Methods in Rhizosphere Biology Research 9789811357671, 9811357676

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Rhizosphere Biology

Didier Reinhardt Anil K. Sharma Editors

Methods in Rhizosphere Biology Research

Rhizosphere Biology Series Editor Anil K. Sharma Biological Sciences, CBSH, G.B.Pant University of Agriculture & Technology, Pantnagar, Uttarakhand, India

The Series Rhizosphere Biology, emphasizes on the different aspects of Rhizosphere. Major increase in agricultural productivity to meet growing food demands of human population is imperative, to survive in the future. Along with methods of crop improvement, an understanding of the rhizosphere biology, and the ways to manipulate it, could be an innovative strategy to deal with this demand of increasing productivity. This Series would provide comprehensive information for researchers, and encompass all aspects in field of rhizosphere biology. It would comprise of topics ranging from the classical studies to the most advanced application being done in the field. Rhizoshpere is a dynamic environment, and a series of processes take place to create a congenial environment for plant to grow and survive. There are factors which might hamper the growth of plants, resulting in productivity loss, but, the mechanisms are not very clear. Understanding the rhizosphere is needed, in order to create opportunities for researchers to come up with robust strategies to exploit the rhizosphere for sustainable agriculture. There are titles already available in the market in the broad area of rhizosphere biology, but there is a major lack of information as to the functions and future applications of this field. These titles have not given all the up-to-date information required by the today’s researchers and therefore, this Series aims to fill out those gaps.

More information about this series at http://www.springer.com/series/15861

Didier Reinhardt • Anil K. Sharma Editors

Methods in Rhizosphere Biology Research

Editors Didier Reinhardt Department of Biology University of Fribourg Fribourg, Switzerland

Anil K. Sharma Biological Sciences, CBSH G.B.Pant University of Agriculture & Technology Pantnagar, Uttarakhand, India

ISSN 2523-8442 ISSN 2523-8450 (electronic) Rhizosphere Biology ISBN 978-981-13-5766-4 ISBN 978-981-13-5767-1 (eBook) https://doi.org/10.1007/978-981-13-5767-1 Library of Congress Control Number: 2019934376 © Springer Nature Singapore Pte Ltd. 2019 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, express or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Springer imprint is published by the registered company Springer Nature Singapore Pte Ltd. The registered company address is: 152 Beach Road, #21-01/04 Gateway East, Singapore 189721, Singapore

Preface

The thin layer surrounding the roots, the rhizosphere, is one of the most important biological niches impacting on plant health and productivity, soil fertility and soil structure, and biodiversity belowground as well as aboveground. Hence, the rhizosphere is of central importance for natural ecosystems, from tropical forests to temperate environments, and even for arctic regions. In addition, the rhizosphere impacts on crop health and productivity and thereby influences crop yield and quality. Many aspects of the rhizosphere have long remained a black box, due to its inaccessibility and its complexity. However, in recent years, many aspects of the important functions of the rhizosphere in these different contexts have become increasingly acknowledged. New-generation sequencing techniques have unveiled one aspect of the rhizosphere: its astounding richness and the diversity of its inhabitants. Additional innovations include sophisticated compartment systems to assess transport phenomena and exchange of resources. Other important innovations are cutting-edge imaging techniques that allow to visualize processes in living tissues, to unveil subcellular structures at the sub-micrometer scale, and to assess nutrient element distributions in fixed material. This book intends to provide an overview on some of the most important techniques and methods applied to the rhizosphere to study processes and interactions in this unique biological compartment. Fribourg, Switzerland Pantnagar, Uttarakhand, India

Didier Reinhardt Anil K. Sharma

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Contents

Part I 1

2

3

Root Symbioses

Synthetic Plasmids to Challenge Symbiotic Nitrogen Fixation Between Rhizobia and Legumes . . . . . . . . . . . . . . . . . . . . . . . . . . . Jovelyn Unay and Xavier Perret In Vivo Analysis of Rhizosphere Enzyme Activities by the Use of Plastic Syringes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ru-Jie Li, Yi-Han Wang, Jie Cai, Jie Liu, Zhi-Ping Xie, and Christian Staehelin Characterization of Arbuscular Mycorrhizal Communities in Roots of Vineyard Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Alice Drain, Laurent Bonneau, Ghislaine Recorbet, Diederik van Tuinen, Daniel Wipf, and Pierre-Emmanuel Courty

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Molecular Methods for Research on Actinorhiza . . . . . . . . . . . . . . Hassen Gherbi, Valérie Hocher, Mariama Ngom, Nathalie Diagne, Joëlle Fournier, Alyssa Carre-Mlouka, Luis G. Wall, Louis S. Tisa, and Sergio Svistoonoff

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Molecular and Functional Characterization of Beneficial Bacteria Associated with AMF Spores . . . . . . . . . . . . . Monica Agnolucci, Alessandra Turrini, and Manuela Giovannetti

Part II

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27

35

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Plant Pathogens and Microbial Plant Protection

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Oomycete-Root Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jacob Hargreaves and Pieter van West

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Exploitation of Rhizosphere Microbiome Services . . . . . . . . . . . . . . 105 Valentina Riva, Elisa Terzaghi, Lorenzo Vergani, Francesca Mapelli, Elisabetta Zanardini, Cristiana Morosini, Giuseppe Raspa, Antonio Di Guardo, and Sara Borin

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Methods for Detecting Biocontrol and Plant Growth-Promoting Traits in Rhizobacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 133 Gustavo Santoyo, Juan M. Sánchez-Yáñez, and Sergio de los Santos-Villalobos

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A Split-Root Method to Study Systemic and Heritable Traits Induced by Trichoderma in Tomato Plants . . . . . . . . . . . . . . . . . . . . . . . . . . 151 M. B. Rubio, H. A. de Medeiros, M. E. Morán-Diez, P. Castillo, R. Hermosa, and E. Monte

Part III

Experimental Approaches and Analytical Techniques in Rhizosphere Biology

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Using a Tri-Isotope (13C, 15N, 33P) Labelling Method to Quantify Rhizodeposition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 169 Pierre Stevenel, E. Frossard, S. Abiven, I. M. Rao, F. Tamburini, and A. Oberson

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Microscopic Techniques Coupled to Molecular and Genetic Approaches to Highlight Cell-Type Specific Differences in Mycorrhizal Symbiosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 197 Valentina Fiorilli, Veronica Volpe, and Raffaella Balestrini

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From Imaging to Functional Traits in Interactions Between Roots and Microbes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 227 Yukari Kuga, Klaus Schläppi, and Didier Reinhardt

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Live Imaging of Arbuscular Mycorrhizal Symbiosis . . . . . . . . . . . . 241 Yoshihiro Kobae

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Microcosm Approaches to Investigate Multitrophic Interactions between Microbial Communities in the Rhizosphere of Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 255 Michael Bonkowski

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Analysis of Common Mycorrhizal Networks in Microcosms . . . . . . 271 Laurent Bonneau, Ghislaine Recorbet, Diederik van Tuinen, Daniel Wipf, and Pierre-Emmanuel Courty

About the Editors

Prof. D. Reinhardt is a researcher at the University of Fribourg, Switzerland. After completing his doctoral studies in the field of plant–pathogen interactions at the University of Basel, and a postdoc stay at the Salk Institute in La Jolla (California), he worked at the University of Bern. His main area of interest encompasses various aspects of arbuscular mycorrhizal symbiosis, with particular emphasis on the genetic factors in the host required for the intracellular accommodation of the fungal symbiont, and on the regulation of AM by exogenous factors such as nutrient availability. In addition, he is currently investigating organ formation and regulation of the body plan of plants. This includes the positioning of leaves and flowers around the growth axis, a phenomenon known as phyllotaxis. He has published over 50 research and review articles in various scientific journals. Prof. Anil Kumar Sharma is a professor at the Department of Biological Sciences, CBSH G.B. Pant University of Agriculture and Technology in Pantnagar. He completed his postdoctoral studies at the GSU, Louisiana (USA), and has been a visiting scientist at the University of Basel, Switzerland (2003) and University of Helsinki, Finland (2013). He has extensive research and teaching experience, and served as a reviewer for various funding bodies and international research journals. He also holds two patents in plant biology and microbiology, respectively. He is a recipient of various prestigious grants. He has published more than 60 research articles and 30 review articles, as well as 2 books with prominent publishers.

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Part I Root Symbioses

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Synthetic Plasmids to Challenge Symbiotic Nitrogen Fixation Between Rhizobia and Legumes Jovelyn Unay and Xavier Perret

Abstract

Growth of most angiosperms, including cereal crops, is constrained by a limited accessibility of nitrogen in soils. Until now, agriculture relied extensively upon chemical fertilizers to compensate for NPK deficiencies in fields, often at considerable ecological costs. Making better use of plant-bacteria associations could lower the environmental impact of agriculture while retaining good yields. For example, soil bacteria known as rhizobia reduce enough atmospheric nitrogen (N2) to secure growth and seed production of legumes. The positive impact on soil fertility of growing legumes in fields and pastures has been known for centuries. Yet, molecular mechanisms governing rhizobia-legume symbioses were only extensively deciphered recently. Conversion of N2 into ammonia by rhizobia nitrogenase occurs almost exclusively inside plant cells of specialized root (or more rarely stem) organs called nodules. Thus, rhizobia established on root surfaces must infect plant tissues and gain access to the cytoplasm of nodule cells before becoming proficient symbionts. Concomitantly, host plants must also prevent systemic infections by non-symbiotic bacteria while securing the development of the nodule organs. Bacterial infection and nodule development are coordinated by molecular signals exchanged by legumes and rhizobia. Amongst these signals, plant-made flavonoids and rhizobial nodulation (Nod) factors are instrumental in securing harmonious symbiosis. As many symbiotic signals and cognate receptors/sensors have been identified in recent years, genetic engineering offers new opportunities to study but also to harness the benefits of legumerhizobia symbioses. Here, we review the major molecular mechanisms involved in the development of proficient nodules and describe a framework for assembling a subset of symbiotic loci into small synthetic plasmids capable of

J. Unay · X. Perret (*) Department of Botany and Plant Biology, Sciences III, University of Geneva, Geneva, Switzerland e-mail: [email protected] © Springer Nature Singapore Pte Ltd. 2019 D. Reinhardt, A. K. Sharma (eds.), Methods in Rhizosphere Biology Research, Rhizosphere Biology, https://doi.org/10.1007/978-981-13-5767-1_1

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converting soil bacteria into beneficial plant symbionts. As proof of concept, the nodulation phenotype conferred by the synthetic plasmid pMSym2 is detailed.

1.1

Introduction

In natural ecosystems, land plants rely extensively upon soil microbes to compensate for essential macronutrient deficiencies. Symbiotic associations with arbuscular mycorrhizal fungi (AMF) and nitrogen-fixing bacteria provide higher plants with, respectively, the phosphate and reduced forms of nitrogen needed for growth. By contrast, intensive agriculture relies upon large quantities of chemical fertilizers to foster crop development and achieve the yields needed to secure food production. Because of its considerable ecological impact, chemical fertilization is under debate, and alternative solutions are actively sought. Legumes have been cultivated since the dawn of agriculture because of both the protein-rich grains they produce and perhaps more importantly their positive effect on soil fertility. Indeed, the use of nitrogenfixing legumes in crop rotation systems has long been known to improve the nitrogen and organic contents as well as the structure of farming soils. For these beneficial characteristics, legume species are often referred to as “green manure”. A key feature of most legumes is their ability to recruit specialized soil bacteria – collectively known as rhizobia – that provide host plants with reduced forms of nitrogen. These nitrogen-fixing symbioses give legumes an important ecological advantage particularly in nitrate-poor soils where non-fixing plants cannot prosper (Jackson et al. 2008). Similarly, in legume fields, inoculation of seedlings with selected rhizobia strains can replace nitrogen fertilizers to secure normal plant growth and seed production (Brewin 2010; Gresshoff et al. 2015).

1.1.1

Formation of Effective Nodules on Legume Roots

Symbiotic associations between rhizobia and legumes come in many forms and shapes (Masson-Boivin et al. 2009; Sprent et al. 2017). Yet, efficient nitrogen fixation by rhizobia occurs almost exclusively within plant cells, where rhizobia encounter a cellular environment tuned to maximize the activity of the nitrogenase complex (Downie 2005). Together, plant cells infected with rhizobia form specialized root (or occasionally stem) organs called nodules. Major legume crops form either one or two nodule types that can be distinguished according to their ontogeny and development (Ferguson et al. 2010). Indeterminate nodules (IDN) originate with dividing inner cortical cells and possess at their distal extremity a persistent meristem. Mature IDN are elongated and characterized by a longitudinal gradient, with plant cells and rhizobia at distinct stages of differentiation. Instead, determinate nodules (DN) originate with external cortical cells, become spherical and differentiate in a more synchronous manner, as the nodule meristem is only

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transient. Regardless of the nodule type, rhizobia must first establish persistent intracellular colonies prior to differentiating into fully active nitrogen-fixing bacteroids (Kereszt et al. 2011). Inside IDN of plants of the galegoid clade (e.g. Medicago sativa, Trifolium spp., Vicia sativa), bacteroids generally undergo a series of profound cellular changes leading to terminally differentiated microorganisms that have lost their capacity to divide (Mergaert et al. 2006). Inside the IDN of non-galegoid species (e.g. Mimosa pudica and Leucaena leucocephala), however, the bacteroids are not necessarily terminally differentiated (Marchetti et al. 2011). By contrast, DN nodules of many legumes such as bean, cowpea and soybean contain bacteroids that differ little from free-living rhizobia and can colonize again the rhizosphere once the nodules senesce and disaggregate. Both, nodule type and levels of bacteroid differentiation, are controlled by the legume host. Nodule development is normally restricted to situations when roots of nitrogenstarved plants contact compatible rhizobia (Masson-Boivin et al. 2009; Andrews and Andrews 2017). The successive developmental and cellular processes involved in the making of nitrogen-fixing nodules can be assigned to four distinct and yet complementary steps: (I) colonization of the host rhizosphere by symbiotic rhizobia (whether inoculated or indigenous to soils), (II) development of morphologically mature nodules, (III) concomitant colonization of root and nodule tissues by rhizobia and eventually (IV) implementation of symbiotic nitrogen fixation. As specific sets of plant and bacterial genes are dedicated to steps II and IV of symbiosis, characterization of the genetic bases for nodulation and nitrogen fixation was facilitated. By contrast, what makes rhizobia competitive in the legume rhizosphere (step I) and/or facilitates the infection of root tissues (step III) differs between strains and often involves the recruitment of housekeeping functions as, for example, the synthesis of surface polysaccharides. As each of the infected nodule cells will eventually contain as many as thousands of bacteroids, a precise modulation of the host immune system allows rhizobial colonization but prevents a systemic infection of roots by rhizobia or by non-symbiotic or pathogenic bacteria that also populate soils (Gourion et al. 2015). To coordinate root nodule formation with legitimate infection of plant tissues, legumes and rhizobia sequentially exchange a number of molecular cues (Perret et al. 2000; Oldroyd and Downie 2008; Oldroyd et al. 2011; Masson-Boivin and Sachs 2017). Although exceptions to the rule do exist (Okazaki et al. 2016), initiation of nodulation on roots of legume crops (step II) generally involves two major categories of signal molecules. First, legume roots exude complex cocktails of biologically active compounds amongst which phenolic flavonoids are recognized by rhizobia as signals announcing the presence of a suitable host. Root exudates partly determine the specificity of symbiotic associations as each rhizobial strain responds to a specific set of flavonoids. Within rhizobia, compatible flavonoids activate the transcription of nodulation (nod, nol and noe) genes via transcriptional regulators of the LysR family known as NodD proteins (Mulligan and Long 1985). NodD proteins activate transcription from conserved nod box (NB) sequences found in the promoter of nod operons (Fisher and Long 1993). Most nod genes are involved in the synthesis of strain-specific lipo-chito-oligosaccharides known as

