Pooecetes gramineus (Vesper Sparrow; Figure 123-Figure 124) Ammodramus savannarum (Grasshopper Sparrow; Figure 125Figure 126) Passerella iliaca (Fox Sparrow; Figure 127) Melospiza lincolnii (Lincoln’s Sparrow; Figure 128) Zonotrichia albicollis (White-Throated Sparrow; Figure 129)

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Zonotrichia querula (Harris’s Sparrow; Figure 130) Zonotrichia leucophrys (White-Crowned Sparrow; Figure 131Figure 132) Zonotrichia atricapilla (Golden-Crowned Sparrow; Figure 133) Junco hyemalis (Dark-Eyed Junco; Figure 134-Figure 137) Junco phaeonotus (Yellow-Eyed Junco; Figure 138) Calcarius lapponicus (Lapland Longspur; Figure 139-Figure 140) Plectrophenax nivalis (Snow Bunting; Figure 141)

Figure 121. Spizella arborea, American Tree Sparrow. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 124. Pooecetes gramineus, Vesper Sparrow, nestlings in nest, begging. Photo by Kati Fleming, through Creative Commons.

Figure 122. Spizella arborea, American Tree Sparrow, nest and nestlings. Photo from USFWS, through public domain.

Figure 125. Ammodramus savannarum, Grasshopper Sparrow. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 123. Pooecetes gramineus, Vesper Sparrow. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 126. Ammodramus savannarum, female Grasshopper Sparrows in nest. Photo by Janet Ruth, USGS, through public domain.

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Chapter 16-7: Bird Nests – Passeriformes, part 2

Figure 127. Passerella iliaca, Fox Sparrow. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 128. Melospiza lincolnii, Lincoln's Sparrow. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 129. Zonotrichia albicollis, White-throated Sparrow. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 130. Zonotrichia querula, Harris's Sparrow. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 131. Zonotrichia leucophrys, White-crowned Sparrow. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Figure 132. Zonotrichia leucophrys, White-Crowned Sparrow, nest with eggs. Photo by Jacob W. Franks, NPS, through public domain.

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Figure 133. Zonotrichia atricapilla, Golden-crowned Sparrow. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

Junco (Junco hyemalis) The common Junco (Junco hyemalis; Figure 134) spends its winter in snowy places in the northern USA, then returns to even more northern locations in late April to build its nest of grasses, moss, and rootlets nestled in a mossy bank (Figure 135) or along a woodland trail (Figure 136) (Harrison 2000). Ken-ichi Ueda found a similar construction in a stream bank (Figure 137).

Figure 134. Junco hyemalis, Dark-eyed Junco. Members of this species use bryophytes in their nests. Photo by USFWS, through public domain.

Figure 135. Junco hyemalis, Dark-eyed Junco, nest with eggs in mossy cavity. Photo from USFWS, through public domain.

Figure 136. Junco hiemalis, Dark-eyed Junco, nest with Hedwigia ciliata. Photo courtesy of Susan Studlar.

Figure 137. Junco nest in mossy stream embankment. Photo by Ken-ichi Ueda, through Creative Commons.

Figure 138. Junco phaeonotus, Yellow-eyed Junco. Members of this species use bryophytes in their nests. Photo by Tom Grey, with permission.

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Chapter 16-7: Bird Nests – Passeriformes, part 2

Figure 139. Calcarius lapponicus, Lapland Longspur. Members of this species use bryophytes in their nests. Photo by Ómar Runólfsson, through Creative Commons.

Figure 142. Pipilo erythrophthalmus, Eastern Towhee male. Members of this species that use setae of Polytrichum ohioense (Figure 144) to line their nests in the southeastern USA. Photo by Bill Thompson, through Creative Commons.

Figure 140. Calcarius lapponicus, Lapland Longspur, nest. Photo by James K. Lindsey, with permission.

Figure 143. Pipilo erythrophthalmus, Eastern Towhee, nest. Photo by Bill Thompson, through Creative Commons.

Figure 141. Plectrophenax nivalis, Snow Bunting,. Members of this species use bryophytes in their nests. Photo by Cephas, through Creative Commons.

Eastern Towhee (Pipilo erythrophthalmus) The Eastern Towhee (Pipilo erythrophthalmus; Figure 142), formerly the Rufous-sided Towhee, nest (Figure 143) is somewhat unusual in its moss component. The lining can consist of a single material – 70-80 strands of Polytrichum ohioense setae (Figure 144) interwoven to form the lining (Breil & Moyle 1976). A few had gametophyte (leafy plants) fragments or capsules attached.

Figure 144. Polytrichum ohioense showing setae that can be used to line the nests of the Eastern Towhee (Pipilo erythrophthalmus). Photo by Bob Klips, with permission.

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Savannah Sparrow (Passerculus sandwichensis) Mosses comprised more than 30% of the mass of nesting materials in the southeastern Ontario, Canada, populations of the ground-nesting Savannah Sparrow (Passerculus sandwichensis; Figure 145-Figure 146) compared to less than 20% in the northern Manitoba populations (Crossman et al. 2011). Although these differences were not statistically significant (p >0.05), they may reflect the somewhat smaller, more compact nests in the northern Manitoba population. But does it vary with climate as an adaptive means to maintain more favorable temperatures? Indeed Crossman and coworkers found that whereas the external dimensions of the nest did not differ, the inner nest cup was significantly shallower in northern Manitoba, indicating a thicker bottom that could provide greater insulation in the northern Manitoba population. But alas, we do not know if the mosses contributed to any insulating properties.

Figure 145. Passerculus sandwichensis, Savannah Sparrow, a species for which moss usage and nest size vary with latitude. Photo by Tom Grey, with permission. Figure 147. Nest composition for materials comprising ≥1% of nest mass of the Savannah Sparrows (Passerculus sandwichensis; Figure 145-Figure 146) that bred in southeastern Ontario (white bars) and northern Manitoba (grey bars). Bars represent dominant nesting materials ≥1% of nest dry mass. Those materials comprising 30 lemmings per hectare), they rapidly exhaust the graminoids and must live on a diet of 100% mosses (Turchin & Batzli 2001). With the low digestibility of mosses (Batzli & Cole 1979), it is not surprising that captive Nearctic brown lemmings (Lemmus trimucronatus; Figure 58) lost weight on a moss-only diet, supporting the suggestion that mosses must serve some function other than as a source of energy. Batzli and Cole (1979) suggest that the high concentrations of calcium, magnesium, and iron may be beneficial. In a feeding experiment using Funaria hygrometrica (Figure 28), the lemmings of Devon Island ate only the capsules (Pakarinen & Vitt 1974). Pakarinen and Vitt suggested that the choice of capsules may have been related to the high lipid content of the spores. The availability of the highly polyunsaturated fatty acid arachidonic acid (Gellerman et al. 1972) almost exclusively in mosses (and also Equisetum) may be especially important to these small mammals that must run about on and under the snow (Prins 1982b). Northern climates seem to increase the predation on mosses, perhaps because the arachidonic acids might help to keep the fats in the foot pads from changing from a liquid to a solid phase on the cold ground in winter (Prins 1982a), or perhaps because there are fewer choices for food. Arachidonic acid has a low melting point of -49.5oC, supporting the foot pad theory. Few other plants have arachidonic acid, yet it is present in high concentrations in the blood of Arctic animals, perhaps contributing to increased limb mobility and protecting cell membranes at low temperatures. Interestingly, Hansen and Rossi (1991) found that arachidonic acid comprised 30% of the fatty acids in Rhytidiadelphus squarrosus (Figure 69) and Eurhynchium striatum (Figure 70) at 20ºC, but concentrations shifted toward more eicosapentaenoic acid at lower temperatures, with a slight decrease in arachidonic acid. Synaptomys borealis – Northern Bog Lemmings The range of the northern bog lemming (Synaptomys borealis; Figure 71) extends from Alaska, USA, eastward to Labrador, Canada, and southward to southeastern Manitoba, then southward in the USA to Washington, Montana, and northern New England (Clough & Albright 1987; Cassola 2017).