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nodulation (Nod) factors (NF). The basic NF structure consists of three to six units of N-acetyl-D-glucosamine with a fatty acyl chain at the non-reducing end of the NF molecule [for a review see (Perret et al. 2000)]. Synthesis of NF backbones always involves three major enzymes: an N-acetyl-glucosaminyltransferase encoded by nodC, a deacetylase (NodB) and the acyltransferase NodA that links the acyl chain to the acetyl-free C-2 carbon of the NF non-reducing end. It is the NF synthesized by rhizobia that initiates nodule formation on host roots (Demont-Caulet et al. 1999) and also that allows rhizobia to enter into legume roots (Relić et al. 1994b). Thus, rhizobia mutants incapable of synthesizing NF are nodulation deficient (Nod-) on legumes that require NF to initiate symbiosis. Depending on the strain, a number of chemical NF substitutions can be added leading to the secretion of diverse NF mixtures. Occasionally, the substituents attached to the oligosaccharide NF backbone were reported as determinants of legume-rhizobia specificity (Lewin et al. 1990; Dénarié et al. 1996), but not necessarily on all legume hosts. Like that of the canonical NodA, NodB and NodC enzymes, expression of the NF-modifying enzymes is also flavonoid- and NodD-dependent (Perret et al. 2000). Genetic analyses showed that specificity in symbiotic interactions also lies in the properties of specialized plant receptors that perceive NF and are required to initiate nodule ontogeny and infection by rhizobia (Fliegmann and Bono 2015; Kelly et al. 2017). On most seed legumes, rhizobia generally enter roots via infection threads that guide infecting rhizobia across several cortical cell layers and are under the strict developmental control of host plants (Gage 2004; Oldroyd and Downie 2008). Once infection threads reach the nodule primordium, rhizobia are released into the cytoplasm of nodule cells as infection droplets surrounded by a plasma membrane of plant origin (the peribacteroid membrane). At this stage, endocellular rhizobia normally differentiate into proficient bacteroids, which, together with the peribacteroid membranes that surround them, form cellular compartments called symbiosomes (Roth et al. 1988). Bacteroid differentiation leads to a metabolic switch that converts free-living rhizobia into N2-fixing organelles (Fix+ phenotype) that express the nif and fix genes required for nitrogenase assembly and functioning (Fischer 1994; Poole et al. 2018). While NodD represents the flavonoid-dependent master switch for nodulation genes, the σ54-dependent NifA regulator controls the expression of essential nif and fix loci. In return for ammonia, host plants provide rhizobia with the carbon and energy sources required to fuel symbiotic N2 fixation mostly in the form of dicarboxylic acids such as malate and succinate, amino acids and often homocitrate as well (Poole et al. 2018).

1.1.2

Genes of Rhizobia Modulating Nodule Formation and Invasion

In addition to the NF-biosynthetic genes, rhizobia carry sets of strain-specific loci that contribute to symbiosis and whose expression is also flavonoid-dependent but not necessarily NodD-dependent. For example, several rhizobia use components of protein secretion machineries and/or secreted proteins to interact with legume hosts

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[for reviews see (Fauvart and Michiels 2008; Deakin and Broughton 2009)]. Rhizobium leguminosarum bv. viciae secretes via a constitutively expressed type I secretion system the NodO protein whose transcription is NodD- and flavonoiddependent (Fauvart and Michiels 2008). Notably, a nodO mutant of Rhizobium sp. strain BR816 showed an impaired symbiosis on P. vulgaris but not on L. leucocephala (Vlassak et al. 1998). In other strains, the presence of inducing flavonoids activates type III secretion systems (T3SS) or type IV secretion systems (T4SS) [for reviews see (Fauvart and Michiels 2008; Deakin and Broughton 2009)]. T3SS can deliver cocktails of effector proteins into the cytoplasm of eukaryotic target cells, which are needed to attenuate immune responses. Initially identified as an important virulence factor of plant and animal pathogens, the T3SS can also contribute to mutualistic symbioses. The first T3SS active during nodulation was characterized in the promiscuous Sinorhizobium fredii strain NGR234 (Viprey et al. 1998). Since then, symbiotic T3SS were also found in numerous rhizobia (Deakin and Broughton 2009), including the ß-proteobacterium Cupriavidus taiwanensis (Saad et al. 2012). Instead of T3SS, other rhizobia such as Mesorhizobium loti strain R7A use T4SS to target plant cells with effector proteins (Hubber et al. 2004). The effects on nodulation of T3SS or T4SS and the nodulation outer proteins (Nop) they secrete are host-dependent (Staehelin and Krishnan 2015). On some plants Nop secretion facilitates symbiosis, while on other hosts’ functional T3SS or T4SS and/or specific Nop impair the formation of Fix+ nodules (Miwa and Okazaki 2017). On a few soybean cultivars, the T3SS of Bradyrhizobium elkanii strain USDA61 was even shown to supersede NF signalling and to allow nodule formation on plant mutants otherwise incapable of detecting NF (Okazaki et al. 2013). This indicated that effector proteins could circumvent the normal NF-dependent regulatory cascade that triggers nodule formation (Miwa and Okazaki 2017). The propagation of infection threads through several layers of root cells and the release into nodule cells of rhizobia contained within branched infection threads are processes that also require appropriate signal molecules. Because of the intimate contact between plant and rhizobial cells, it was proposed that T3SS activity influences infection thread development (Viprey et al. 1998; Deakin and Broughton 2009; Staehelin and Krishnan 2015). Other types of rhizobial (signal) molecules are also involved at this stage of the infection process, however. For example, mutants of Sinorhizobium meliloti strain 1021 that lack type I exopolysaccharides (EPS I, also known as succinoglycan) form empty nodules on roots of Medicago. Although EPS probably protect rhizobia from plant-related stresses that may occur within infection threads or inside nodules, a number of observations indicate that the primary symbiotic role of EPS is to avoid or suppress plant defence responses (Gibson et al. 2008). Surface polysaccharides other than EPS may also support the rhizobial infection process, as in Lotus japonicus where Mesorhizobium loti cyclic β-glucans reduced production of antimicrobial phytoalexins (D’Antuono et al. 2008). In other symbioses, rhizobia mutants with altered lipopolysaccharides (LPS) frequently formed non-functional nodules. LPS are complex macromolecules made of a hydrophobic lipid A membrane anchor linked to an oligosaccharide core, which can be further modified by the addition of a variable O-antigen polysaccharide (Raetz and

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Whitfield 2002). Often LPS from pathogenic bacteria stimulate immune responses. In the case of NGR234, flavonoids induced the formation of rhamnose-rich O antigens known as rhamnan, and mutants impaired in rhamnan synthesis formed nodules but failed to fix nitrogen with certain hosts (Fraysse et al. 2002; Reuhs et al. 2005; Broughton et al. 2006). In addition to protective functions, surface polysaccharides also have signalling roles as demonstrated by the cognate exopolysaccharide receptor 3 (EPR3) recently identified in L. japonicus ecotype Gifu (Kawaharada et al. 2015). Unlike the NF receptors NFR1 and NFR5 that are expressed even in the absence of a compatible rhizobia strain, EPR3 was synthesized only when strains producing appropriate NF were inoculated onto L. japonicus roots, thus allowing plants to screen for rhizobia with correct EPS signals (Kawaharada et al. 2015; Long 2015). Together, these results show how rhizobia and legumes use combinations of molecular «keys» to define the specificity of their associations. Some «keys» such as flavonoids and NF are almost ubiquitous and ensure the onset of the symbiotic process leading to the development of the nodules. In contrast, additional strain-specific nodulation keys (e.g. LPS, EPS, T3SS and/or Nop) and the associated cognate plant receptors (such as EPR3 or those detecting effectors) finetune the infection process that directs rhizobia towards the cytoplasm of nodule cells.

1.1.3

From Specific to Promiscuous Symbioses

Because many rhizobia-legume associations were initially thought to be specific, strains of rhizobia were soon divided into cross-inoculation groups that defined proficient combinations of symbionts (Fred et al. 1932). As explained above, many signal molecules contribute to define symbiotic specificity including plant flavonoids, NF structures, effector proteins secreted by T3SS or T4SS, bacterial surface components (either expressed constitutively or modified during symbiosis), etc. Yet, what makes a strain promiscuous or, by contrast, defines a narrow host range in rhizobia is still unclear. Azorhizobium caulinodans strain ORS571, which was isolated from stem nodules of Sesbania rostrata (Dreyfus and Dommergues 1981), is often considered as the strain with the most narrow host range, even though it makes nodules on Sesbania punctata and other legume species such as Phaseolus vulgaris and Leucaena leucocephala (Waelkens et al. 1995). By contrast, S. fredii strain NGR234 triggers nodule formation on roots of plants belonging to more than 110 legume genera and fixes nitrogen in association with >130 legumes that form either DN or IDN (Pueppke and Broughton 1999). With so many hosts, NGR234 has the broadest host range so far described. Strain NGR234 was isolated by MJ Trinick in Papua New Guinea in the early 1960s. With a generation time of 3.5 h, NGR234 was the only fast growing of 30 isolates from nodules of Lablab purpureus L. (previously Dolichos lablab L.) (Trinick 1980). In the following years, NGR234 became known under different denominations including WU508 (University of Western Australia), ANU240 (Australian National University) or MPIK3030 (Max-Planck-Institut Köln). In 1980, Trinick reported that in addition to L. purpureus L., NGR234 also formed

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N2-fixing nodules on 7 of 12 tropical legumes that were thought to preferentially associate with slow-growing rhizobia (Trinick and Galbraith 1980). Mobilization and hybridization experiments later confirmed that essential symbiotic genes of NGR234 were carried by a 536 kb plasmid called pNGR234a (Pankhurst et al. 1983; Broughton et al. 1984). As the heat-cured derivative strain ANU265 was cultivable but unable to nodulate siratro, pNGR234a was defined as essential for symbiosis but dispensable for free-living growth (Morrison et al. 1983). Sequence analysis of pNGR234a highlighted the mosaic nature of this replicon and provided some clues on the genetic basis for symbiotic promiscuity (Freiberg et al. 1997; Perret et al. 2000). For example, it explained the particularly broad spectrum of NF (>80 different structures) synthesized by NGR234 (Price et al. 1992, 1996; Freiberg et al. 1997) and showed that the activity of a T3SS was not necessarily restricted to pathogenic interactions (Freiberg et al. 1997; Viprey et al. 1998). Later, the complete genome sequence of NGR234 confirmed that most genes essential for symbiosis were carried by pNGR234a but that a number of chromosomal loci were required for a sustained infection of nodules (e.g. ndvB) or an efficient nitrogen fixation ( fixNOPQ) (Schmeisser et al. 2009). In NGR234, the flavonoid-dependent expression of known symbiotic loci is mediated by a cascade that includes NodD1, NodD2, SyrM2 and TtsI as transcription regulators (Kobayashi et al. 2004). In turn, these regulators promote the sequential expression of more than 75 genes found downstream of cognate binding sites: 19 nod boxes (NB1 to NB19) for NodD1 and/or NodD2, 1 SyrM box (SB2) and 10 out of 11 TtsI-binding boxes (TB1 to TB4 and TB6 to TB11) (Kobayashi et al. 2004; Marie et al. 2004). As a nodD1 mutant of NGR234 (strain NGRΩnodD1) was Nod- regardless of the host tested (Relić et al. 1993), NodD1 was deemed as the master switch to begin the symbiotic dialogue with legume hosts (Kobayashi et al. 2004). By contrast, TtsI and NodD2 regulate the expression of various nodulation functions including T3SS and Nop’s secretion (TtsI role) or IAA and rhamnan synthesis (NodD2) (Viprey et al. 1998; Marie et al. 2003, 2004; Kobayashi et al. 2004. Mutants in nodD2 (NGRΩnodD2) and ttsI (NGRΩy4xI) were Nod+ on all of the inoculated legumes (Fellay et al. 1998; Viprey et al. 1998). Symbiotic proficiencies of NGRΩnodD2 and NGRΩy4xI often differed significantly, however. For example, on V. unguiculata NGRΩy4xI was as effective as the parent strain (Viprey et al. 1998), whereas NGRΩnodD2 formed poorly infected and ineffective nodules (Fellay et al. 1998). These results confirmed the host-specific character of mutations in regulators downstream of NodD1.

1.1.4

Are Many Keys Responsible for “Much Harmony”?