Figure 69. Rhytidiadelphus squarrosus, a species in which dominance of arachidonic acid is shifted to dominance of eicosapentaenoic acid at low temperatures. Photo by Johan N., through Creative Commons.

Figure 70. Eurhynchium striatum with capsules, a species in which dominance of arachidonic acid is shifted to dominance of eicosapentaenoic acid at low temperatures. Photo by J. C. Schou, with permission.

Figure 71. Synaptomys borealis, a species that prefers mossy habitats. Painting by Todd Zalewski, Smithsonian Institutes, through public domain.

Mosses seem to play a prominent role in habitat preference. In the Athabaska-Mackenzie Region of Canada, Preble (1908) reported habitats for the northern bog lemming (Synaptomys borealis; Figure 71). These

Chapter 17-2: Rodents – Muroidea: Non-Muridae

included the border of a small meadow, a wet, swampy area, proximity of small muskeg ponds, and a marsh. To these, Banfield (1974) reported Canada black spruce bogs as the primary habitat, but also wet subalpine meadows, alpine, and sagebrush. In Churchill, Manitoba, Scott and Hansell (1989) found them in the Carex-moss-Salix community and the Salix community; Wrigley (1974) similarly found them in a sedge-moss tundra (Figure 72). Cowan (1939) found them in muskegs in British Columbia, Canada. Booth (1947) also considered them to be inhabitants of wet, boggy places in the North Cascades, Canada, as did Manville and Young (1965) and Osgood (1904) for Alaska, USA. Groves and Yensen (1989) (also Bursik 1993) reported them from Sphagnum bogs (Figure 73) in Idaho, USA, as did Johnson and Cheney (1953) for Idaho and Washington and Layser and Burke (1973) for Washington. In Montana, Reichel and Beckstrom (1993, 1994) found them in thick mats of Sphagnum (Figure 74), and found this habitat to be the best predictor for finding them. For Minnesota, USA, Coffin and Pfannmuller (1988) listed the habitat as dominated by Sphagnum and graminoids, including forested bogs and open ericaceous shrublands. Christian et al. (1999) concurred, but expanded the Minnesota habitats to include spruce forest (Figure 73) with moss on the forest floor, wet alpine meadows, and alpine tundra. Clough and Albright (1987) reported them from wet sedge meadows in the northeastern USA. Near the base of Mount Washington, New Hampshire, USA, Preble (1899) found them in swampy habitats densely carpeted with moss. On the other hand, in Montana, USA, Pearson (1991) found them in an old-growth hemlock Tsuga heterophylla forest (Figure 75) that lacked the typical bog/fen habitat, although most of the sites were more typical.

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Figure 73. Mountain bog/fen in Idaho, USA, with spruce forest in the background. Photo by Robert Marshall, through Creative Commons.

Figure 74. Sphagnum capillifolium, a common bog/fen species. Photo by David Holyoak, with permission.

Figure 72. Sedge-moss tundra, Nunavut, northern Canada. Photo by A. Dialla, through Creative Commons.

In British Columbia, Canada, Cowan (1939) found that Synaptomys borealis (Figure 71) creates a honeycomb of tunnels in the mossy carpets of the muskegs. These tunnels are strewn with fecal pellets, indicating where feeding occurred. The nests are above ground in winter and below ground in summer (Banfield 1974).

Figure 75. Tsuga heterophylla forest. Photo by pxhere, through Creative Commons.

The "house" that is less likely to disappear is a house of Sphagnum (Figure 74) (Cowan 1939). The bog lemmings Synaptomys borealis (Figure 71) usually live in small colonies among the wet mosses (Osgood 1904). Their runways are among the mosses rather than among the

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Chapter 17-2: Rodents – Muroidea: Non-Muridae

grasses and other weeds. Although rare even in Alaska, they tend to be more common in peatlands (Preble 1908; Osgood 1909), where they make nests beneath the moss (Headstrom 1970). For these lemmings in their more southern extensions of their range, where they are also rare, it is in the peatlands that they survive (Coffin & Pfannmuller 1988). Runways not only carry clippings of new bryophyte species, but open habitat to mosses that otherwise could not occur there. Among these in Arctic Alaska is the colonizing species, Funaria polaris (Batzli et al. 1980). While it is clear that mosses, especially Sphagnum (Figure 74), are important in defining the habitat of the northern bog lemming, it is less clear why. Perhaps a small indication is the presence of Hylocomiastrum pyrenaicum (Figure 76) in the mouth of one individual (Harper 1961), but this may just be a gathering to line the nest. Moisture could be an important factor, but there seem to be no physiological studies to test this idea.

Figure 77. Synaptomys cooperi, bog lemming, makes tunnels under Sphagnum. Photo by Phil Myers, through Creative Commons.

Despite its typical bog habitat, Hamilton (1941) found this species in quite different circumstances in Albany County, New York, USA. These "bog" lemmings were in a beech-hemlock forest with a forest floor of spring perennials and lots of black leaf litter. Mosses were apparently not an important component. The bog lemming eats grasses, sedges, mosses, fungi, fruit, bark, and roots (EOL 2017m). Using fecal analysis, Linzey (1984) found that even in southwestern Virginia, USA, the bog lemming subsisted on the broom grass Andropogon (Figure 78) in the summer but on mosses in winter. Both of these foods are low in digestible nutrients.

Figure 76. Hylocomiastrum pyrenaicum, a species that has been seen in the mouth of a northern bog lemming (Synaptomys borealis). Photo by Michael Lüth, with permission.

Rand (1945) provides examples that support this suggestion of the importance of moisture. In this study, seven individuals were captured in wet grassy glades and twelve in marshy sedges of dwarf birch flats (Yukon and Northwest Territories, Canada), although another seven trapped by Rand were in typical spruce swamps with mosses. The common factor is moisture. Synaptomys cooperi – Southern Bog Lemming The bog lemming (Synaptomys cooperi; Figure 77), as its name implies, is a bog species (Connor 1959; Banfield 1974), ranging from southern Manitoba, Canada, south to Arkansas and Tennessee, USA (EOL 2017m). Nevertheless, it can occupy a wide range of habitats, including grasslands, mixed deciduous and coniferous woodlands, spruce-fir forests, and freshwater wetlands (EOL 2017m). In Minnesota, USA, Christian et al. (1999) found that it was significantly more abundant in bogs than in sedge meadows or lowland conifer habitats. Connor (1959) reported it from New Jersey. Goodwin (1932) found this species in Connecticut, USA, on a dark forest floor that was overgrown with ferns, Sphagnum (Figure 74), and other mosses. No surface runways were visible, but there were definite tunnels beneath the surface.

Figure 78. Andropogon virginicus, summer food for the bog lemming (Synaptomys cooperi) in Virginia, USA. Photo by P. B. Pelser, through online permission.

Dicrostonyx – Collared Lemming Once again, we encounter recent changes in our understanding of the species. Dicrostonyx torquatus sensu stricto (Figure 79) is now considered to be distributed only in the Arctic and sub-Arctic tundra and forest-tundra in the Palaearctic region – i.e., in Northern Europe and Asia (Wilson & Reeder 2005). Dicrostonyx is the only rodent (order Rodentia) that changes to white for the winter.

Chapter 17-2: Rodents – Muroidea: Non-Muridae

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Figure 79. Dicrostonyx torquatus, the collared lemming in the Palaearctic region. Photo by Ellicrum, through Creative Commons.