Almost 20 years ago, Broughton et al. proposed a molecular key-lock system to explain host specificity in rhizobia-legumes symbioses (Broughton et al. 2000). Accordingly, it was proposed that NGR234 had a particularly broad host range because of the following reasons and corollaries: (i) NodD1 is activated by a large spectrum of plant inducers (Le Strange et al. 1990), hence making NGR234 capable of detecting many hosts; (ii) NGR234 synthesizes a cocktail of >80 NF, which

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allows the strain to trigger nodulation on numerous legume species; (iii) T3SS and associated Nop together with complex surface polysaccharides are additional «keys» to open the «locks» found inside infection threads, thus reinforcing the capacity of NGR234 to reach and infect nodules. In other words, a large key ring made NGR234 more promiscuous than other rhizobia equipped with fewer keys. A number of observations questioned this model, however. First, in spite of the many chemical substitutions found on NF of NGR234, only a nodS mutant (strain NGR234Ω25) exhibited a restricted host range compared to wild type and only on L. leucocephala (Lewin et al. 1990; Jabbouri et al. 1995). This suggested that noeK, noeJ, noeL, nolK, nodZ, noeI, noeE, nolL, nolO and nodU were dispensable for NGR234 nodulation. As a nodS mutant of NGR234 was found to secrete lower NF levels (Relić et al. 1994a), it was perhaps the quantity of NF rather than their chemical nature that made NGR234 promiscuous. Second, the comparison of the symbiotic properties of mutant strains in nodD2, ttsI, rhcN (in which the T3SS was inactivated) or either of the genes coding for Nop showed no clear relationship between a missing (mutated) “key” and a restriction in the mutant’s host range. In fact, it was concluded that the sum of Nop rather than individual effectors defined whether T3SS was beneficial or detrimental to symbiosis on a specific host (Deakin and Broughton 2009). Furthermore, as bacterial surface polysaccharides were reported to modulate plant defences, LPS may well counterbalance the effect of T3SS on some hosts. Given the limited number of successive mutations one can easily introduce in a bacterium, we searched for alternative ways to identify which of the many nodulation keys in NGR234 are essential and confer symbiotic promiscuity.

1.2

A Forward Genetic Approach to Study Nodulation Genes

The recent development of easy-to-use and reliable protocols for de novo synthesis and assembly of new replicons made genetic engineering more accessible (Gibson 2012; de Kok et al. 2014). In the field of rhizobia-legume symbioses, several ongoing initiatives attempt at manipulating plant and bacterial genomes, for example, to engineer nitrogen-fixing cereals (Mus et al. 2016), to assemble nitrogenase components in yeast mitochondria (Buren et al. 2017), to reduce S. meliloti genome to a minimal set of core genes (diCenzo et al. 2016) or to build libraries of shuttle vectors to engineer rhizobia genomes (Dohlemann et al. 2017). Although these research avenues may offer mid- to long-term solutions to the immediate problem of making agriculture more sustainable, they are unlikely to boost plant productivity in the near future. Instead, a more efficient use of rhizobial inoculants may rapidly improve legume and other crop yields. The poor survival and low competitiveness of rhizobia introduced into foreign soils is a well-known phenomenon (Streeter 1994), which often hinders the use of commercial inoculants, however. A possible solution to this problem would be to convert into N2-fixing symbionts some of the local bacteria that are perfectly adapted to existing soil conditions. In principle, this could be achieved by introducing into selected recipient bacteria a minimal set of nod, nif

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Fig. 1.1 Simplified scheme for assembling the series of mini-symbiotic plasmid pMSym. Left panel describes the genetic modules assembled together to create the synthetic replicons: the ca. 7.5 kb black Module 1 (pMSym1) carries all functions necessary for propagation and selection of constructs in E. coli, rhizobia and non-symbiotic bacteria; Addition of red Module 2 (yielding pMSym2) allows for synthesis of NF in response to compatible flavonoids and mediates nodule formation (Nod+ phenotype). To promote infection of root tissues and establishment of intracellular colonies within nodule cells (Inf+), blue Module 3 is added to make pMSym3. Introduction of yellow Module 4, which codes for nif and fix genes and the regulatory functions for symbiotic nitrogen fixation (Fix+), makes pMSym4. Right panel describes the workflow for synthesizing pMSym replicons: step 1, PCR-amplified gene blocks are assembled using Gibson protocol; step 2, assembly mixes are transformed into E. coli to obtain sufficient quantities of constructs to verify structures and DNA sequences; and step 3, conjugation secures transfer of synthetic replicons into selected hosts. The ANU265 derivative strain cured of the large pNGR234a symbiotic plasmid was selected as a host to optimize expression of NGR234 genes

and fix genes sufficient to entice the formation of proficient nodules on roots of as many legume crops as possible. In order to define this limited set of symbiotic genes, we chose a forward genetic approach that consists in introducing into a genomic platform lacking symbiotic functions various synthetic plasmids that carry selected loci of strain NGR234. The rational for choosing NGR234 as a gene reservoir stems from its unsurpassed ability to nodulate a broad set of seed legumes and fix nitrogen inside both DN and IDN of many legume species (Pueppke and Broughton 1999). Major characteristics of the synthetic replicons and steps towards assembling a complete symbiotic plasmid are detailed in Fig. 1.1. Selected gene blocks are amplified by PCR, independently cloned to verify or alter sequences if needed, and brought together using the Gibson assembly protocol (Gibson 2012). To facilitate engineering steps, assembled replicons are first maintained in Escherichia coli

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for sequence verification and then mobilized into recipient strains by conjugal transfer. Mini-symbiotic plasmids (pMSym) consist of four main modules. Module 1 includes functions required for maintenance and selection in E. coli and rhizobia as well as an origin transfer; Module 2 mediates nodule formation on legume roots; Module 3 carries genes to secure infection of host plants and Module 4 includes loci required for assembly and functioning of the nitrogenase enzyme. Each of the modules includes the regulatory functions needed to secure the timely expression of the selected gene blocks. Depending on the recipient strain inside which the synthetic symbiotic plasmid must function or the legume host that is targeted, a fifth module conferring additional regulatory or metabolic functions might be needed, however. Given that symbiotic genes of rhizobia are generally clustered into chromosomal islands [e.g. M. loti strain R7A (Sullivan et al. 2002)] or found on large symbiotic plasmids, synthetic pMSym must be maintained as low- to unit-copy number plasmids to minimize possible adverse effect on the metabolism of recipient bacteria. For phenotypic assessment, synthetic constructs are initially mobilized into ANU265. Using a derivative strain of NGR234 cured of its symbiotic plasmid as a genomic platform to test synthetic constructs ensures proper expression of the selected symbiotic functions.

1.2.1

pMSym2: A Small Replicon That Confers Nodulation to Recipient Bacteria

Synthetic construct pMSym2 includes genetic modules 1 and 2, whose sequences and gene organizations cannot be detailed here because of proprietary issues. Module 2 encodes a minimal set of nodulation genes of NGR234 required to synthesize and secrete a unique kind of NF in response to plant flavonoids. To test whether this construct was sufficient to induce nodule formation on legume roots, pMSym2 was mobilized into ANU265, and the resulting transconjugants were inoculated onto sterile seedlings of Vigna unguiculata grown in Magenta jars. Nodulation assays were conducted as previously described (Fumeaux et al. 2011), and plants were harvested at 7, 14, 21 and 28 days postinoculation (dpi) to follow the development of nodules (see Fig. 1.2). At 28 dpi, plants inoculated with NGR234 carried an average of 64 nodules (15.7), while ANU265::pMSym2 enticed the formation of ca. 25 (4.4) apparently normal DN structures. Whereas nodules formed by NGR234 were 2–3 mm in diameter, those enticed by ANU265:: pMSym2 were notably smaller (ca. 1–1.5 mm). Sections of nodules infected with NGR234 showed the presence of leghaemoglobin (Fig. 1.2c), indicative of active nitrogen fixation, even though the shoot dry weight of control and inoculated plants did not differ significantly yet. By contrast, sections of nodules induced by ANU265::pMSym2 showed signs of necrosis in the nodule centre where plant cells are expected to be infected with rhizobia (Fig. 1.2f). As expected, control plants inoculated with the ANU265 recipient strain or grown in the absence of bacteria showed no root outgrowth or nodule. Although numbers of nodules varied

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Fig. 1.2 Nodules of determinate type formed on Vigna unguiculata roots by wild-type S. fredii strain NGR234 (panels a to c) or by the ANU265 transconjugant carrying the synthetic pMSym2 mini-symbiotic plasmid (panels d to f). Nodules were photographed 7 (panels a and d), 14 (b and e) and 28 (c and f) days postinoculation (dpi). Photographs of 28 dpi nodule sections (panels c and f) highlight the presence of leghaemoglobin inside nodules formed by the nitrogen-fixing strain NGR234 (panel c), whereas those formed by the ANU265::pMSym2 strain showed necrotic regions (panel f). Scale bars correspond to 0.2 mm for panels a and d and 1 mm for panels b, c, e and f

slightly, repeated nodulation trials confirmed that the partial nodulation phenotype conferred by pMSym2 to ANU265 was both, stable and robust.

1.3

pMSym2: A Successful Step Towards Constructing Synthetic Symbiotic Replicons

Features embedded into Module 1 differ significantly from those of the repABC-type shuttle vectors (Dohlemann et al. 2017). Yet, pMSym derivatives propagate inside Proteobacteria such as E. coli and ANU265 at single- to low-copy number (data not shown) and with sufficient stability (even in the absence of antibiotic selection) to enable ANU265 recipients to initiate nodule formation once inoculated onto host roots. Nodulation properties of ANU265::pMSym2 on cowpea fits well the genetic data acquired over the three decades of studying the promiscuous NGR234 strain. The small nodulation Module 2 carried by pMSym2, which only includes one tenth of the genes from the complex NodD1-NodD2-SyrM2-TtsI regulon (Perret et al. 2003; Kobayashi et al. 2004), is sufficient for nodule formation on V. unguiculata. As cowpea was reported as a widely compatible legume (Lewin et al. 1987), thus

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probably receptive to a broad spectrum of NF, nodulation by pMSym2 transconjugants was not unexpected. Nodules appeared almost simultaneously with both strains, but those formed by ANU265::pMSym2 appeared to stop developing 2–3 weeks postinoculation. Concomitant to this phenomenon, dark-stained and apparently necrotic lesions were repeatedly observed in the central zone of nodules indicative of an immune response mounted against the pMSym2 transconjugants. Thus, additional nodulation functions (Module 3) must be added to synthetic constructs to allow infecting bacteria to colonize cowpea nodules and establish persistent colonies within nodule cells as efficiently as the parent NGR234 does. Unlike previous attempts to transfer entire large symbiotic plasmids to confer nodulation to recipient strains (Broughton et al. 1986), our work confirms that even a small set of selected nodulation genes assembled in a synthetic plasmid is sufficient to trigger the development of nodules on roots of cowpea. The symbiotic response was incomplete as necrotic lesions developed in the infection zone, indicating that cowpea may perceive infecting ANU265::pMSym2 cells as non-legitimate rhizobia. Nevertheless, nodules appeared more developed and structured than the smaller pseudo-nodules or nodule primordia often observed on host roots following the inoculation with poorly matched symbionts. These promising results confirm that it is possible to dissect the molecular bases for symbiotic nitrogen fixation into independent genetic modules, which in turn can be engineered to better suit our needs, and once assembled together, transform non-symbiotic bacteria into proficient rhizobia capable of fostering plant growth. Acknowledgements We would like to thank Natalia Giot for her help in many aspects of this work. Financial support for this project was provided by the University of Geneva and the Swiss National Science Foundation (grants no. 31003A-146,548 and 31003A-173,191).

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In Vivo Analysis of Rhizosphere Enzyme Activities by the Use of Plastic Syringes

2

Ru-Jie Li, Yi-Han Wang, Jie Cai, Jie Liu, Zhi-Ping Xie, and Christian Staehelin

Abstract

Analysis of enzyme activities and metabolites in the rhizosphere requires suitable growth systems. Here, we describe the use of plastic syringes as a test system for measuring extracellular root enzyme activities of a single seedling under sterile conditions. Syringes have been originally used to analyze hydrolysis of rhizobial nodulation signals (Nod factors) by host legumes such as Medicago truncatula. The developed method offers now the opportunity to use syringes for other purposes. Syringe systems can be used to determine any enzyme activity in the rhizosphere, providing that the substrates and products in the growth medium are water-soluble, nontoxic, and detectable. Moreover, syringes can be employed to collect root exudates from a single seedling.

2.1

Introduction

Plant roots form many interactions with soil microbes that may be either parasitic or mutualistic. The rhizosphere is the microecological zone where microbial root colonization and infection events take place (van Dam and Bouwmeester 2016).

R.-J. Li · Y.-H. Wang · J. Cai · J. Liu State Key Laboratory of Biocontrol and Guangdong Key Laboratory of Plant Resources, School of Life Sciences, Sun Yat-sen University, Guangzhou, China Z.-P. Xie (*) · C. Staehelin (*) State Key Laboratory of Biocontrol and Guangdong Key Laboratory of Plant Resources, School of Life Sciences, Sun Yat-sen University, Guangzhou, China Shenzhen Research and Development Center of State Key Laboratory of Biocontrol, School of Life Sciences, Sun Yat-sen University, Shenzhen, China e-mail: [email protected]; [email protected] © Springer Nature Singapore Pte Ltd. 2019 D. Reinhardt, A. K. Sharma (eds.), Methods in Rhizosphere Biology Research, Rhizosphere Biology, https://doi.org/10.1007/978-981-13-5767-1_2

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Analysis of enzyme activities and metabolites related to microbial infection requires suitable growth systems. Here, we describe the use of plastic syringes as a smallscale culture system to measure enzyme activities in the rhizosphere of individual young seedlings. In our laboratory, we are working on nodulation signals (Nod factors) of nitrogen-fixing rhizobia. These bacteria induce root nodules on host legumes such as Medicago truncatula. Nod factor synthesis is stimulated in response to host flavonoids in the rhizosphere. Nod factors are required for bacterial entry into host plants, as rhizobia mutated in Nod factor synthesis usually cannot infect host plants. Nod factors are lipo-oligosaccharides (modified chitin oligosaccharides) consisting of a sugar backbone and an N-linked fatty acid (acyl chain) at the nonreducing end (Perret et al. 2000). The sugar backbone of many Nod factors is chemically modified with additional groups, which vary in a strain-specific manner and can determine host specificity. Sinorhizobium meliloti produces Nod factors with a sulfate group at the reducing end (Lerouge et al. 1990; Schultze et al. 1992) (Fig. 2.1a). Nod factors of S. meliloti are perceived by Nod factor receptors of M. truncatula (Limpens et al. 2003; Ben Amor et al. 2003; Arrighi et al. 2006). Nod factor signaling in M. truncatula culminates in expression of symbiosis-related host genes. One of them is MtNFH1 (MEDICAGO TRUNCATULA NOD FACTOR HYDROLASE1). Nod factor hydrolysis by the MtNFH1 enzyme plays a role in the fine-tuning of the symbiosis at the stage of rhizobial root hair infection and in nitrogen-fixing nodules. Remarkably, MtNFH1-deficient mutant plants formed enlarged nodules that showed abnormal nodule branching (Cai et al. 2018). MtNFH1 efficiently cleaves Nod factors produced by S. meliloti (Tian et al. 2013). Hydrolysis of Nod factors by MtNFH1 results in formation of lipo-disaccharides that can be separated from the non-hydrolyzed Nod factors by high-pressure liquid chromatography (HPLC) as shown in Fig. 2.1a (Staehelin et al. 1994). MtNFH1 is an extracellular enzyme and rapidly degrades purified Nod factors of S. meliloti applied to the M. truncatula rhizosphere (Tian et al. 2013). Pretreatment of M. truncatula roots with low concentrations of S. meliloti Nod factors results in increased MtNFH1 expression and consequently in increased Nod factor hydrolysis (Cai et al. 2018). Similar Nod factor cleaving activities were previously characterized in the rhizosphere of Medicago sativa, Vicia sativa, and Pisum sativum (Heidstra et al. 1994; Staehelin et al. 1994, 1995; Ovtsyna et al. 2000, 2005). In the course of these studies, it turned out that plastic syringes represent a convenient test system for measuring rhizospheric Nod factor hydrolysis in vivo. Here, we illustrate the use of syringes to analyze Nod factor hydrolysis in the rhizosphere of Medicago sp. (Staehelin et al. 1994, 1995; Tian et al. 2013; Cai et al. 2018). Syringes are also suitable to determine other enzyme activities in the rhizosphere and to prepare root exudates from a single seedling. We hope that this chapter stimulates researchers to extend the described syringe method to other plants.