Dicrostonyx groenlandicus – Northern Collared Lemming The northern collared lemming (Dicrostonyx groenlandicus; see related species in Figure 80) is distributed in northern Greenland and Queen Elizabeth Islands to northern North America above the tree line, including northern Alaska, USA (Musser & Carleton, in Wilson & Reeder 2005). Like other genera of lemmings, mosses form part of the diet of Dicrostonyx. Not just any moss will do either. It is perhaps not surprising to learn that northern collared lemmings (Dicrostonyx groenlandicus) graze on Polytrichum (Figure 46-Figure 49) gametophytes during summer on both Devon Island and Ellesmere Island (Pakarinen & Vitt 1974; Longton 1980). But when they were offered fruiting material of Funaria arctica, only capsules were eaten (Pakarinen & Vitt 1974). Pakarinen and Vitt suggested that this preference may be related to the high lipid content of some moss spores. Mosses generally provide less than 10% of the diet of the collared lemming (cf. Figure 79) in Alaska (Batzli & Jung 1980). It appears that this Alaskan lemming must now be Dicrostonyx groenlandicus, although it was reported as D. torquatus. The common sedge Carex aquatilis (Figure 81) contains one or more compounds that are deleterious to collared lemmings (Batzli & Jung 1980). The common evergreen shrub (Ledum palustre; Figure 82) is likewise deleterious to the collared lemming, but also to the tundra vole (Microtus oeconomus; Figure 83) and brown lemmings (Lemmus sibiricus; Figure 68). Differing secondary compounds separate the diets of the two lemmings, but the tundra vole is more of a generalist, overlapping the diets of both lemmings.

Figure 81. Carex aquatilis, a species that is deleterious if eaten by the collared lemming (Dicrostonyx). Photo by Matt Lavin, through Creative Commons.

Figure 82. Ledum palustre with flowers, a species that is deleterious if eaten by the collared lemming (Dicrostonyx). Photo by Kristian Peters, through Creative Commons.

Figure 83. Microtus oeconomus, a species that suffers deleterious effects from eating Ledum palustre. Photo by аимаина хикари, through Creative Commons. Figure 80. Dicrostonyx nelsonii (=D. exsol ), one of three North American species, and a bryophyte consumer. Photo courtesy of Tim Menard.

Gut content analysis indicates that moss capsules form a substantial part of the diet of several North American and Eurasian Arctic lemming species (Batzli & Jung 1980).

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Chapter 17-2: Rodents – Muroidea: Non-Muridae

And Ron Lewis Smith (Bryonet, 21 November 2006) reports large-scale grazing by lemmings on the capsules of Polytrichum (Figure 46-Figure 49) and Polytrichastrum (Figure 44-Figure 45) in northern Sweden. When grazing on capsules, lemmings prefer mature capsules in which the spores have a high lipid content (Pakarinen & Vitt 1974). Wooding (1982) reported the diet of Canadian brown lemmings (Lemmus trimucronatus?; Figure 58) was comprised of willow buds, fruits, flowers, grasses, and twigs. However, in captivity they will eat mushrooms and mosses. This supports the concept that availability is an important determinant of the diet. Rodgers and Lewis (1985) came to an interesting conclusion regarding diet differences between the brown lemming (Lemmus trimucronatus; Figure 58) and the northern collared lemming (Dicrostonyx groenlandicus). The brown lemming preferred graminoids and moss, whereas the northern collared lemming preferred shrubs and herbs. They demonstrated that diet preferences were heritable. The diet preferences for both species were based on macronutrients and caloric content, but the differences between the species depended on secondary compounds and physical characteristics of the plants. They concluded that the northern collared lemming has a greater capacity to deal with secondary compounds or the presence of plant hairs than does the brown lemming.

reflexum (Figure 85), Dicranum fuscescens (Figure 86), D. polysetum (Figure 10), D. scoparium (Figure 11), Hylocomium splendens (Figure 63), Pleurozium schreberi (Figure 8), Ptilium crista-castrensis (Figure 7), Pohlia nutans (Figure 55), Polytrichum commune (Figure 46), P. juniperinum (Figure 48), and Rhodobryum roseum (Figure 87). In eastern Finland, Dicranum and Polytrichum seem to be their favorites, which happen also to have the highest nitrogen content, even though Pleurozium schreberi and Hylocomium splendens are more abundant (Eskelinen 2002). They rejected most herbaceous species, but only rejected a few bryophytes such as Ptilidium ciliare (Figure 9) and Plagiothecium denticulatum (Figure 88) (Kalela et al. 1963a, b). In one area this species used Aulacomnium palustre (Figure 89) extensively, but this seems to be a rare occurrence (Lepp 2008).

Myopus schisticolor – Wood Lemming Wood lemmings, Myopus schisticolor (Figure 84), are distributed in the northern Palaearctic, ranging from western Norway, through Sweden and Finland through northern and central Russia to the Pacific coast and Sakhalin Island (Russia) (Shenbrot & Krasnov 2005). They live in mossy bogs and coniferous forests in cool climates. In the Ural Mountains, they are rare and are restricted to swampy moss habitats (Bolshakov & Berdjugin 1990). Their runways often traverse moss beds as well as under fallen trees and roots. Figure 85. Brachythecium reflexum, one of the preferred forest mosses of the wood lemming. Photo by Michael Lüth, with permission.

Figure 84. Myopus schisticolor by its path through the moss Hylocomium splendens. Photo by Risto S. Pynnönen, through Wikimedia Commons.

Using food preference experiments, Kalela et al. (1963a, b) showed that in northern Sweden, the wood lemmings highly preferred a large number of the most abundant forest mosses, including Brachythecium

Figure 86. Dicranum fuscescens, one of the preferred forest mosses of the wood lemming. Photo by Hermann Schachner, through Creative Commons.

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Dicranum scoparium (Figure 11) > Hylocomium splendens (Figure 63) > Pleurozium schreberi (Figure 64) > Sphagnum girgensohnii (Figure 90). This order provides an interesting contrast to the choices of the heather vole (Phenacomys intermedius; Figure 18, Figure 21-Figure 22) that Kate Frego described. That vole seemed uninterested in Dicranum scoparium. The wood lemming in Finland had some similar preferences to those in Sweden, with Dicranum and Polytrichum (Figure 46Figure 49) as top choices, despite a greater availability of Pleurozium and Hylocomium (Lepp 2008; Figure 91).

Figure 87. Rhodobryum roseum, one of the preferred forest mosses of the wood lemming. Photo by Hermann Schachner, through Creative Commons.

Figure 90. Sphagnum girgensohnii, a preferred moss for food by Myopus schisticolor. Photo by Hermann Schachner, through Creative Commons.

Figure 88. Plagiothecium denticulatum, one of the rejected forest mosses of the wood lemming. Photo by Christian Peters, with permission.

Figure 91. Percent grazing vs cover represented in a lemming habitat in Sweden. Based on data from Lepp 2008.

Figure 89. Aulacomnium palustre, a species that is sometimes eaten as a major food source by the wood lemming. Photo by Kristian Peters, through Creative Commons.

During the snow-free season Myopus schisticolor (Figure 84) feeds on only the green topshoots of the mosses, whereas during the snow-covered season, these lemmings bite off the shoots at the base (Kalela et al. 1963a, b). Their order of preference in Sweden seems to be

The species choices changed somewhat in the winter storage holes, which were located in drier sites (Lepp 2008). About 85% of their stored mosses were Dicranum (Figure 10), 11% Pleurozium schreberi (Figure 64), and only 3% Hylocomium splendens (Figure 63). They did still forage in winter, still preferring Dicranum, but their second highest nibblings were on Ptilium (Figure 7), which occurred in only 30% of the study plots. In fact, for whatever reason, they did not forage on Polytrichum (Figure 46-Figure 49) in winter, despite its greater abundance than that of Ptilium. The wood lemming will graze for a long time on the same moss species, hence making it possible to identify its recent food by the color of the feces (Lepp 2008). Those with Pleurozium schreberi (Figure 64) and Hylocomium splendens (Figure 63) are light brown, Polytrichum