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Fig. 2.1 Illustration of the described method. (a) Analysis of Nod factors and cleavage products by reverse-phase HPLC (Nova Pak C18, 3.9  150 mm from waters, 35% (v/v) acetonitrile/water containing 40 mM ammonium acetate as the mobile phase). The chromatogram shows the tetrameric Nod factor NodSm-IV(C16:2, S) and the lipo-disaccharidic cleavage product NodSm-II (C16:2) separated into anomers (double peak). (b) Preparation of M. truncatula seedlings. (c) Transfer of a M. truncatula seedling onto the top of a 1 mL syringe filled with Jensen medium and NodSm-IV(C16:2, S). Foam rubber was used to keep the syringe in a vertical position. (d) A syringe containing a transferred seedling. (e, f) Seedlings covered by plastic caps (1 mL pipette tips sealed on a flame). (g) Analysis of β-N-acetylglucosaminidase activity in the M. truncatula rhizosphere using p-nitrophenyl N-acetyl-β-D-glucosaminide as substrate

2.2

Description of the Method and Discussion

Seeds of M. truncatula were surface-sterilized with diluted sodium hypochlorite for 20 min on a shaker. The seeds were then washed with sterilized water three times and transferred to 0.8% (w/v) agar plates. The plates were incubated in the dark at 4  C

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for 48 h. For germination, the plates were placed upside down (agar with attached seeds on the top) and incubated in the dark at 20  C for ca. 16–20 h. In this way, the germinated seedlings formed uniform and straight primary roots (Fig. 2.1b). Nod factors of S. meliloti were purified from culture supernatants of S. meliloti strain 1021 (pEK327) (Schultze et al. 1992) as described (Staehelin et al. 1994; Tian et al. 2013). Nod factors, such as NodSm-IV(C16:2, S) (Fig. 2.1a), were purified by HPLC and dried. The Nod factors were then dissolved in small quantities of dimethylsulfoxide (DMSO), and defined quantities were added to sterilized deposit-free Jensen medium (Van Brussel et al. 1982) containing 0.5% (v/v) DMSO (final concentration). Sterile 1 mL plastic syringes filled with solution (usually 300 μL) were placed into foam rubber (with appropriate holes) to keep the syringes in a vertical position. The germinated seedlings with short roots (ca. 0.5 cm in length) were then carefully transferred from the agar plate to the syringes with the help of sterile forceps (Fig. 2.1c and d). The syringes were then covered by plastic caps (1 mL pipette tips sealed on a flame) to protect the seedlings and reduce evaporation (Fig. 2.1e and f). After incubation (e.g., 24  C for 18 h), caps and seedlings were removed. Nod factors and acylated cleavage products (lipodisaccharides) in the incubation medium were extracted with an equal volume of nbutanol, dried and analyzed by reverse-phase HPLC as described (Staehelin et al. 1994). A representative HPLC chromatogram is shown in Fig. 2.1a. A substrate concentration of 15 μM NodSm-IV(C16:2, S) in 400 μL Jensen medium was required to analyze Nod factor hydrolysis in the rhizosphere of a single seedling. Alternatively, a concentration of 5 μM NodSm-IV(C16:2, S) in 300 μL Jensen medium was used, and solutions from three plants were combined to obtain sufficient amounts of Nod factors and cleavage products for HPLC analysis. Only low quantities of Nod factors or cleavage products could be extracted from incubated roots after washing with n-butanol. Hence, analysis of the incubation medium was sufficient to measure Nod factor hydrolysis in the rhizosphere. Instead of Nod factor hydrolysis, other rhizospheric enzyme activities can be measured with the established syringe system. For example, we recently used the substrate p-nitrophenyl N-acetyl-β-D-glucosaminide (Sigma) to determine β-Nacetylglucosaminidase activity in the rhizosphere of M. truncatula (Fig. 2.1g). Formed p-nitrophenol (yellow coloration) in the incubation medium was quantified with a spectrophotometer. The design of experiments with syringes can be varied in different ways as shown in Fig. 2.2. For example, seedlings incubated in a given substrate or inoculated with rhizobia can be used for further purposes such as nodulation tests or seed propagation. Alternatively, RNA from seedlings incubated with a given substrate can be analyzed, and expression of a given gene can be compared with the corresponding enzyme activity in the rhizosphere (Fig. 2.2a). Furthermore, the syringe system can be used to analyze the effects of various pretreatments such as application of low concentrations of Nod factors, inoculation with bacteria, or application of chemicals that promote or inhibit nodulation signaling. Different root exudates can be prepared,

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Fig. 2.2 Design of possible enzyme assays with syringes. (a) Analysis of Nod factor hydrolysis (or other activities) in a single step. Removed seedlings are used for further purposes. Substrate and cleavage products accumulated in the incubation medium are measured. (b) Analysis of pretreatment effects on enzyme activities in root exudates. Seedlings are incubated with or without the test compound at a defined concentration. The enzyme assay is performed in the absence of the seedlings. (c) Analysis of pretreatment effects on enzyme activities in the rhizosphere. Seedlings are pretreated in syringes with a given test compound and then transferred to syringes containing the substrate. The enzyme assay is performed in the presence of seedlings. To vary the pretreatment time, seedlings are first incubated in additional syringes under control conditions and then transferred at different times to syringes containing the test compound

and activities in the incubation medium are then assayed by addition of a given substrate (Fig. 2.2b). Such root exudation experiments can provide information on the properties of a given enzyme, i.e., whether it is secreted and active in the incubation medium. Finally, effects of different pretreatments can be examined for enzyme activities that are associated with the root surface. In this case, plants are pretreated in a first syringe and then transferred to a second syringe with a given substrate (Fig. 2.2c). Using such an experimental design, M. sativa (Staehelin et al. 1995) and M. truncatula (Cai et al. 2018) showed increased Nod factor hydrolysis when the seedlings were pretreated with S. meliloti Nod factors for at least 2 h. In our recent study on MtNFH1, we found a good correlation between MtNFH1 expression and Nod factor hydrolysis in the rhizosphere. A pretreatment of seedlings with 10 nM NodSm-IV(C16:2, S) was sufficient to reach half-maximal induction of MtNFH1 activity (Cai et al. 2018).

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Conclusions and Outlook

Syringes represent a convenient test system for measuring enzyme activities in the rhizosphere, providing that the substrates and products in the growth medium are water-soluble, nontoxic, and detectable. In this system, the root is not in contact with the plastic wall of the syringe, and touch-induced stress can be avoided. The level of the solution in the syringe can be adjusted by the plunger in order to submerge the root but not the hypocotyl in the solution. Activities of seedlings can be determined under sterile conditions. The small incubation volume (200–400 μL) allows analysis of small quantities of a given molecule. Syringe systems are therefore particularly suitable when commercial substrates are expensive, and preparation of laboratoryprepared substrates is time-consuming. Another advantage of the syringe system is that a single plant can be analyzed. Each syringe can be considered as an independent sample, thus allowing statistical analysis of many biological replicates, or phenotyping of individual seedlings in a segregating population. Syringes can be used in different ways. Enzyme activities secreted into the incubation medium or associated with the root surface can be determined. Furthermore, syringes are suitable to compare pretreatment effects. Finally, syringes can be employed for preparation of root exudates, while removed seedlings can be used for other purposes such as gene expression analysis. A challenge for the future will be to adopt the described method to other plants. P. sativum, for example, possesses bigger seeds than Medicago, and it was necessary in previous studies to prepare a suitable syringe type by cutting off the tip of the syringe (Ovtsyna et al. 2000, 2005). Another future modification of the syringe system would be to fill the syringes with soil substrate in order to measure enzyme activities and metabolites under natural conditions. Acknowledgments We acknowledge financial support from the National Natural Science Foundation of China (grant 31670241), from the Department of Science and Technology of Guangdong Province, China (grant 2016A030313299), from the Science Foundation of the State Key Laboratory of Biocontrol (grants SKLBC 16A01 and SKLBC 322017A09), and from the Guangdong Key Laboratory of Plant Resources (grant 2014B030301026).

References Arrighi JF, Barre A, Ben Amor B, Bersoult A, Soriano LC, Mirabella R, De Carvalho-Niebel F, Journet EP, Ghérardi M, Huguet T, Geurts R, Dénarié J, Rougé P, Gough C (2006) The Medicago truncatula lysin motif-receptor-like kinase gene family includes NFP and new nodule-expressed genes. Plant Physiol 142:265–279 Ben Amor B, Shaw SL, Oldroyd GE, Maillet F, Penmetsa RV, Cook D, Long SR, Dénarié J, Gough C (2003) The NFP locus of Medicago truncatula controls an early step of Nod factor signal transduction upstream of a rapid calcium flux and root hair deformation. Plant J 34:495–506 Cai J, Zhang LY, Liu W, Tian Y, Xiong JS, Wang YH, Li RJ, Li HM, Wen J, Mysore KS, Boller T, Xie ZP, Staehelin C (2018) Role of the Nod factor hydrolase MtNFH1 in regulating Nod factor levels during rhizobial infection and in mature nodules of Medicago truncatula. Plant Cell 30:397–414

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Heidstra R, Geurts R, Franssen H, Spaink HP, Van Kammen AB, Bisseling T (1994) Root hair deformation activity of nodulation factors and their fate on Vicia sativa. Plant Physiol 105:787–797 Lerouge P, Roche P, Faucher C, Maillet F, Truchet G, Promé JC, Dénarié J (1990) Symbiotic hostspecificity of Rhizobium meliloti is determined by a sulphated and acylated glucosamine oligosaccharide signal. Nature 344:781–784 Limpens E, Franken C, Smit P, Willemse J, Bisseling T, Geurts R (2003) LysM domain receptor kinases regulating rhizobial Nod factor-induced infection. Science 302:630–633 Ovtsyna AO, Schultze M, Tikhonovich IA, Spaink HP, Kondorosi É, Kondorosi Á, Staehelin C (2000) Nod factors of Rhizobium leguminosarum bv. viciae and their fucosylated derivatives stimulate a Nod factor cleaving activity in pea roots and are hydrolyzed in vitro by plant chitinases at different rates. Mol Plant-Microbe Interact 13:799–807 Ovtsyna AO, Dolgikh EA, Kilanova AS, Tsyganov VE, Borisov AY, Tikhonovich JA, Staehelin C (2005) Nod factors induce Nod factor cleaving enzymes in pea roots. Genetic and pharmacological approaches indicate different activation mechanisms. Plant Physiol 139:1051–1064 Perret X, Staehelin C, Broughton WJ (2000) Molecular basis of symbiotic promiscuity. Microbiol Mol Biol Rev 64:180–201 Schultze M, Quiclet-Sire B, Kondorosi É, Virelizer H, Glushka JN, Endre G, Géro SD, Kondorosi Á (1992) Rhizobium meliloti produces a family of sulphated lipooligosaccharides exhibiting different degrees of plant host specificity. Proc Natl Acad Sci U S A 89:192–196 Staehelin C, Schultze M, Kondorosi É, Mellor RB, Boller T, Kondorosi Á (1994) Structural modifications in Rhizobium meliloti Nod factors influence their stability against hydrolysis by root chitinases. Plant J 5:319–330 Staehelin C, Schultze M, Kondorosi É, Kondorosi Á (1995) Lipo-chitooligosaccharide nodulation signals from Rhizobium meliloti induce their rapid degradation by the host plant alfalfa. Plant Physiol 108:1607–1614 Tian Y, Liu W, Cai J, Zhang LY, Wong KB, Feddermann N, Boller T, Xie ZP, Staehelin C (2013) The nodulation factor hydrolase of Medicago truncatula: characterization of an enzyme specifically cleaving rhizobial nodulation signals. Plant Physiol 163:1179–1190 Van Brussel AAN, Tak T, Wetselaar A, Pees E, Wijffelman CA (1982) Small Leguminosae as test plants for nodulation of Rhizobium leguminosarum and other rhizobia and agrobacteria harbouring a leguminosarum sym-plasmid. Plant Sci Lett 27:317–325 van Dam NM, Bouwmeester HJ (2016) Metabolomics in the rhizosphere: tapping into belowground chemical communication. Trends Plant Sci 21:256–265

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Characterization of Arbuscular Mycorrhizal Communities in Roots of Vineyard Plants Alice Drain, Laurent Bonneau, Ghislaine Recorbet, Diederik van Tuinen, Daniel Wipf, and Pierre-Emmanuel Courty

Abstract

We developed a new protocol to study arbuscular mycorrhizal fungal (AMF) communities in Bordeaux vineyards in a standardized way, in order to compare results obtained over years and between locations. To this end, we first used grapevine root samples instead of soil samples to avoid AMF spores or hyphae interacting with cover plants. We next increased the number of grapevine samples to obtain more representative coverage of AMF communities while decreasing variability intra-vineyard, especially for the larger parcels. In addition, we adapted the DNA extraction protocol dedicated to soil samples to grapevine roots, as a way to increase the yield and the purity of samples. These features, coupled to the choice of the LSU-D2 region as the molecular marker for highthroughput sequencing (MiSeq® technology), further allowed us to assess the AMF populations of Bordeaux vineyards.

3.1

Introduction

Arbuscular mycorrhiza (AM) is an ancestral symbiosis established between soilborne fungi of the subphylum Glomeromycotina (Spatafora et al. 2016) and up to 80% of land plants, in which both partners obtain advantages. The plant allocates a part of carbohydrates originating from photosynthesis to the arbuscular mycorrhizal fungi (AMF) that are obligate symbionts. In return, the fungal partner transfers to the host plant mineral nutrients and water scavenged from the soil through the development of an extensive extra-radical hyphal network (Smith and Read 2008). In this manner, AMF give plants access to nutrients located beyond the depletion zone that A. Drain · L. Bonneau · G. Recorbet · D. van Tuinen · D. Wipf · P.-E. Courty (*) Agroécologie, AgroSup Dijon, CNRS, INRA, Université Bourgogne Franche-Comté, Dijon, France e-mail: [email protected]; [email protected] © Springer Nature Singapore Pte Ltd. 2019 D. Reinhardt, A. K. Sharma (eds.), Methods in Rhizosphere Biology Research, Rhizosphere Biology, https://doi.org/10.1007/978-981-13-5767-1_3

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develops around the roots and thus support plant development. Moreover, AM colonization is known to stimulate plant immunity, acting like a “vaccine” that can prime systemic defences against potential pathogen attacks (Cameron et al. 2013). In addition to these beneficial effects on plant physiology and health, AM can also improve soil stability and prevent soil erosion (Rillig et al. 2002). Through all these features, AM fungi appear as good candidates in a sustainable agriculture context to limit chemical fertilization and pesticide application in fields, together with supplying essential protection against many biotic and abiotic stresses (Gianinazzi et al. 2010). To date, very few AMF strains have been commercialized, and most of experimental data relate to the use of the ubiquitous strain DAOM197198 (Rhizophagus irregularis). Consequently, to obtain the best advantages of AMF for a greener agriculture, we need to improve our knowledge on Glomeromycotina species contribution to ecosystem services and interactions between different strains, host plants and other soil microorganisms (Basu et al. 2018). The global comprehension of AMF biology and ecology is a prerequisite to improve the selection and the use of efficient and locally adapted strains. In France, Vitis vinifera cultures represent a major economical and ecological challenge. Indeed, many winegrowers have started to use natural practices to improve land productivity while decreasing agricultural inputs. As AMF emerge as strong candidates due to their beneficial services provided to grapevine cultures (Trouvelot et al. 2015), there is a growing need to monitor the endogenous AMF diversity in vineyards. With the goal to investigate AMF diversity in 22 vineyards within the Bordeaux wine region using high-throughput sequencing of the Glomeromycota phylogenetic marker LSU, we have designed a protocol that aims at minimizing the intra-vineyard variability together with reflecting the AMF community interacting with grapevine roots rather than AMF soil populations.

3.2

Approaches, Techniques and Results

3.2.1

Root Sampling in Vineyard

AMF represent an important soil microbial group that interacts with a large spectrum of hosts distributed across many plant families. To date, most of the studies that have investigated AMF communities in vineyards have been performed on soil samples (Lumini et al. 2010; Holland et al. 2013). With the aim to appraise the genuine AMF partners of grapevine with more confidence, we have decided to sample grapevine roots, instead of vineyard soil, the latter likely containing fungal hyphae and spores interacting with roots of cover plants instead of grapevine, together with AMF spores present in a quiescent state. To investigate AMF diversity within the Bordeaux wine region, 22 different vineyards were chosen in the wine-growing region of Bordeaux for a large-scale sampling. We included parcel surfaces that reached up to two hectares and which displayed various soil parameters and grapevine physiological states (i.e. mineral

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Fig. 3.1 Sampling method. (a) Map of a selected parcel, with the 36 sampled plants represented by yellow plots across the parcel. (b) Young grapevine roots are found at the base of the vine-stock in about 20 cm depth. (c) Only thin and young roots with about 1 mm diameter were harvested

and water deficiencies). As these parameters can also greatly vary within the same parcel, we have sought to minimize their incidence by increasing the number of samples, avoiding border rows and by selecting nine consecutives plants in four homogenous rows across the vineyard (Fig. 3.1a). In order to reduce intra-vineyard variability, a total of 36 plants were then sampled for each site and combined in 12 pools (3 neighbours each; Fig. 3.1a). This procedure provides considerably better coverage than the sample sizes used in previous studies (Bouffaud et al. 2016). In addition, in order to harvest only grapevine roots, we sampled at a soil depth of at least 20 cm at the base of each selected plant (Fig. 3.1b).

3.2.2

DNA Extraction

The main steps of the DNA extraction procedure that was applied to all the samples studied in this work are depicted in Fig. 3.2. In the laboratory, roots were first cleaned under a running water tap to eliminate residual soil particles, and only the youngest roots were selected for DNA extraction (Fig. 3.1b). The DNA extraction method was adapted from the “ISO Standard 11063 DNA Extraction Procedure” previously developed for soil samples (Plassart et al. 2012). Cell lysis was performed in 15 ml Falcon tubes containing 1 g (fresh weight) of cleaned roots and 5 ml of lysis buffer (100 mM Tris-EDTA pH ¼ 8, 100 mM NaCl, 2% SDS). In each tube, 2 g silica beads (100 μm diameter), 1 g ceramic beads (1.4 mm diameter) and 1.3 g glass beads (4 mm diameter) were added to physically disrupt root tissues under agitation for 3  30 s runs at 4 m.s1 in a FastPrep®-24 device (MP-Biomedicals) at room temperature. The samples were then incubated for 1 h at 70  C and centrifuged at 7000  g for 5 min at 20  C to pellet cell debris. Proteins were precipitated and removed from the lysate (1 ml) by incubation for 10 min on ice after adding 1/10 volume of 3 M potassium acetate (pH 5.5) and then centrifugation at 14,000  g for 5 min at 4  C. Finally, after overnight precipitation with one volume of ice-cold isopropanol (and centrifugation?), the nucleic acid pellet was washed with 70% ethanol and resuspended in 200 μl nuclease free water. In order to remove phenolic

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Fig. 3.2 “From roots to reads” protocol. Roots were cleaned under water and sorted to keep the thinnest one for DNA extraction. Crude DNA was loaded on PVPP column to retain residual impurities and finally completely purified with GeneClean® kit, allowing to obtain high quality DNA for PCR amplification. The LSU-D2 marker was amplified by nested PCR, amplicons were then sequenced according to MiSeq® technology, and produced reads were filtered and clustered on FROGS pipeline Table 3.1 Molecular results/features at different steps of the analyses

Maximum Minimum Median

DNA yield (ng) 12,225 117 2694

OD 260/280 2,12 1,36 1,88

OD 260/230 2,86 0,67 1,67

Reads after filtering 94,435 1058 22,348

Total OTU 130 15 56

An average of 2.7 μg of DNA per about 1 g of roots was obtained, with good quality indices, leaded to about 22,348 paired-reads, clustered into an average of 56 OUT per sample

compounds and alkaloids contained within residual tannins and humic acids, total crude DNA was next loaded onto mini-columns (BIORAD) filled with 100 to 160 mg of PVPP (polyvinylpolypyrrolidone) and centrifuged at 1000  g for 2 min at 10  C. Nucleic acids were then purified and concentrated using the GeneClean turbo kit (Q-Biogene) with an elution volume of 30 μl nuclease free water. Quantity and quality were evaluated by loading samples on a NanoDrop™ 2000 spectrophotometer (ThermoFisher). Unlike the other kits or classic DNA purification methods we tested, this adapted protocol resulted in high DNA yield with acceptable purity indices for future molecular analysis (Table 3.1). We also found that a longer lysis time tended to increase DNA yield and that an adapted amount of PVPP, depending on the “darkness” of the sample after pellet suspension, is important to avoid elution of impurities or on the contrary DNA retention in the column. The fact that the average values for the OD260/230 ratio were lower than expected (i.e. 1.8) suggests that the DNA still contained residual organic contaminants (Table 3.1).

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3.2.3

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Amplification of AMF Phylogenetic Marker

No consensus DNA barcode is validated for Glomeromycota identification, but the nuclear ribosomal RNA sequences, particularly the SSU (small subunit) and LSU (large subunit) regions, are both widely used in ecological studies (Lindahl et al. 2013; Hart et al. 2015). In our case, we selected the variable D2 region of the LSU, which has proven to be more informative for phylogenetic identification and for species resolution (Stockinger et al. 2010; Schlaeppi et al. 2016). Nested PCRs were performed to amplify the LSU-D2 sequence in sufficient amounts for highthroughput sequencing. PCR was first performed with primers LR1-NDL22 (van Tuinen et al. 1998), which target LSU sequences of all eukaryotes, using 30 ng DNA as template, and the Phusion® High-Fidelity protocol (ThermoFisher). Thermal cycling was performed on a Mastercycler® pro (Eppendorf) with the following conditions: 40 s initial denaturation at 98  C, 25 cycles of 10 s denaturation at 98  C, 30 s annealing at 60  C, 30 s elongation at 72  C and a final elongation of 5 min. After 1/10 dilution of PCR products, 2 μl were used as template with FLR3-T (50 TTG AAA GGG AAA CGA TTG AAG 30 ) and FLR4 primers (Gollotte et al. 2004) modified with the adaptor sequence needed for hybridization to the MiSeq® flow cell. Mix protocol and thermal cycling parameters remained the same as for the first PCR and allowed to produce about 350 bp fragments for all the tested samples. The PCR products were separated by agarose gel electrophoresis (1.2%) and revealed under ultraviolet light after staining with ethidium bromide. The AMF-specific amplicons were obtained for all samples but with different sizes (ranging from 250 to 350 bp) and intensities, thereby suggesting the presence of different species of varied abundance. In cases of primer dimer formation, PCR products were purified from the agarose gel.

3.2.4

High-Throughput Sequencing, Data Processing and Community Analysis

PCR fragments were submitted to the GeT platform of Genotoul (Genopole Toulouse, France) where the routine MiSeq® (Illumina) was applied. This procedure includes an additional PCR amplification of ten cycles to add the terminal part of the adaptor sequence. The libraries obtained were multiplexed on the flow cell and submitted to pair-end sequencing (2 x 250 bp), and an average of 30,000 reads were produced for each library. The FROGS pipeline (Escudié et al. 2016), available on the Galaxy server, was used for the bioinformatic treatment of reads, with adapted parameters. The clustering step was performed with the SWARM aggregation distance of 5 and the abundance threshold of 2x105, resulting in a total of 433 operational taxonomic units (OTU) and a median of 56 OTU per sample (Table 3.1). The taxonomic assignment of the generated OTU was performed using the Maarjam (Öpik et al. 2010) and Silva databases as references. Relative abundances, diversity and dissimilarity indices were obtained using dedicated packages implemented in the R software (Vegan and Phyloseq R packages). As

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Fig. 3.3 Abundance percentages of AMF genera in all samples. (a) General repartition of the AMF genera, dominated by two major genera Glomus and Rhizophagus recovering 96.8%. (b) Repartition of other genera in the remaining 3.2%

already observed from other wine regions (Balestrini et al. 2010; Likar et al. 2012; Holland et al. 2013; Bouffaud et al. 2016), we observed that the majority of sequences belong to the genera Rhizophagus and Glomus (96.8%), but a total of ten different genera of AMF were found associated with grapevine roots (Fig. 3.3).

3.3

Discussion/Conclusions

In our large-scale project, we have established a protocol to study AMF communities in vineyards with a standardized procedure (Fig. 3.2) that yields comparable results from diverse localities and different years. We used root samples rather than soil samples that could contain spores or hyphae from AMF species associated with other plants in the vineyard, e.g. cover plants (Balestrini et al. 2010; Likar et al. 2012) or resting spores without a host. We increased the number of root samples per field plot to obtain more representative coverage of AMF communities, thereby decreasing variability within the vineyards, especially in the case of larger parcels. As expected, dissimilarity indices of Bray-Curtis were on average higher for samples from different vineyards than replication samples from the same vineyard (Table 3.2). This means that the variability intra-vineyard was generally lower than variability inter-vineyard and therefore validates our sampling strategy. In addition, we have modified the DNA extraction procedure of grapevine roots from a protocol dedicated to soil samples (Plassart et al. 2012), in order to increase the yield and the purity of DNA samples and to optimize the subsequent molecular steps. An increasing of lysis time and purification steps has allowed us to obtain DNA in required quantity and quality for PCR amplification and sequencing (Table 3.1). An important point has been the choice of the marker sequence for AMF identification. In our study we have chosen the large subunit (LSU) rRNA gene, known for being phylogenetically slightly more informative than the small subunit (SSU) or the internal transcribed space (ITS) markers (Lee et al. 2013). We have used the FLR3-T/FLR4 primer couple to target the LSU-D2 domain (Gollotte et al. 2004), and we have succeeded in

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Table 3.2 Bray-Curtis dissimilarity indices

Bray-Curtis index Intra-vineyard Inter-vineyard

Mean 0,320 0,631

Min 0,203 0,352

33 Max 0,480 0,840

Dissimilarity indices were calculated between the 22 vineyards, representing 264 samples. Dissimilarity index is always higher in “inter-vineyard” comparison than in “intra-vineyard” comparison

amplification of species from 10 genera, belonging to 6 orders (Acaulosporaceae, Ambisporaceae, Claroideoglomeraceae, Gigasporaceae, Glomeraceae, Paraglomeraceae) of the 11 known orders of the Glomeromycota (Redecker et al. 2013). Moreover very few sequences of pathogenic species or from other fungal taxa were amplified (data not shown), establishing the specificity and the efficiency of this primer couple for studies of AMF populations. We hope that our protocol will allow improving analyses on microbial communities in vineyards and contribute to enlarge the data set on AMF ecology in various regions around the world. Acknowledgements The authors acknowledge the financial support provided by the Bureau Interprofessionnel des Vins de Bourgogne (MYCOVITI Project), the Conseil Interprofessionel du Vin de Bordeaux (project 41971/41975), the Plant Health and Environment department from INRA and funding bodies within the H2020 ERA-net project, CORE Organic Cofund, and with cofunds from the European Commission (BIOVINE project).