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(Figure 46-Figure 49) dark brown, Dicranum (Figure 10) dark green, and Ptilium crista-castrensis (Figure 7) light green. One explanation for the choice of mosses for the wood lemming may be the nitrogen content (Lepp 2008). Dicranum (Figure 10) and Polytrichum (Figure 46-Figure 49) have the highest nitrogen content among the mosses in the study area. Secondary compounds such as phenols may discourage consumption of some species that are abundant, but no data are available for the study site. Since such content could differ based on environmental conditions, we can only speculate. On the other hand, Eskelinen (2002) suggested that the high carbon:nitrogen content of Dicranum (Caut et al. 2009; Codron et al. 2011) might account for Dicranum as the preferred food, and sometimes only food, for this species in Finland. Ericson (1977) found that Myopus schisticolor (Figure 84) had a high preference for many forest moss species in preference experiments. Their preferred mosses were Dicranum scoparium (Figure 11), Hylocomium splendens (Figure 63), Pleurozium schreberi (Figure 64), and Sphagnum girgensohnii (Figure 90). In fact, they rejected most of the herb species. Some bryophytes were also rejected, including the leafy liverwort Ptilidium ciliare (Figure 9) and the moss Plagiothecium denticulatum (Figure 88). In summer the wood lemming eats only the green tops of shoots, but in winter when the bryophytes are snow covered, they eat them down to the base. Young wood lemmings cannot survive on mosses alone; to grow faster, they need to eat other plants as well (Andreassen & Bondrup-Nielsen 1991; Lepp 2008). Adults, however, can subsist on mosses alone. Nevertheless, both growth and reproduction are negatively affected when the diet is 100% moss, compared with a diet that also includes grasses and shrubs.

Figure 93. Cryptomys hottentotus adult showing dense fur. Photo by Daderot, through Creative Commons.

Myoxidae – Dormice and Hazel Mice Muscardinus avellanarius – Hazel Dormouse In England, the hazel dormouse (Muscardinus avellanarius; Figure 94), a somewhat rare nocturnal rodent, gets its name from the Anglo-Norman term dormeus, which means "sleepy" (Wikipedia 2008). This refers to its habit of becoming torpid and cold in the winter, waking only occasionally to eat food stored nearby. Hibernation is triggered by temperatures below 16ºC (Habril & Passig 2008).

Bathyergidae – Blesmoles and Mole Rats Cryptomys hottentotus – Hottentot Mole-rat The Hottentot mole-rat (Cryptomys hottentotus; Figure 92Figure 93) is widely distributed in South Africa (Bishop et al. 2004). Colonies have 2-14 individuals that permanently live in a network of burrows, locating their food as they burrow (Spinks 1998) The Hottentot mole-rat builds hummocks through its burrowing activity (Lynch 1992) in mesic bog soils (Bishop et al. 2004). It may not need a mossy habitat, but some mosses seem to benefit from its presence. The excavated soil is colonized by a lawn-like cover that includes mosses (Lynch 1992).

Figure 92. Cryptomys hottentotus (Hottentot mole-rat), a species that creates habitat for some mosses. Photo by Lloyd Glenn Ingles, through Creative Commons.

Figure 94. Muscardinus avellanarius – hazel dormouse, a species that uses mosses in its winter hibernacula. Photo by Danielle Schwarz, through Creative Commons.

Its habitat is typically an unshaded understory where there is high species diversity (Bright & Morris 1990). Bright and Morris (1991) contend that this species is entirely arboreal, detouring considerable distances to avoid crossing open ground. They seldom venture more than 100 m from the nest. They seem to prefer nesting in tree hollows, but when these are scarce they select a location with shrub cover and proximity to the forest edge (Berg & Berg 1998). Despite living in trees, they do not seem to include mosses in the diet (Bright & Morris 1993). Mosses may be more important for a hibernaculum (shelter occupied during the winter by a dormant animal). The hazel dormice hibernate in winter, 6-7 months in Lithuania (Juškaitis 1999). Bright and Morris (1996) reported that the dormice covered their surface hibernaculum with a thin layer of mosses or leaves. Such shallow surface hibernacula make the hibernating animals vulnerable to floods, trampling, and predation (Juškaitis 1999).

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In a Ukraine study, Zaytseva (2006) found that mosses comprise about 5% of the nesting material in nest boxes used by the hazel dormouse, which sleeps there throughout the day. The globose summer nest is shaped much like a wren's nest with a door (Habril & Passig 2008). Both summer and winter nests often have mosses in them, but the winter nest is more likely to be in a tree hollow or stump. Some dormice may spend their winter on the ground under moss and litter. Van Laar and Dirkse (2010) examined the nesting materials and found that this species used the epiphytic mosses Brachythecium rutabulum (Figure 95) and Orthotrichum lyellii (Figure 96). But they also used the primarily ground-dwelling species Cirriphyllum piliferum (Figure 97), Hypnum cupressiforme (Figure 4), Calliergonella cuspidata (Figure 98), Eurhynchium hians (Figure 99), and Thuidium assimile (Figure 100). All nest materials were pleurocarpous mosses. Van Laar and Dirkse considered the moss choice to be due to the physical properties of the moss that helped the hazel dormouse to maintain a certain degree of humidity in the nests.

Figure 95. Brachythecium rutabulum, an epiphyte used for nesting material by the hazel dormouse, Muscardinus avellanarius. Photo by Michael Lüth, with permission.

Figure 96. Orthotrichum lyellii, an epiphyte used for nesting material by the hazel dormouse, Muscardinus avellanarius. Photo by Michael Lüth, with permission.

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Figure 97. Cirriphyllum piliferum, a ground species used as nesting material for the hazel dormouse, Muscardinus avellanarius. Photo by Michael Lüth, with permission.

Figure 98. Calliergonella cuspidata, a ground species used as nesting material for the hazel dormouse, Muscardinus avellanarius. Photo by Tim Waters, through Creative Commons.

Figure 99. Eurhynchium hians, a ground species used as nesting material for the hazel dormouse, Muscardinus avellanarius. Photo by Michael Lüth, with permission.

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Figure 100. Thuidium assimile, a ground species used as nesting material for the hazel dormouse, Muscardinus avellanarius. Photo by Hermann Schachner, through Creative Commons.

1978), a fact noted much earlier in Britain by Tripp (1888). These bryophytes are useful in building suitable nests. Even in arboreal habitats at warmer latitudes, the Japanese dormouse uses bryophytes for its lair (Watanabe 1978; Minato & Doei 1995; Doei & Minato 1998). After examining 21 nests, Minato and Doei (1995) reported 42 species of mosses and 15 species of liverworts as constituting the majority (53.1% by weight) of the nest materials. Like most of the bird nest bryophytes, the majority of those used by the Japanese dormouse were pleurocarpous, and consistent with the dormouse habitat, they were mostly epiphytic. The six most commonly used species were the leafy liverwort Frullania tamarisci subsp. obscura (Figure 103), and the mosses Hypnum tristoviride (Figure 104), Isothecium subdiversiforme (Figure 105), Anomodon rugelii (Figure 106), Entodon scabridens, Anomodon longinervis. The leafy liverwort Frullania tamarisci subsp. obscura was often the most abundant bryophyte in the nest. This species is typically abundant nearby, spreading over the surface of tree trunks in large mats, often making it easier for the dormouse to harvest.

Gliridae – Dormouse Glirulus japonicus – Japanese Dormouse The Japanese dormouse (Glirulus japonicus; Figure 101)), an endemic to Japan, is nocturnal, searching a relatively large area to find food at night (EOL 2017b). Its name derives from the Anglo-Norman word dormeus, which means sleepy one. However, it is not its daytime sleeping that gives it this name, but rather its long hibernation period. The males awaken in May to find a mate.

Figure 102. Glirulus japonicus sleeping in nest. Photo by Yamaneseisokubunpuik, through Creative Commons.

Figure 101. Glirulus japonicus, a species that uses bryophytes in its lair. Photo by Katuuya, through Creative Commons.

It easily climbs trees, where it feeds on seeds, fruits, insects, and bird eggs (EOL 2017b). It can run as easily on the lower side of a branch as on the upper side. This species lacks a caecum, and thus should not be expected to digest cellulose, making mosses an inefficient food and explaining their absence in the dormouse diet. The Japanese dormouse (Glirulus japonicus; Figure 101) uses bryophytes in its lair (Figure 102) (Watanabe

Figure 103. Frullania tamarasci subsp obscura, a matforming pleurocarpous moss used for nesting material by the Japanese dormouse (Glirulus japonicus). Photo from , through Creative Commons.

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Myoxus glis – Fat Dormouse; Edible Dormouse The fat dormouse (Myoxus glis; Figure 107) occurs throughout much of mainland western Europe and on a number of Mediterranean islands (Milazzo et al. 2003).

Figure 104. Hypnum tristo-viride, a pleurocarpous moss used for nesting material by the Japanese dormouse (Glirulus japonicus). Photo by Jiang Zhenyu, Mou Shanjie, Xu Zaiwen, and Chen Jianzhi, through Creative Commons.

Figure 105. Isothecium subdiversiforme, a pleurocarpous moss used for nesting material by the Japanese dormouse (Glirulus japonicus). Photo from Digital Museum, Hiroshima University, with permission.

Figure 107. Myoxus glis, a species that eats mosses, but most likely accidentally. Photo by Marcus Ostermann through Creative Commons.

Gigirey and Rey (1998) reported that 12 of 32 stomachs of the fat dormouse, Myoxus glis (Figure 107), had moss remains. Gigirey and Rey (1999) subsequently found mosses of this species in the feces. However, in both cases they considered these mosses to be ingested accidentally. Whereas mosses may not be a desirable diet item, they do provide nesting materials (Drăgoi & Faur 2013). They typically construct these nests using leaves and mosses (Grzimek 2003). The mosses are typically pleurocarpous mosses, including the epiphytes Brachythecium rutabulum (Figure 95), Isothecium myosuroides (Figure 108), and Eurhynchium praelongum (Figure 109), but also nearby forest floor species including Brachythecium glareosum (Figure 110), Ctenidium molluscum (Figure 111), Eurhynchium striatum (Figure 70), and Eurhynchium hians (Figure 99) (van Laar & Dirkse 2010).

Figure 106. Anomodon rugelii, a pleurocarpous moss used for nesting material by the Japanese dormouse (Glirulus japonicus). Photo by Janice Glime.

Watanabe (1978) found 25 bryophyte species in 8 nests. He found an average of 4 bryophyte species per nest, whereas Minato and Doei (1995) found an average of 6.8 species.

Figure 108. Isothecium myosuroides, a pleurocarpous epiphyte used for nesting by the edible dormouse (Myoxus glis). Photo by Malcolm Storey, DiscoverLife, with online permission.

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Dryomys nitedula – Forest Dormouse The forest dormouse (Dryomys nitedula; Figure 112) lives in Switzerland through eastern and southern Europe, Asia Minor and the Caucasus to central Russia and central Asia. It is a tree dweller, living in forests (EOL 2017n).

Figure 109. Eurhynchium praelongum, a pleurocarpous epiphyte used for nesting by the edible dormouse (Myoxus glis). Photo by Janice Glime. Figure 112. Dryomys nitedula, the forest dormouse. Photo by Domodi, through Creative Commons.

Like Myoxus glis (Figure 107), Dryomys nitedula (Figure 112) uses mosses in its nests (Drăgoi & Faur 2013). The nests are round with either a side or top entry. The exterior is rough, constructed of branches, but the interior is padded, using grasses, feathers, hair, or mosses. And like the fat dormouse, Dryomys nitedula sometimes uses empty bird nests (Adamik & Kral 2008).

Summary Figure 110. Brachythecium glareosum, a pleurocarpous ground species used for nesting by the edible dormouse (Myoxus glis). Photo by Michael Lüth, with permission.

Figure 111. Ctenidium molluscum, a pleurocarpous ground species used for nesting by the edible dormouse (Myoxus glis). Photo by Michael Lüth, with permission.

They locate their nests high in trees, using the cup formed by branching, although some may use abandoned bird nests (Juškaitis 2006).

Many rodents have mosses in the gut and feces, but these seem to be the result of accidental intake. But some seem to include them as an important part of the diet, often increasing the percentage in winter. Researchers have suggested that this switch may be a need for nitrogen, arachidonic acid, or fiber. In other cases, it may be a simple matter of availability. The shoot tips seem most desirable for food, but in winter the moss may be clipped at the bottom. Some records indicate that moss capsules are eaten. Known consumers of mosses include Chionomys nivalis, and several members of Microtus, Phenacomys, Peromyscus maniculatus (capsules). Lemmings, in particular, are dependent on mosses in the diet. These may provide arachidonic acid, a more pliable fatty acid at cold temperatures. When their population peaks, they may destroy their moss cover under the snow, making them dangerously visible to predators when the snow melts. Many rodents use mosses in the construction of nests, particularly as part of the lining. In bogs, several species may coexist in a single bog, some using them for food or to make nests, tunnels, or runways. Pleurocarpous mosses are preferred by most of the rodents that use mosses as nesting materials. Bryophytes are impacted by the rodents in multiple ways: diminished cover, competition from flowering

Chapter 17-2: Rodents – Muroidea: Non-Muridae

plants. But at other times they may benefit. The rodents can serve as dispersal agents, and runways and burrow openings open new habitats where colonizers like Funaria can grow, increasing diversity.

Acknowledgments This chapter has benefitted greatly from anecdotal records sent to me by bryologists and friends who observed these small rodents interacting with mosses in the field. Steve Juntikka, a former plant taxonomy student of mine, sent me an excited email from Isle Royale after observing the young mouse devouring capsules of Funaria hygrometrica. Leah Vucetich and Rolf Peterson, Isle Royale researchers, provided me with the identification of the juvenile Peromyscus maniculatus based on the picture alone.

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Glime, J. M. 2018. Rodents and Bats – Non-Muroidea. Chapt. 17-3. In: Glime, J. M. Bryophyte Ecology. Volume 2. Bryological Interaction. eBook sponsored by Michigan Technological University and the International Association of Bryologists. Last updated 19 July 2020 and available at .

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CHAPTER 17-3 RODENTS AND BATS – NON-MUROIDEA TABLE OF CONTENTS Soricomorpha .................................................................................................................................................... 17-3-2 Soricidae – Shrews ..................................................................................................................................... 17-3-2 Sorex cinereus – Long-tailed Shrew ................................................................................................... 17-3-3 Sciuromorpha .................................................................................................................................................... 17-3-4 Sciuridae .................................................................................................................................................... 17-3-4 Tamias merriami – Merriam Chipmunk ............................................................................................. 17-3-4 Tamiasciurus hudsonicus – American Red Squirrel ........................................................................... 17-3-4 Sciurus vulgaris – Eurasian Red Squirrel ........................................................................................... 17-3-5 Sciurus carolinensis – Grey Squirrel .................................................................................................. 17-3-6 Spermophilus parryii – Arctic Ground Squirrel.................................................................................. 17-3-6 Glaucomys – Flying Squirrels ............................................................................................................. 17-3-7 Glaucomys sabrinus – Northern Flying Squirrel ................................................................................ 17-3-7 Glaucomys volans – Southern Flying Squirrel.................................................................................... 17-3-7 Lagomorpha – Hares, Rabbits, and Pikas.......................................................................................................... 17-3-7 Leporidae – Rabbits and Hares .................................................................................................................. 17-3-7 Lepus arcticus – Arctic Hare .............................................................................................................. 17-3-7 Oryctolagus cuniculus – European Rabbit .......................................................................................... 17-3-8 Ochotonidae – Pikas................................................................................................................................. 17-3-12 Ochotona princeps – American Pika ................................................................................................ 17-3-12 Ochotona collaris – Collared Pika .................................................................................................... 17-3-14 Erinaceidae – Hedgehogs ......................................................................................................................... 17-3-15 Chiroptera – Bats............................................................................................................................................. 17-3-15 Pteropidae – Flying Foxes........................................................................................................................ 17-3-15 Pteropus conspicillatus – Spectacled Flying Fox ............................................................................. 17-3-15 Summary ......................................................................................................................................................... 17-3-16 Acknowledgments ........................................................................................................................................... 17-3-17 Literature Cited ............................................................................................................................................... 17-3-17

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Chapter 17-3: Rodents and Bats – Non-Muroidea

CHAPTER 17-3 RODENTS AND BATS – NON-MUROIDEA

Figure 1. Lepus arcticus in its summer coloring. Photo from Gilad.rom, through Creative Commons.