References Balestrini R, Magurno F, Walker C, Lumini E, Bianciotto V (2010) Cohorts of arbuscular mycorrhizal fungi (AMF) in Vitis vinifera, a typical Mediterranean fruit crop. Environ Microbiol Rep 2:594–604 Basu S, Rabara RC, Negi S (2018) AMF: the future prospect for sustainable agriculture. Physiol Mol Plant Pathol 102:36–45 Bouffaud M-L, Bernaud E, Colombet A, van Tuinen D, Wipf D, Redecker D (2016) Regional-scale analysis of arbuscular mycorrhizal fungi: the case of Burgundy vineyards. J Int Sci Vigne Vin 50:1–8 Cameron DD, Neal AL, van Wees SCM, Ton J (2013) Mycorrhiza-induced resistance: more than the sum of its parts? Trends Plant Sci 18:539–545 Escudié F, Auer L, Bernard M, Mariadassou M, Cauquil L, Vidal K, Maman S, HernandezRaquet G, Combes S, Pascal G (2016) FROGS: find, rapidly, OTUs with galaxy solution. Bioinformatics 40:1–4 Gianinazzi S, Gollotte A, Binet M-N, van Tuinen D, Redecker D, Wipf D (2010) Agroecology: the key role of arbuscular mycorrhizas in ecosystem services. Mycorrhiza 20:519–530 Gollotte A, van Tuinen D, Atkinson D (2004) Diversity of arbuscular mycorrhizal fungi colonising roots of the grass species Agrostis capillaris and Lolium perenne in a field experiment. Mycorrhiza 14:111–117 Hart MM, Aleklett K, Chagnon P-L, Egan C, Ghignone S, Helgason T, Lekberg Y, Öpik M, Pickles BJ, Waller L (2015) Navigating the labyrinth: a guide to sequence-based, community ecology of arbuscular mycorrhizal fungi. New Phytol 207:235–247 Holland TC, Bowen P, Bogdanoff C, Hart MM (2013) How distinct are arbuscular mycorrhizal fungal communities associating with grapevines? Biol Fertil Soils 50:667–674

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Lee E-H, Eo J-K, Ka K-H, Eom A-H (2013) Diversity of arbuscular mycorrhizal fungi and their roles in ecosystems. Mycobiology 41:121 Likar M, Hančević K, Radić T, Regvar M (2012) Distribution and diversity of arbuscular mycorrhizal fungi in grapevines from production vineyards along the eastern Adriatic coast. Mycorrhiza 23:209–219 Lindahl BD, Nilsson RH, Tedersoo L, Abarenkov K, Carlsen T, Kjøller R, Kõljalg U, Pennanen T, Rosendahl S, Stenlid J et al (2013) Fungal community analysis by high-throughput sequencing of amplified markers - a user's guide. New Phytol 199:288–299 Lumini E, Orgiazzi A, Borriello R, Bonfante P, Bianciotto V (2010) Disclosing arbuscular mycorrhizal fungal biodiversity in soil through a land-use gradient using a pyrosequencing approach. Environ Microbiol 12:2165–2179 Öpik M, Vanatoa A, Vanatoa E, Moora M, Davison J, Kalwij JM, Reier Ü, Zobel M (2010) The online database MaarjAM reveals global and ecosystemic distribution patterns in arbuscular mycorrhizal fungi (Glomeromycota). New Phytol 188:223–241 Plassart P, Terrat S, Thomson B, Griffiths R, Dequiedt S, Lelievre M, Regnier T, Nowak V, Bailey M, Lemanceau P et al (2012) Evaluation of the ISO standard 11063 DNA extraction procedure for assessing soil microbial abundance and community structure. PLoS One 7:e44279 Redecker D, Schüßler A, Stockinger H, Stürmer SL, Morton JB, Walker C (2013) An evidencebased consensus for the classification of arbuscular mycorrhizal fungi (Glomeromycota). Mycorrhiza 23:515–531 Rillig MC, Wright SF, Eviner VT (2002) The role of arbuscular mycorrhizal fungi and glomalin in soil aggregation: comparing effects of five plant species. Plant Soil 238:325–333 Schlaeppi K, Bender SF, Mascher F, Russo G, Patrignani A, Camenzind T, Hempel S, Rillig MC, van der Heijden MGA (2016) High-resolution community profiling of arbuscular mycorrhizal fungi. New Phytol 212:780–791 Spatafora JW, Chang Y, Benny GL, Lazarus K, Smith ME, Berbee ML, Bonito G, Corradi N, Grigoriev I, Gryganskyi A et al (2016) A phylum-level phylogenetic classification of zygomycete fungi based on genome-scale data. Mycologia 108:1028–1046 Smith SE, Read D (2008) Mycorrhizal Symbiosis, 3rd edn. Academic Press, Amsterdam Stockinger H, Krüger M, Schüßler A (2010) DNA barcoding of arbuscular mycorrhizal fungi. New Phytol 187:461–474 Trouvelot S, Bonneau L, Redecker D, Tuinen DV, Adrian M, Wipf D (2015) Arbuscular mycorrhiza symbiosis in viticulture: a review. Agron Sustain Dev 35:1449–1467 van Tuinen D, Jacquot E, Zhao B, Gollotte A, Gianinazzi-Pearson V (1998) Characterization of root colonization profiles by a microcosm community of arbuscular mycorrhizal fungi using 25S rDNA-targeted nested PCR. Mol Ecol 7:879–887

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Molecular Methods for Research on Actinorhiza Hassen Gherbi, Valérie Hocher, Mariama Ngom, Nathalie Diagne, Joëlle Fournier, Alyssa Carre-Mlouka, Luis G. Wall, Louis S. Tisa, and Sergio Svistoonoff

Abstract

Actinorhizal root nodules result from the interaction between a nitrogen-fixing actinomycete from the genus Frankia and roots of dicotyledonous trees and shrubs belonging to 25 genera within 8 plant families. Most actinorhizal plants can reach high rates of nitrogen fixation comparable to those found in root nodule symbiosis of the legumes. As a consequence, these trees are able to grow in poor and disturbed soils and are important elements in plant communities worldwide. While the basic knowledge of these symbiotic associations is still poorly understood, actinorhizal symbioses emerged recently as original systems to explore developmental strategies to form nitrogen-fixing nodules. Many tools have been developed in recent years to explore the interaction between Frankia and actinorhizal plants including molecular biology, biochemistry, and genomics. However, technical difficulties are often encountered to explore these symbiotic H. Gherbi · V. Hocher Laboratoire des Symbioses Tropicales et Méditerranéennes (LSTM), IRD/CIRAD/INRA/ Université Montpellier/Supagro, Montpellier, France M. Ngom Laboratoire Commun de Microbiologie (IRD/ISRA/UCAD), Centre de Recherche de Bel-Air, Dakar, Sénégal Laboratoire Mixte international Adaptation des Plantes et microorganismes associés aux Stress Environnementaux (LAPSE), Dakar, Senegal N. Diagne Laboratoire Mixte international Adaptation des Plantes et microorganismes associés aux Stress Environnementaux (LAPSE), Dakar, Senegal Centre National de Recherches Agronomiques, Institut Sénégalais de Recherches Agricoles (ISRA/ CNRA) BP 53, Bambey, Sénégal J. Fournier Laboratory of Plant-Microbe Interactions (LIPM), CNRS/INRA/ Université Paul Sabatier, Castanet-Tolosan, France © Springer Nature Singapore Pte Ltd. 2019 D. Reinhardt, A. K. Sharma (eds.), Methods in Rhizosphere Biology Research, Rhizosphere Biology, https://doi.org/10.1007/978-981-13-5767-1_4

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interactions, mainly linked to the woody nature of the plant species and to the lack of genetic tools for their bacterial symbionts. In this chapter, we report an inventory of the main recent molecular tools and techniques developed for studying actinorhizae.

4.1

Introduction

4.1.1

Actinorhizal Plants

Actinorhizal plants represent about 200 species encompassing 25 genera in 8 different angiosperm families, in 3 different orders: the Fagales, the Rosales, and the Cucurbitales (Huss-Danell 1997). They are defined as a group by their ability to be nodulated by the diazotroph actinomycete Frankia. In some families, all members are nodulated (Coriariaceae, Elaeagnaceae, Datiscaceae, and Casuarinaceae), whereas in others only a fraction of the genera are nodulated (Betulaceae, Myricaceae, Rhamnaceae, and the Rosaceae). In at least one case (Dryas), nodulation apparently does not extend to all members of a single genus (Kohls et al. 1994). Actinorhizal plants are, with the exception of the Datisca genus, woody perennial dicotyledonous angiosperms widely distributed worldwide except in Antarctica. They can fix high rates of nitrogen which are comparable to those found in legumes. Actinorhizal species are usually known as pioneers on nitrogen-deficient soils and are frequently found in harsh sites, such as glacial till, recent volcanic soils, sand A. Carre-Mlouka Laboratoire des Symbioses Tropicales et Méditerranéennes (LSTM), IRD/CIRAD/INRA/ Université Montpellier/Supagro, Montpellier, France Molécules de Communication et Adaptation des Microorganismes (MCAM, UMR 7245 CNRSMNH N), Muséum national d’Histoire naturelle, Centre national de la Recherche scientifique (CNRS), Sorbonne Universités, Paris, France L. G. Wall LBMIBS, Departamento de Ciencia y Tecnología, Universidad Nacional de Quilmes, Bernal, Argentina L. S. Tisa Department of Molecular, Cellular, and Biomedical Sciences, University of New Hampshire, Durham, NH, USA S. Svistoonoff (*) Laboratoire des Symbioses Tropicales et Méditerranéennes (LSTM), IRD/CIRAD/INRA/ Université Montpellier/Supagro, Montpellier, France Laboratoire Commun de Microbiologie (IRD/ISRA/UCAD), Centre de Recherche de Bel-Air, Dakar, Sénégal Laboratoire Mixte international Adaptation des Plantes et microorganismes associés aux Stress Environnementaux (LAPSE), Dakar, Senegal e-mail: [email protected]

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dunes, clear cuts, and desert (Schwencke and Carú 2001). Actinorhizal plants are thus essential for the N cycle and for the revegetation of various landscapes. They are often used for land stabilization and soil reclamation (Sprent and Parsons 2000).

4.1.2

The Actinomycete Frankia

The microsymbiont Frankia is a filamentous, branching, gram-positive actinobacterium. There is not much knowledge about free-living Frankia in soil (Chaia et al. 2010), while most of the knowledge comes from the symbiotic form or isolates cultured from N2-fixing nodules. Frankia displays three structures in pure culture: vegetative hyphae for the multiplication form, sporangia which constitutes the dissemination form, and vesicles known as the site of nitrogen fixation. No Frankia strain specific to a single actinorhizal host plant species has been described to date. However host specificity is observed at different levels, and correspondence was proposed between the phylogenies of Frankia strains and actinorhizal plants (Wall 2000).

4.1.3

Development of Actinorhizae, the Actinorhizal Nodules

In condition of low nitrogen, actinorhizal nodule development occurs after the emission of unknown signals by plant roots that are perceived by Frankia. Depending on the host plant, two modes of infection of actinorhizal plants by Frankia have been described: intercellular root invasion and intracellular root hair infection (Berry and Sunell 1990; Duhoux et al. 1996; Wall and Berry 2008). Intracellular infection (e.g., of Casuarina, Alnus, Myrica) starts with root hair curling induced by an unknown Frankia signal (Cérémonie et al. 1999; Svistoonoff et al. 2014). Following Frankia penetration through curled root hairs, infection progresses intracellularly in the root cortex (Lalonde and Knowles 1975; Berg 1990). Concomitantly, cell divisions start in the cortex, giving rise to the development of a small external protuberance called the prenodule, whose cells are infected with Frankia (Berry and Sunell 1990). Actinorhizal prenodules do not evolve into nodules and in parallel to prenodule development; cell divisions are observed in pericycle cells opposite to a protoxylem pole, giving rise to an actinorhizal lobe primordium that will be colonized by Frankia. For intercellular infection (e.g., Discaria, Ceanothus, Elaeagnus, Hippophae), Frankia hyphae penetrate between adjacent rhizoderm cells and proceed apoplastically through the root cortex within an electron-dense matrix secreted into the intercellular spaces (Miller and Baker 1986; Racette and Torrey 1989; Liu and Berry 1991; Valverde and Wall 1999a, b; Ibáñez et al. 2016; Fournier et al. 2018). Intercellular-infected actinorhizal species do not develop prenodules, and when the nodule primordium has developed from the pericycle cells, intracellular penetration by Frankia starts acropetally with respect to the root tip in cortical cells of the nodule lobe primordium.

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Specific Issues

Actinorhizal species show long generation times making genetic approaches very difficult to undertake. In addition actinorhizal plants are not model species; standardized protocols are usually not available and have to be optimized for each species. Very often those difficulties are related to lignified tissues present in most actinorhizal plants which render the extraction of proteins or nucleic acids more difficult compared to herbaceous species. The accumulation of dark or opaque compounds also sometimes precludes histological or cell biology approaches. On the Frankia side, one of the major obstacles is the lack of genetics tools. The reasons for the protracted development of genetic tools are complex and include the slow growth rate of Frankia and the difficulty of obtaining single genomic units. Frankia has also a high G + C DNA content (Simonet et al. 1990; Benson and Silvester 1993). Special care is needed when designing PCR primers, and optimization is needed when performing hybridization experiments such as microarrays (Alloisio et al. 2010). Frankia has also a different codon usage compared to model gram-negative bacteria, thus complicating experiments involving heterologous expression of Frankia genes in gram-negative bacteria (Persson et al. 2015; Lurthy et al. 2018) and vice versa (Kucho et al. 2013).

4.2

Approaches, Techniques, and Results

4.2.1

Plant Side

4.2.1.1 Growing Actinorhizal Plants Many actinorhizal plant species are not cultivated; seeds are usually harvested directly from plants growing in the wild or in gardens (Chaia 1998; Valverde and Wall 1999a, b; Gtari et al. 2004; Pawlowski et al. 2007; Gabbarini and Wall 2011; Hocher et al. 2011; Demina et al. 2013). For Casuarinaceae, seeds can be purchased from the Australian Tree Seed Centre (ATSC; https://www.csiro.au/en/Research/ Collections/ATSC/Purchasing-seed). Seeds are usually germinated in soil/vermiculite or soil/sand mixture under greenhouse conditions (Alloisio et al. 2010; Hocher et al. 2011; Gabbarini and Wall 2011; Demina et al. 2013). For experiments requiring axenic conditions, seeds are disinfected, scarified, and sown in petri dishes containing solid agar mineral medium (Smouni et al. 2002; Imanishi et al. 2011; Ktari et al. 2017a, b). 4.2.1.2 Nodulation Experiments Nodulation in Hydroponics To follow nodule development, culture in soil/vermiculite or soil/sand mixture can be used (Demina et al. 2013; Ngom et al. 2016a, b). As an alternative, hydroponic culture is often used with liquid mineral medium free of a nitrogen source (Alloisio et al. 2010; Abdel-Lateif et al. 2013; Imanishi et al. 2014).