Soricomorpha Soricidae – Shrews In 25 bogs and ombrotrophic mires of Poland, Ciechanowski et al. (2012) found that shrews dominated among the mammals captured in pitfall traps. The traps produced 598 individuals distributed among 12 mammal species. Typical wetland species included Neomys fodiens (Eurasian water shrew; Figure 2), Neomys anomalus (Mediterranean water shrew; Figure 3), and Microtus oeconomus (tundra vole; Figure 4). The most numerous species was the Eurasian pigmy shrew (Sorex minutus; Figure 5), and it was sometimes the only rodent present in the habitat. It was most common in undisturbed, treeless parts of bogs where Sphagnum (Figure 6) dominated.

Figure 2. Neomys fodiens, The Eurasian water shrew, a typical wetland species that is found in bogs and mires. Photo from Saxifraga – Rudmer Zwerver, with online permission.

Chapter 17-3: Rodents and Bats – Non-Muroidea

Figure 3. Neomys anomalus (Eurasian water shrew), a typical wetland species that is found in bogs and mires. Photo by Mnolf, through Creative Commons.

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Sorex cinereus – Long-tailed Shrew The long-tailed shrew (Sorex cinereus; Figure 7) occurs from Alaska, USA, east to Labrador/Newfoundland, Canada, south in the USA to Washington, Utah, New Mexico, Northern Great Plains, southern Indiana and Ohio, through the Appalachian Mountains to northern Georgia and western South Carolina, and on the east coast to New Jersey and northern Maryland, where it commonly occurs with mosses (Youngman 1975; Whitaker 2004). It seems often to be present in traps set for lemmings. Hamilton (1941) found Sorex cinereus near the summit of Big Black Mountain in Harlan County, Kentucky, USA, at ~1220 m. Of these, six of the seven specimens were taken from runways at the sides of moss-covered logs in damp, deciduous thickets. In the thickets of Maine and New Hampshire, USA, traps set for lemmings also captured shrews (Clough & Albright 1987). These included Blarina brevicauda (northern short-tailed shrew; Figure 8) and Sorex cinereus. Groves and Yesen (1989) likewise found species of Sorex in lemming traps in a Sphagnum "bog" in Idaho, USA (Figure 9), as did Pearson (1991) in Glacier National Park and Reichel and Beckstrom (1993) in western Montana.

Figure 4. Microtus oeconomus (tundra vole), a typical wetland species. Photo from Saxifraga, Janus Verkerk, with online permission.

Figure 5. Sorex minutus (Eurasian pigmy shrew), the most common rodent species in Polish bogs. Photo from Saxifraga – Rudmer Zwerver, with online permission.

Figure 6. Sphagnum rubellum, in a genus that dominates bogs. Photo by Michael Lüth, with permission.

Figure 7. Sorex cinereus (long-tailed shrew), a species that seems to have an affinity for moss-covered logs in its runways. Photo by Phil Myers, through Creative Commons.

Figure 8. Blarina brevicauda (northern short-tailed shrew), a species caught in lemming traps in thickets of Maine and New Hampshire, USA. Photo by Gilles Gonthier, through Creative Commons.

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Figure 9. Mountain bog (poor fen?) in Idaho, USA. Photo by Robert Marshall, through Creative Commons.

Figure 11. Chipmunk (Tamias merriami), caught in the act by a camcorder as it eats mosses, Syntrichia princeps). Photo courtesy of Brent Mishler.

Sciuromorpha Sciuridae Records indicating that squirrels use mosses to line their nests are old (Tripp 1888). But sometimes, the mosses use squirrel activity to their advantage (Ken Adams, Bryonet 30 April 2020). In the Epping Forest, UK, Zygodon viridissimus competes with Z. forsteri for space on the grooves created by squirrel gnawing. The former often out-competes the latter. Tamias merriami – Merriam Chipmunk The Merriam chipmunk (Tamias merriami) has a small distribution in central and southern California, USA (Harvey & Polite 1999). There seems to be little documentation of chipmunks eating or using mosses. Imagine the surprise when Brent Mishler and his team (Mishler & Hamilton 2002) caught a chipmunk (Figure 10Figure 11) grabbing a chunk of the moss Syntrichia princeps (Figure 12-Figure 13) from the very middle of their field of view (Figure 12) through a CAMcorder (see Grant et al. 2006 for setup). Mishler (pers. comm. 12 January 2008) suggests that the Merriam chipmunk (Tamias merriami; Figure 10-Figure 11) may have been after the water adhering to the moss (Syntrichia princeps), as it had just been moistened earlier in the day for an experiment; the surroundings were dry.

Figure 12. Syntrichia princeps with red ellipse indicating where moss was removed by Tamias merriami. Photo courtesy of Brent Mishler.

Figure 13. Syntrichia princeps with capsules. Photo by F. Guana, Modoc National Forest.

Tamiasciurus hudsonicus – American Red Squirrel

Figure 10. Tamias merriami, a chipmunk that harvests mosses. Photo by James Maughn, through Creative Commons.

The American red squirrel (Tamiasciurus hudsonicus; Figure 14) seems to eat just about anything. It is more tame than most squirrels, and I have even had a confused squirrel climb my leg! It also seems to like decorating its

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abode, using paper, moss, and other local objects it can find (Hanrahan 2012).

Figure 16. Sciurus vulgaris, a species that uses pleurocarpous mosses as nesting materials. Photo from Saxifraga – Mark Zekhuis, with online permission. Figure 14. Tamiasciurus hudsonicus (American red squirrel) uses mosses to decorate its home. Photo by Cephas, through Creative Commons b

Sciurus vulgaris – Eurasian Red Squirrel The Eurasian red squirrel (Sciurus vulgaris; Figure 15-Figure 16) is distributed across the northern parts of Europe (Greene 1887). It makes a nest in the fork of a tree. This nest is an interwoven structure of twigs, leaves, and mosses.

Figure 17. Hypnum cupressiforme, a moss used in nests of Sciurus vulgaris. Photo by Michael Lüth, with permission.

Figure 15. Sciurus vulgaris, a species that uses pleurocarpous mosses in its nest boxes. Photo from Saxifraga – Mark Zekhuis, with online permission.

Nest boxes used by the Eurasian red squirrel (Sciurus vulgaris; Figure 15-Figure 16) displayed pleurocarpous mosses (van Laar & Dirkse 2010). Two of these were ground species [Hypnum cupressiforme (Figure 17), Homalothecium sericeum (Figure 18)]. The Eurasian red squirrel used only one epiphytic species (Orthotrichum sp.; Figure 19), but van Laar and Dirkse suggested that all of the mosses may have been collected from a nearby tree. The nest included ~470 g spruce twigs and ~180 g of these mosses. In addition, the squirrel had included insulation material from the roof of a nearby house. Quinton (1997) reported finding a nest under Sphagnum (Figure 6) in the boreal forest of North America.

Figure 18. Homalothecium sericeum, a moss used in nests of Sciurus vulgaris. Photo by Michael Lüth, with permission.

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The summer nest is typically flimsy and located among small branches.

Figure 19. Orthotrichum cupulatum with capsules, a moss used in nests of Sciurus vulgaris. Photo by Jutta Kapfer, with permission.

Pulliainen and Raatikainen (1996) studied the effect of various nesting materials on nest temperature of the red squirrel in Finland. The wind speed had a large effect on differences between inside and outside the nest. During windless times, the temperature difference could be as much as 30ºC in nests made of mosses, proving mosses to be superior insulators to the beard lichen (Usnea; Figure 20). Juniper bark provided the poorest insulation among the materials tested. A plastic plate under grass greatly increased the inside temperature by restricting the air current throughout the nest.

Figure 20. Usnea filipendula, a nest material that has less insulating ability than the tested mosses. Photo by Jerzy Opioła, through Creative Commons.

TalkTalk (2011) describes the nest of the red squirrel as having a layer of twigs with a layer of moss or bark fragments. It is likely that availability is a major influence on the nest materials used. Sciurus carolinensis – Grey Squirrel The grey squirrel (Sciurus carolinensis; Figure 21Figure 22) lives in the eastern USA, but is an invasive in Europe (Steele et al. 1996; Goheen & Swihart 2003). It builds a nest the size of a football (YPTE 2011). It is comprised of twigs, often with their leaves remaining attached, and is perched high in a tree. The squirrels line the nest with dry grass, shredded bark, moss, and feathers.

Figure 21. Sciurus carolinensis, grey squirrel, a species that uses mosses as one of its nest lining materials. Photo by Janice Glime.

Figure 22. Sciurus carolinensis, a species that uses mosses as one of its nest lining materials. Photo by John White, with permission.

Spermophilus parryii – Arctic Ground Squirrels Like the pikas, it appears that Arctic ground squirrels (Spermophilus parryii; Figure 23-Figure 24) survive winter in the "warmth" of hibernacula (Barnes 1989). These rodents can wake up and run around when their core temperature is as low as -2.9°C. Temperatures much lower than that can be lethal for such small homeotherms. Maintenance of a temperature as low as -3°C could save up to ten times as much energy as maintenance of a body temperature above 0°C. It is quite possible that for the pikas, the mosses permit the maintenance of sufficiently "warm" temperatures to survive.

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Figure 23. Spermophilus parryii and tunnel entrances. Photo from National Park Service, through public domain. Figure 25. Northern flying squirrel, Glaucomys sabrinus, a species that uses mosses in its nests. Photo by Phil Myers, through Creative Commons.

The northern flying squirrel (Glaucomys sabrinus; Figure 25) builds a cavity nest, using various mosses (Patterson et al. 2007). Patterson and coworkers found trace amounts of peat moss (Sphagnum; Figure 6), dried grasses, cedar leaves, and twigs in the nests in southern Ontario. Glaucomys volans – Southern Flying Squirrel

Figure 24. Spermophilus parryii, Arctic ground squirrel, a species that seems to benefit from the insulating ability of mosses in the nest. Photo Jim McCarthy, through public domain.

The smaller southern flying squirrels (Glaucomys volans; Figure 26) occur along the southern USA north to New England (Marchand 2001). They have tiny bodies, weighing only 57-113 g. They are nocturnal, thus most people have never seen them. Nevertheless, they are the most abundant squirrel in the eastern US.

Arctic ground squirrels actually cache bryophytes. They preferentially decapsulate bryaceous mosses and store the capsules in their nests for winter food reserves (Zazula et al. 2006). Nest materials for these Arctic ground squirrels in the Yukon include mosses and lichens and these are the most common materials found in the pouches of females (Gillis et al. 2005). Carrying these materials was most common prior to and during lactation. These mosses and lichens are absent in male pouches. Glaucomys – Flying Squirrels Glaucomys are active all year, but have little resistance to cold (Marchand 2001). Instead, they keep warm by huddling together in tree cavities lined with grass, moss, or bark. The nests can be as much as 30º warmer than the surrounding air outside the nest. These huddles typically have about 10 squirrels, but there may be as may as 50.

Figure 26. Southern flying squirrel, Glaucomys volans, a species that uses mosses in its nests. Photo by Ken Thomas, through Creative Commons.

Glaucomys sabrinus – Northern Flying Squirrel

Leporidae – Rabbits and Hares

The northern flying squirrels (Glaucomys sabrinus; Figure 25) has a wide distribution throughout northern North America from Alaska, across Canada to the eastern provinces, with several extensions into northern USA. Like the southern flying squirrel, this squirrel is nocturnal (IUCN 2017).

Lagomorpha – Hares, Rabbits, and Pikas Lepus arcticus – Arctic Hare In the high Arctic, the Arctic hare (Lepus arcticus; Figure 1, Figure 27) seems to prefer eating developing bryophyte capsules (Catherine LaFarge, Bryonet 30 March 2016). LaFarge often found decapitated sporophytes, although the lemmings helped in the consumption.

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Figure 29. Dicranum scoparium with capsules, a species that the European rabbit dislikes. Photo by Janice Glime. Figure 27. Lepus arcticus in white phase. Chmee2, through Creative Commons.

Photo by

Oryctolagus cuniculus – European Rabbit The European rabbit (Oryctolagus cuniculus) is present in all Western European countries, Ireland and UK, Austria, Sweden, Poland, Czech Republic, Hungary, Romania, Ukraine, and Mediterranean, Croatia, and Slovakia (Smith & Boyer 2008). Rabbits, with their noses to the ground, would seem ideally suited for nibbling on bryophytes. However, it seems they may not find them to their liking. Bhadresa (1977) reported that in a food preference test, the rabbit Oryctolagus cuniculus (European rabbit – the only domesticated rabbit; Figure 28) in Norfolk – actually disliked Dicranum scoparium (Figure 29). But then, that is only one moss. Davidson et al. (1990) found leaf fragments of Mnium (Figure 30-Figure 31), Brachythecium (Figure 32), Hypnum (Figure 17), and Polytrichum (Figure 36) species in feces of rabbits in southeast England, but never forming more than 5% of the plant material in a fecal pellet. Rabbits eat a mixed diet (European Rabbit 2009), and it appears that mosses may be part of it – or they are ingested accidentally.

Figure 28. European rabbit, Oryctolagus cuniculus, a species that consumes at least some mosses. Photo by Aiwok, through Creative Commons.

Figure 30. Mnium spinosum cushions, in a genus found in the feces of the European rabbit. Photo by George Shepherd, through Creative Commons.

Figure 31. Mnium spinosum, in a genus found in the feces of the European rabbit. Photo by Michael Lüth, with permission.

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Figure 34. Ceratodon purpureus, a species that rebuilds organic matter after a fire. Photo by Janice Glime. Figure 32. Brachythecium rutabulum, in a genus found in the feces of the European rabbit. Photo by J. C. Schou, with permission.

Rabbits can have a negative impact on bryophytes. After a fire in the heathlands of Brittany, rabbits, along with roe-deer, damaged the bryophytes by scraping (Clément & Touffet 1981). The bryophytes were important as initial colonizers after the fire, so the scraped areas suffered from their loss in succeeding plant and animal colonization. The mosses Funaria hygrometrica (Figure 33) and Ceratodon purpureus (Figure 34) are important in rebuilding the organic matter following fires and their loss is unfavorable to invertebrate development. Polytrichum s.l. species have a strong competitive ability compared to tracheophytes in colonizing these nutrient-poor sites. In particular, Polytrichastrum formosum (Figure 35) and Polytrichum commune (Figure 36) have a higher density and growth rate and can produce 7-8 tons ha-1 yr-1, preventing new species from becoming established and retarding the growth of those already present. As in cases with other rodents, the rabbits may facilitate the development of these Polytrichaceae colonies.

Figure 35. Polytrichastrum formosum with capsules, a species that is highly competitive on nutrient-poor sites opened up by browsing. Photo from UBC Botany website, with permission.

Figure 36. Polytrichum commune, a species that is highly competitive on nutrient-poor sites opened up by browsing. Photo by Michael Lüth, with permission.

Figure 33. Funaria hygrometrica, a species that rebuilds organic matter after a fire. Photo by Michael Lüth, with permission.