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Plant nodulation assay in pouches is a very useful technique where the root grows inside a flat transparent plastic bag with a sheet of paper inside the plastic bag that works as a support device that is saturated by the nutrient solution by capillarity. The growing root is slightly attached to the paper, and the device allows to follow the growth of the root and the development of nodules. Since it is possible to do permanent marks in the plastic bag without hurting the root, it is possible to do measurements of nodule distribution related to the position of the root tip at the moment of inoculation and to analyze this distribution afterward. Using this versatile technique, it was possible to follow the timing of nodule development (Valverde and Wall 1999b) and to study the impact of many environmental factors such as P (Valverde et al. 2002), N (Valverde et al. 2000; Wall et al. 2000, 2003), and Ca (Valverde et al. 2009) that modulate nodulation in actinorhizal plants, as well as autoregulatory factors controlling infection and nodulation (Valverde and Wall 1999b; Valverde et al. 2000) and phytohormone effects (Valverde and Wall 2005). The advantage of growing plants in pouches is also to evidence differential degrees of specificity and recognition (Chaia et al. 2006) as well as the involvement of diffusible signals in the process of early infection before nodule development (Gabbarini and Wall 2011). Spot Inoculation Since Frankia is a non-motile actinomycete, it was possible to develop a simple spot inoculation technique that, combined with the pouch growth method, allows to follow localized plant responses (Obertello and Wall 2015). This is particularly useful with reporter genes fused to promoters of genes suspected to be involved in early plant-microbe interaction (Fournier et al. 2018). Split Root Culture System In order to look for systemic plant signaling related to the symbiotic interaction, split root systems were successfully used in different actinorhizal plant species (Wall and Huss-Danell 1997; Gentili et al. 2006). This kind of experimental approach could be improved using transformed plants to search for the symbiotic plant physiology under a particular genetic condition.

4.2.1.3 Functional Genomics Nucleic Acid Extraction Functional genomics studies often begin with nucleic acid extraction. For different species (Alnus, Casuarina, Discaria), small amounts of RNA can be extracted with standard RNA isolation kits often with modifications suited for lignified species (https://www.qiagen.com/us/resources/resourcedetail?id¼9f16addc-8414-4a5db969-c620e73b3235&lang¼en). Fontainebleau sand can improve sample crushing, and PVPP can be added to the extraction buffer to improve the extraction. RNAs can then be used for further Q-RT-PCR expression analysis (Alloisio et al. 2010; Imanishi et al. 2014; Abdel-Lateif et al. 2013). However, to reach high quantities and quality RNA, protocols based on purification with ultracentrifugation through a

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cesium chloride cushion (Morcillo et al. 2006) or on CTAB extraction buffer (Chomczynski 1993) were successfully applied to Casuarina glauca and Datisca glomerata (Hocher et al. 2011; Demina et al. 2013). DNA extraction can be done using standard DNA extraction kits (Qiagen). Highquality genomic DNA for whole genome sequencing can be obtained with a modified CTAB-based method developed by the company ADNiD (www.adnid.fr; Montpellier, France) followed by quantification with PicoGreen DNA assay (Invitrogen, USA) and quality evaluation with gel electrophoresis, endonuclease degradation, and HinfI digestion tests. Plant Genomic Resources The first genomic platform devoted to the symbiotic process between Frankia and actinorhizal species was developed by Hocher et al. (2006, 2011). Starting from cDNA libraries obtained from non-inoculated roots (controls), inoculated roots (2, 4, and 7 days post-inoculation (dpi)), and nodules (3 weeks post-inoculation), expressed sequence tags (ESTs) were sequenced for C. glauca and Alnus glutinosa. Valid ESTs were processed using a bioinformatic custom pipeline, resulting in about 15,000 unigenes per plant species searchable through http://esttik.cirad.fr/cgi-bin/ public_quick_search.cgi. Transcriptional changes occurring during symbiotic interaction between A. glutinosa and C. glauca and, respectively, Frankia alni and Frankia casuarinae were assessed by developing a custom 15 K Agilent oligonucleotide chip for each species. The comparison of two biological conditions for each plant species (non-inoculated roots and 3-week-old nodules) resulted in the identification of differentially expressed genes (Hocher et al. 2011). The Casuarina 15 K chip was also hybridized with cDNA from C. glauca arbuscular mycorrhiza, giving thus information about arbuscular mycorrhizae in actinorhizal species (Tromas et al. 2012). Expression data are available on NCBI under references GPL10929 for C. glauca and GSE24153 for A. glutinosa, respectively. More recently, a combination of serial analysis of gene expression (SAGE)-type cDNA libraries with 454 GS FLX (Roche) and RNA Seq (Illumina) sequencing technologies was used to generate D. glomerata transcriptome data available at http://fido.nsc.liu.se (Demina et al. 2013). Furthermore, in order to rapidly clone candidate genes identified from transcriptomic studies in C. glauca, a BAC library prepared from nuclei extracted from young stems was constructed in collaboration with CNRGV (INRA, France) and constitutes now an additional genomic resource (Gherbi et al. unpublished). Like for model legume species, the development of -omics data for actinorhizal species allowed analyses which brought new insights in the understanding of these nonlegume symbioses. Global analyses of transcriptomes revealed a similar regulation of gene expression during actinorhizal nodulation very close to what is described for model legumes, suggesting that similar genetic programs govern root nodule symbioses (Hocher et al. 2011; Demina et al. 2013). Several gene families were focused on putative nodulation-associated transcription factors (TF) (Diédhiou et al. 2014), plant defensins (Demina et al. 2013; Carro et al. 2015), auxin signaling genes (Champion et al. 2015), and genes linked to the flavonoid biosynthesis pathway (Auguy et al. 2011; Abdel-Lateif et al. 2013). Orthologs of most legume

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genes described as actors of symbiotic signaling pathways were also identified in C. glauca, A. glutinosa, and D. glomerata, and their expression was found to be comparable between the three actinorhizal species but also comparable to those found in legumes (Hocher et al. 2011; Demina et al. 2013) supporting thus the hypothesis of a conserved signaling pathway for endosymbioses and reinforcing the hypothesis of a common genetic ancestor of the nodulating clade (Hocher et al. 2011). Transcriptomics was also used to search genes induced during AM in the actinorhizal tropical tree C. glauca, and a group of genes was identified corresponding to core functions needed for AM symbiosis, mainly encoding proteases, cytochromes P450, and transporters (Tromas et al. 2012). RNA Seq projects are now developed on the C. glauca-Frankia symbiotic interaction aiming 1) to identify early induced genes linked to the symbiotic process (funded by ANR, France) and 2) to identify genes linked to salt tolerance in C. glauca (funded by DOE-JGI, USA). Recently, sequencing of the first genomes for actinorhizal plants, C. glauca, D. glomerata, A. glutinosa, Dryas drummondii, and Discaria trinervis, was done through the EVO NOD project funded by Beijing Genome Institute (BGI), China (Griesmann et al. 2018). All these data will be made available for scientific community through a Web platform.

4.2.1.4 Genetic Transformation of Actinorhizal Plants Genetic transformation is an essential tool for functional studies on actinorhizae, but its development in woody non-model actinorhizal species has been a major challenge. Several experimental procedures had to be established including the set-up of an efficient plant tissue culture and transformation protocol, the identification of a suitable marker to select transformed cells, and a reliable in vitro regeneration technique. Up to now, Agrobacterium tumefaciens-based transformation has been reported for three species of the Casuarinaceae family: Allocasuarina verticillata (Franche et al. 1997), C. glauca (Smouni et al. 2002), and Casuarina cunninghamiana (Jiang et al. 2015). A. rhizogenes-mediated transformation has also been described for different actinorhizal plants including C. glauca (Diouf et al. 1995), D. glomerata (Markmann et al. 2008), and D. trinervis (Imanishi et al. 2011) (Table 4.1). Agrobacterium tumefaciens-Mediated Transformation of Actinorhizal Plants (Stable) The capacity of actinorhizal plants for genetic transformation was first reported by Mackay et al. (1988) who showed that A. glutinosa and Alnus incana were able to develop tumors after transformation by wild-type strains of A. tumefaciens. The establishment of an efficient transformation protocol using A. tumefaciens-mediated gene transfer was described for A. verticillata (Franche et al. 1997). Transgenic plants were recovered from wounded mature zygotic embryos co-cultivated with the disarmed strain C58C1 (GV2260) in the presence of acetosyringone, NAA (naphthaleneacetic acid), and BAP (benzylaminopurine). The binary vector used for DNA transfer contained the kanamycin resistance gene (NPTII) and the 35S

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Table 4.1 Summary of genetic transformation studies in actinorhizal plants Actinorhizal species Alnus acuminata Alnus glutinosa

Agrobacterium tumefaciens / Tumor induction

Agrobacterium rhizogenes Hairy root induction Hairy root induction

Alnus incana Allocasuarina verticillata Casuarina cunninghamiana Casuarina glauca

Tumor induction Fully transformed plants Fully transformed plants Fully transformed plants /

/ Fully transformed plants /

Composite plants

Diouf et al. (1995), Smouni et al. (2002) Markmann et al. (2008)

/ /

Composite plants “Pseudoactinorhizae”

Imanishi et al. (2011) Savka et al. (1992)

Datisca glomerata Discaria trinervis Elaeagnus angustifolia

Composite plants

References Savka et al. (1992) Mackay et al. (1988), Savka et al. (1992) Mackay et al. (1988) Franche et al. (1997), Phelep et al. (1991) Jiang et al. (2015)

promoter driving the ß-glucuronidase reporter gene. This transformation protocol was improved and further optimized for C. glauca (Smouni et al. 2002). Epicotyls constitute now better targets for T-DNA transfer and calli induction and have a good regeneration ability (Smouni et al. 2002). Transgenic plants can afterward be maintained for years through clonal propagation (Svistoonoff et al. 2010). The availability of an efficient gene transfer method has opened new avenues to better understand the molecular and physiological mechanisms involved in the symbiotic interaction with Frankia and provides an invaluable tool for studying the function of symbiotic genes. The use of ß-glucuronidase (UidA ¼ GUS) or fluorescent proteins (green fluorescent protein ¼ GFP, DsRED, mCherry) as reporter systems allowed the analysis of promoters and cis-acting elements involved in plant gene regulation during the infection process and nodule formation. Thus, spatiotemporal expression of several plant genes playing a key role in the establishment and the development of the nodulation process has been studied (reviewed in PerrineWalker et al. 2011; Svistoonoff et al. 2014). In addition, ectopic activation of promoters from other species has provided information on the conservation of symbiotic mechanisms and the evolution of nitrogen-fixing symbiosis (Franche et al. 1998; Laplaze et al. 2000a; Obertello et al. 2005; Svistoonoff et al. 2010; Sy et al. 2006, 2007). The characterization of Cg12, a gene encoding a subtilase protein, demonstrated the usefulness of reporter systems. The expression of ProCg12:GUS or ProCg12: GFP fusions was linked to infection of root hairs and cortical cells during nodule development of C. glauca and A. verticillata (Svistoonoff et al. 2003) confirming earlier observations made using in situ hybridization (Ribeiro et al. 1995; Laplaze et al. 2000a, b). More recently, the characterization of the CgNIN (¼Nodule INception) the C. glauca ortholog of one of the key early symbiotic TFs in legumes,

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demonstrated that the expression of ProCgNIN:GUS or ProCgNIN:GFP fusions in C. glauca was not only correlated with infection by Frankia but was also induced at preinfection stages by bacterial diffusible factors present in cell-free culture supernatant fluids (Clavijo et al. 2015). This important result opens the way for using ProCgNIN transgenic C. glauca as a bioassay to purify Frankia symbiotic factors (Clavijo et al. 2015; Chabaud et al. 2016). Genetic transformation was also used in loss-of-function approaches using RNA interference (RNAi) targeting CgNIN. Stable RNAi transgenic plants were severely affected in the nodulation process demonstrating the crucial role of CgNIN in the actinorhizal symbiosis (Clavijo et al. 2015). Another great interest of A. tumefaciens-mediated gene transfer is the possibility to introduce diverse tools for cell biology approaches (see Sect. 4.2.1.5). Agrobacterium rhizogenes-Mediated Transformation of Actinorhizal Plants (Hairy Root) A. rhizogenes-based transformation has proven to be a very effective tool for molecular analysis of symbiotic genes during the nodulation process in legumes and actinorhizal plants. The possibility to generate composite plants that consist of a non-transgenic aerial part from which transformed hairy roots can be induced provides an alternative to difficult and laborious production of fully transformed plants using A. tumefaciens. A. rhizogenes-mediated transformation procedure opens the possibility of rapid analysis of gene expression in roots and nodules. Composite plants can be generated within 2–3 months, and nodulation of plants can be achieved 2 months later. Several actinorhizal species have been shown to be susceptible to different A. rhizogenes strains. Alnus glutinosa and A. acuminata were able to develop hairy roots after inoculation of hypocotyls with different A. rhizogenes strains, A4, 1855, 8196, and K599 (Savka et al. 1992), while Elaeagnus angustifolia displayed the formation of nodule-like structures, called pseudoactinorhizae, following inoculation of excised cotyledons with the K599 strain (Savka et al. 1992). The establishment of a transformation and regeneration protocol for actinorhizal plants using A. rhizogenes was first described by Phelep et al. (1991). Fully transformed A. verticillata plants were regenerated by cultivating hairy roots on nutrient medium in the absence of growth regulators. Although transgenic plants were able to form root nodules, they displayed a reduction of apical dominance of the shoots and highly branched and non-geotropic roots. The development of A. rhizogenesmediated transformation method as a tool for the study of the nodulation process in actinorhizal plants was first described by Diouf et al. (1995). In vitro growing seedlings of C. glauca were wounded with a needle dipped in a fresh colony of A4RS strain (Jouanin et al. 1987). Hairy roots were induced in 90% of inoculated plants. After approximately 3 weeks, the original root system was excised, and co-transformed roots containing the transgene from the binary vector were selected. About 50% of roots were co-transformed and 40% of composite plants were nodulated. Transgenic nodules had similar characteristics compared to non-transformed nodules (Diouf et al. 1995). Another A. rhizogenes strain, ARqua1 (Quandt et al. 1993), characterized by low virulence, was also able to induce hairy roots in C. glauca (Abdel-Lateif et al. 2013).