But rabbits (Oryctolagus cuniculus; Figure 28) can also create habitat for bryophytes. Callaghan (2015) reports that some mosses thrive due to grazing activities by rabbits in the UK. A more spectacular find occurred at an old tin works in Cornwall, where the rare copper moss Scopelophila cataractae (Figure 37) benefits by the creation of habitats by rabbits. As succession proceeds on the exposed mineral soil, the tracheophytes replace the bryophytes. However, when the rabbits arrive, the rabbits

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create a network of runways and tunnels, exposing the metal-rich soil where the copper moss thrives. These serve as refugia for this moss species that is disappearing as the more coarse vegetation develops. The entrances to burrows are clothed in a mat of protonemata (Figure 38) that have abundant gemmae (Figure 39). Callaghan speculates that the rabbits must disperse thousands of these gemmae on their fur, and the entrance to the tunnel is often the benefactor substrate.

Figure 37. Mature plants of Scopelophila cataractae, a species that benefits from rabbits making tunnels. Photo by Blanka Shaw, with permission.

The European rabbit has multiplied from the 24 introduced to Australia in 1859 to over 600 million in less than a century (European Rabbit 2009), suggesting that this rapid multiplier could present considerable destruction to mosses, or could favor their increase by destroying lichens. In areas where rabbits have been introduced, they often have no natural enemies. Australia is a case in point. In such cases, the virus causing myxomatosis may be their only enemy. While this has been used successfully to help control the rabbits, the ones currently remaining in Australia are now immune to it. In a dune system in Wales, the advent of myxomatosis caused changes in the vegetation. This area had been the site of severe rabbit grazing. In 1954, myxomatosis began to spread to the area and Ranwell (1960) anticipated the loss to the rabbit population. In May of 1955 rabbit pellets were common and thick on the transects across turf areas. Mosses were very evident among the 1-2 cm high turf, but were much less evident in the deep turf. During the succeeding years of rabbit decline, grasses, sedges, and pleurocarpous mosses [Ditrichum flexicaule (Figure 40), Pseudoscleropodium purum (Figure 41), Rhytidiadelphus squarrosus (Figure 42), R. triquetrus (Figure 43)] increased, surviving in the ungrazed turf. Eurhynchium praelongum (Figure 44) and Plagiomnium undulatum (Figure 45) also increased during the study period. At the same time, decreases were evident in the acrocarpous mosses Bryum sp. (Figure 46), Climacium dendroides (Figure 47), Dicranum scoparium (Figure 29), Syntrichia ruralis (Figure 48). Rhodobryum roseum (Figure 49) disappeared from 1955 to 1958. Overall, the bryophyte richness remained unchanged. The greatest losses of mosses occurred only after 3-4 years of recovery from grazing.

Figure 38. Scopelophila cataractae protonemata in a rabbit hole. Photo courtesy of Des Callaghan.

Figure 39. Scopelophila cataractae protonema and gemma. Photo by Des Callaghan, with permission.

Figure 40. Ditrichum flexicaule in Norway, a species that increased when rabbits declined. Photo by Michael Lüth, with permission.

Chapter 17-3: Rodents and Bats – Non-Muroidea

Figure 41. Pseudoscleropodium purum, a species that increased when rabbits declined. Photo by Janice Glime.

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Figure 44. Eurhynchium praelongum, a moss that increased in response to rabbit population decline. Photo by Michael Lüth, with permission.

Figure 42. Rhytidiadelphus squarrosus, a species that increased when rabbits declined. Photo by Jan-Peter Frahm, with permission.

Figure 45. Plagiomnium undulatum, a moss that increased in response to rabbit population decline. Photo by Michael Lüth, with permission.

Figure 43. Rhytidiadelphus triquetrus, a species that increased when the rabbit population declined. Photo courtesy of Eric Schneider.

Figure 46. Bryum caespiticium, in a moss genus that declined when rabbit population declined. Photo by Bob Klips, with permission.

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Figure 47. Climacium dendroides, a moss that declined when rabbit populations declined. Photo by Janice Glime.

Figure 48. Syntrichia ruralis ssp ruralis, a moss that declined when rabbit populations declined. Photo by Barry Stewart, with permission.

grazed area. These are all small and 10 of the 11 are acrocarpous. As in the Ranwell (1960) study, Watt found that mosses in the ungrazed turf are tall and mostly pleurocarpous. The small mosses seem to be unable to survive competition with taller vegetation, including competition for light. The larger mosses, on the other hand, seem to thrive in the ungrazed conditions. Watt considered these results to support the hypothesis that "in the grazed community the competitive power of the potentially taller growing plants is reduced by grazing sufficiently to allow the smaller species to survive and that in the ungrazed the unchecked growth of taller growing species eliminates or tends to eliminate the smaller, whether they are annual or perennial of varied life-forms." Gillham (1955) also stressed the importance of rabbit grazing, considering it to be less important than exposure. This contention was supported by the abundance of mosses that are intolerant of extreme exposure, but that are able to reach their maximum in the "closely nibbled swards." Heavy grazing caused moss cover to reach 25%, mostly of the moss Ceratodon purpureus (Figure 34) – a moss that is not shy of sunlight. In early spring, when the rabbits were most hungry, the lanes between the grazed heather bushes were dominated by the mosses Rhytidiadelphus squarrosus (Figure 42) and Hypnum cupressiforme (Figure 17). Gillham (1954) found that bryophyte fragments were only occasionally present in the rabbit dung and concluded that they were probably only eaten when mixed with other plant material. Although the bryophytes are important components of the turf in heavily grazed inland areas, they have little importance on sea cliffs due to their exposure to wind and salt there (Gillham 1955). Ochotonidae – Pikas Ochotona princeps – American Pika The American pika (Ochotona princeps; Figure 50) is distributed widely in British Columbia and the western USA (Defenders of Wildlife 2017). Mosses are often a dominant feature of their landscape.

Figure 49. Rhodobryum roseum, a species that disappears when rabbit herbivory declines. Photo by Hermann Schachner, through Creative Commons.

The results of Ranwell (1960) differ somewhat from those of Watt (1957), who showed that disappearance of rabbits resulted in the decrease of mosses in ungrazed pasture over long periods of time. Watt found 29 bryophyte species, but Rhytidiadelphus squarrosus (Figure 42) is found only in the ungrazed community. This is in contrast to its common presence in grazed pasture on the South Downs and other locations in Breckland, England. On the other hand, 11 species occur exclusively in the

Figure 50. Ochotona princeps among mosses. courtesy of Mallory Lambert, through Johanna Varner.

Photo

The presence of pikas is usually a good indicator of regions with rocky, mesic, cool habitat (Figure 51) with long winters and short summers (Simpson 2009). Although

Chapter 17-3: Rodents and Bats – Non-Muroidea

the American pikas (Ochotona princeps; Figure 51) are a high elevation species in western North America, in the Columbia River Gorge they live near sea level (Horsfall 1925; Varner & Dearing 2014a, b). But at low elevations in the southern part of the Columbia River Gorge, Oregon, USA, the known temperature range was extended and the long winters and typical snow accumulation were not present.

Figure 51. Ochotona princeps among the rocks and mosses of a talus slope. Photo courtesy of Johanna Varner.

Dr. Erik Beever (pers. comm.), research ecologist for the National Park Service Inventory & Monitoring program, reported to me that pikas occur at low elevations (less than 150 m) in a valley fed by a snowmelt river in the Cascade Range of western USA. The valley is cold, and he theorizes that their ability to survive the winter without their usual snow cover is due to the thick (>20 cm) moss mats that provide cover and insulation for them (Figure 52).

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pikas are able to travel long distances beneath the thick moss cover. Even their extensive moss consumption only removes about 0.002% of the moss in their home ranges in one year. Hence, unlike the lemmings, the pikas can enjoy the cover of the mosses without the danger of eliminating it. In this unusual habitat they subsist on what is for most rodents an unusual food – mosses (Varner & Dearing 2014a, b). These mosses comprise more than 60% of the diet at the two sites studied. At this rate, the pikas consume ~7.31 g/day and 2.67 kg/year of mosses. The mosses are available all year, thus making food caches unnecessary. Richardson (1981) considered mosses to be a difficult food for mammalian herbivores, having a high fiber content, low nitrogen, and low digestible energy compared to other food choices. Varner and Dearing (2014a) reported the same high fiber and low nitrogen (