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Datisca glomerata and Discaria trinervis were also successfully transformed. Composite plants of D. glomerata were obtained by inoculation with A. rhizogenes strain LBA 1334 through stem injection of 12-week-old plants (Markmann et al. 2008). Hairy root transformation of D. trinervis was set up using either A4RS or ARqua1 strains and consisted in the inoculation of hypocotyls of seedlings with a needle or sectioning of the radicle with a knife dipped in the bacterial culture. Inoculation of D. trinervis seedlings with ARqua1 strain under ex vitro conditions appeared to give better results (Imanishi et al. 2011). Besides the fact that A. rhizogenes-mediated transformation is a fast system for generating transformed roots, it provides some additional advantageous features: transgenic hairy roots can be easily phenotyped; co-transformed roots can be detected using either an antibiotic resistance gene (NPTII) or reporter systems (fluorescent proteins) included in the T-DNA of the binary vector. Although kanamycin selection of co-transformant roots has been performed for C. glauca (Gherbi et al. unpublished data), fluorescent markers appear to be more convenient as they can be easily visualized and are thus usually used to select co-transformed roots in actinorhizal species (Gherbi et al. 2008; Imanishi et al. 2011; Markman et al. 2008). Composite plants were used to study the molecular mechanisms involved in the actinorhizal symbiosis. The utilization of promoters from early expressed symbiotic genes driving reporter systems made possible the study of the first steps of the infection process in C. glauca (Svistoonoff et al. 2010). Composite plants were also used to downregulate candidate genes through RNAi strategy. Knockdown of CgSYMRK, a receptor-like kinase gene belonging to the common symbiotic signaling pathway (CSSP) and required for both nodulation and mycorrhization in legumes, led to a severe alteration of the nodulation and mycorrhization processes in C. glauca and D. glomerata (Gherbi et al. 2008; Markmann et al. 2008). Other examples include a chalcone synthase (Abdel-Lateif et al. 2013), a calcium- and calmodulin-dependent kinase (CCaMK) (Svistoonoff et al. 2013), and the CgNIN TF (Clavijo et al. 2015) in C. glauca. Genetic engineering of actinorhizal species also offers the possibility to generate gain-of-function mutants. Thus, the expression of auto-active CgCCaMK in C. glauca and D. trinervis led to the induction of spontaneous nodules independently of Frankia as it was previously shown in legumes (Svistoonoff et al. 2013). In Casuarinaceae, the possibility to obtain stable transformant plants using A. tumefaciens offers the opportunity to introduce additional genes using A. rhizogenes transformation of the transgenic shoots.

4.2.1.5 Imaging the Early Stages of Frankia Colonization and Associated Host Responses in Living Tissues In recent years, the development of live tissue imaging approaches has revealed novel aspects of the cellular dynamics of host responses associated with the early steps of the colonization process for both the AM and the legume-rhizobia

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endosymbioses (Genre et al. 2005, 2008; Fournier et al. 2008). The availability of Agrobacterium-mediated transformation techniques for several actinorhizal species has now made it possible to efficiently express fluorescent reporters or cellular markers to explore early calcium signaling and early colonization by in vivo confocal microscopy. Calcium concentration oscillations (a.k.a. calcium spiking) are a central feature of early endosymbiotic signaling and can be detected in vivo using so-called cameleon proteins that are Forster resonance energy transfer (FRET)-based fluorescent calcium concentration reporters. In order to explore the occurrence of nuclear calcium spiking in response to Frankia diffusible signals, a nuclear-targeted cameleon driven by the CaMV 35S promoter (Sieberer et al. 2009) was stably expressed in C. glauca. With this tool, calcium spiking was detected in root hairs of C. glauca following exposure of transgenic root segments to Frankia casuarinae Cci3 exudates previously shown to induce CgNIN expression (Clavijo et al. 2015; Chabaud et al. 2016). The calcium spiking response together with CgNIN expression was subsequently used as a bioassay to further characterize the signal molecules present in the Frankia exudates (Chabaud et al. 2016). Alternatively, microinjection of a calcium indicator dye (Oregon Green) in root hairs can be used to detect oscillations of calcium concentration in vivo in response to a Frankia extract when transformation techniques are not available as is the case for A. glutinosa (Granqvist et al. 2015); however, in contrast to transformation, this approach is limited to root hairs. Regarding early colonization, the lack of a routine transformation protocol for Frankia strains has so far precluded the generation of fluorescently labeled microsymbiont strains and thus the in vivo visualization of penetrating filaments at early colonization stages, limiting in vivo investigation of the associated host responses. This is particularly crucial in species undergoing intercellular infection that lack external signs of infection such as root hair deformations (Ibáñez et al. 2016). Alternative approaches to identify early colonization sites have been developed based on the use of host gene promoters for which expression profiles are tightly associated to the initial colonization by Frankia as tools to target fluorescent reporters either to preinfection responsive cells (ProCgNIN, Clavijo et al. 2015; Chabaud et al. 2016) or cells associated to early penetration sites (ProCg12, Svistoonoff et al. 2003). Remarkably, recent work has shown that ProCg12 can also be used to target early infection sites in D. trinervis, which undergoes intercellular Frankia colonization (Valverde and Wall 1999a, b; Fournier et al. 2018). The expression of a ProCg12-driven fluorescent reporter to target root regions undergoing intercellular infection in D. trinervis can be combined with staining of Frankia filaments with the nucleic acid dye SYTO 9 (5 μM, Molecular Probes) within sections taken from these root zones. This approach has allowed to visualize Frankia filaments in intercellular spaces in the vicinity of ProCg12-expressing cells in D. trinervis (Fournier et al. 2018).

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Bacterial Side

4.2.2.1 Frankia Cultivation and Isolation Frankia is characterized by a slow growth rate and high G + C DNA content (Simonet et al. 1990; Benson and Silvester 1993). After a century of unsuccessful attempts, the first successful isolation of Frankia in culture occurred in 1978 (Callaham et al. 1978). Reasons for these failed attempts are complex and involve the slow growth rate of the bacteria and its original identification as fungus because of its hyphal nature. Although many Frankia isolates have been obtained in the following years, several actinorhizal plant genera have not yielded a Frankia isolate that can fulfill Koch’s postulates. Most Frankia strains are commonly grown in BAP medium (Schwencke 1991). With some Frankia strains, a higher growth rate can be obtained by stirring the cultures (Schwencke 1991) or adding phosphatidylcholine (Selim and Schwencke 1994). Frankia strains belonging to cluster II were thought to be unculturable, but recently Nouioui et al. (2017) reported the successful isolation and cultivation of Frankia coriariae using an alkaline medium. 4.2.2.2 Molecular and “Omics” Tools to Study the Bacterial Partner Frankia Molecular tools were developed for Frankia and used in both investigations in planta or in the soil environment (Samant et al. 2014; Rodriguez et al. 2016; Ben Tekaya et al. 2017). Our discussion will focus on these techniques related to culture work and to in planta studies. In general, genetic analysis of Frankia has been restricted to gene cloning, phylogenetic analyses of selected gene sequences, and some limited studies on plasmid isolation and mutagenesis. Initially, a few studies on spontaneous mutants in Frankia spp. have been reported (Lechevalier et al. 1987; Faureraynaud et al. 1990; Cournoyer and Normand 1994), and chemical (nitroguanidine and ethyl methanesulfonate treatments) and physical (UV) mutagenesis approaches have also been investigated with limited success (Caru and Cabello 1998; Myers and Tisa 2004; Kakoi et al. 2014; Kucho et al. 2017). However, targeted mutagenesis procedures have not yet been established for Frankia. A few Frankia strains contain plasmids (Normand et al. 1983), and several of these plasmids have been cloned and sequenced (Normand et al. 1985; Johnson et al. 1999; John et al. 2001; Lavire et al. 2001; Xu et al. 2002). However, none of these indigenous Frankia plasmids have been transferred to another Frankia strain. There have been three published attempts of DNA transfer in Frankia that have resulted in a low efficiency of transfer or unstably transformed cells (Cournoyer and Normand 1992; Myers and Tisa 2003; Kucho et al. 2009). Because of these impediments and to provide vital information on the symbiosis, the genomes of many Frankia strains have been sequenced (Normand et al. 2007a, b; Tisa et al. 2016), and these databases have opened up the use of “omics” approaches (Benson et al. 2011) discussed below. Analysis of the Frankia genomes revealed the presence of integrated phages (Normand et al. 2007a) and integrating conjugative elements (ICE) in several Frankia strains (Normand et al. 2007a; Ghinet et al. 2011), but so far these elements

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have not been employed as genetic tools. Recently, the successful transformation of several Frankia strains was demonstrated and confirmed by molecular techniques (C. Pesce, R. Oshone, V. Kleiner, S.G. Hurst IV, and L.S. Tisa submitted). The vector introduced into Frankia contained a GFP gene, which was expressed during hyphal growth. Another vector having a cloned Frankia gene from a salt-tolerant strain was also introduced into another Frankia salt-sensitive strain. The cloned gene was expressed in the transformed Frankia salt-sensitive strain providing increased salt tolerance. To gain insight into actinorhizal symbiosis, the genomes of three Frankia isolates that represent two of the four major lineages within the genus were initially sequenced and compared (Normand et al. 2007a, b). The most striking difference among the three genomes was their size, which ranged from 5.43 Mbp for a narrow host range Casuarina strain (Frankia casuarinae strain CcI3) to 7.50 Mbp for a medium host range Alnus strain (Frankia alni strain ACN14a) to 9.04 Mbp for a broad host range Elaeagnus strain (Frankia sp. Strain EAN1pec). The smallest genome belongs to the narrow host range and geographically limited representative strain Cci3, and the largest is from strain EAN1pec that belongs to the broadest host range group. Over the past years, the database of Frankia genome sequences has increased dramatically with over 39 genomes including representatives from all four lineages (Tisa et al. 2016) such as the Cluster 3 lineage in which a broad host range dominates, the Cluster 1 lineage that is characterized by a more restrictive host range, and Cluster 2 that has broad host range, but whose symbionts have proven difficult to isolate in pure culture. Cluster 4 is a special case, since these isolates are unable to reinfect their host plants, or they infect and produce an ineffective symbiosis. Genomes of the Cluster 2 lineage consists of the uncultured Frankia symbionts of D. glomerata [Dg1 and Dg2] (Persson et al. 2011; Nguyen et al. 2016) (Normand et al. 2007a, b) and two recent isolates [Frankia coriariae BMG5.1 and BMG5.30] (Gtari et al. 2015; Nouioui et al. 2016). Analysis of the genomes for both Cluster 2 isolates revealed the absence of common nod genes (Gtari et al. 2015), but the Cluster 3 isolate Frankia sp. strain NRRL B-16219 genome contains canonical nodABCH genes (Ktari et al. 2017a). A PCR-sequencing approach suggested that nod genes are only widespread in Ceanothus americanus microsymbionts. Nod genes could not be detected in any Frankia genomes except for Frankia sp. strain NRRL B-16219. The presence of nod genes in this isolate and in the two Candidatus Frankia Dg1 and Dg2, and the absence in all other genomes, suggests that potentially two distinct nodulation pathways could be involved in actinorhizal symbioses. For the majority of Frankia species, the host plants would be infected through the nod-independent pathway, while the use of a nod-dependent pathway may be involved in some hosts like Ceanothus. These genome datasets are providing a wealth of information and have been used in genome mining (Niemann and Tisa 2008; Perrine-Walker et al. 2010; Furnholm and Tisa 2014; Rehan et al. 2014; Furnholm et al. 2017), comparative genomics (Normand et al. 2007a; Mastronunzio et al. 2008; Sen et al. 2008; Bickhart et al. 2009; Udwary et al. 2011), transcriptomics (Alloisio et al. 2010; Bickhart and Benson 2011; Popovici et al. 2011; Lee et al. 2013; Oshone et al. 2017; Lurthy

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et al. 2018), and proteomics approaches (Alloisio et al. 2007; Bagnarol et al. 2007; Mastronunzio et al. 2008, 2009; Mastronunzio and Benson 2010; Ktari et al. 2017b; Oshone et al. 2017; Ghedira et al. 2018). An analysis of the annotated genome sequence data from the Frankia strains has also revealed several notable genes that are thought to be important for the actinorhizal symbiosis. Bioinformatic analysis of these genomes by the use of the antiSMASH (Blin et al. 2017) and other programs (Udwary et al. 2011) revealed the presence of high numbers of secondary metabolic biosynthetic gene clusters. The Frankia genomes maintain a rich natural product biosynthetic potential comparable to Streptomyces, and many of these compounds are potential signaling molecules (Udwary et al. 2011; Tisa et al. 2016). About 20–30 biosynthetic gene clusters per Frankia genome were identified that encode polyketide synthases, non-ribosomal peptide synthetases, terpenoids, or other specialized lipids which are known to generate natural products. These numbers of biosynthetic gene clusters per genome were comparable to Streptomyces and Salinispora genomes. In many cases, chemical structures of the compounds could be predicted, and these include presumed antibiotics, siderophores, lipids, pigments, and signaling molecules. Despite the high variability of chromosome sizes in these Frankia strains, the numbers of biosynthetic gene clusters were roughly equivalent. Proteomic experiments have validated the presence of similar set of enzymes, while preliminary mass spectrometry studies have confirmed the production of some of the corresponding products (Udwary et al. 2011). This approach was used to identify, isolate, and characterize a polyketide compound from Frankia strain EAN1pec, frankiamicin A (Ogasawara et al. 2015). The potential of these symbionts to generate useful natural products will be important in the identification of novel bioactive molecules and will be continued as more Frankia genomes are sequenced. Bioinformatic analysis of the predicted Frankia secretomes indicated that they are reduced in size compared to those of other soil bacteria (Mastronunzio et al. 2008). Proteome studies on Frankia-secreted proteins have confirmed these findings (Mastronunzio et al. 2009). Transcriptome studies on Frankia have used two different techniques: DNA arrays and RNASeq. Both of these techniques require high-quality RNA from cultures grown under different conditions. To increase the efficiency of both techniques, the amount of stable rRNA present in the total RNA needs to be removed or depleted. Array methods have been used to investigate the global expression of genes under different conditions including under nitrogen stress (Bickhart and Benson 2011; Lee et al. 2013; Lurthy et al. 2018) and in planta (Alloisio et al. 2010). An RNASeq approach has been used to determine global gene expression of Frankia under salt stress (Oshone et al. 2017) and in nodules (Nguyen et al. 2016). Two different approaches have also been used to determine the global expression of the Frankia proteome: 2D gel proteomics and next-generation non-matrix proteomics. The 2D gel approach allows the visual comparison of the protein profiles of Frankia under different conditions but is limited by the number of protein spots that can be identified by MS-MS. It is the most laborious and time-consuming method. However, it has been used for differential global expression of Frankia under

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nitrogen stress (Alloisio et al. 2007), salt stress (Oshone et al. 2017), naphthalene stress (Baker et al. 2015), and exposed to plant extracts (Bagnarol et al. 2007; Popovici et al. 2010, 2011). Next-generation proteome techniques have overcome the use of matrix gel to separate the proteins. This approach has been used to investigate the Frankia response to heavy metals (Furnholm et al. 2017), osmotic stress (Ghedira et al. 2018), in planta (Mastronunzio and Benson 2010), in culture (Alloisio et al. 2007; Mastronunzio et al. 2009), and exposed to plant extracts (Ktari et al. 2017b).

4.2.3

Molecular Tools Used to Analyze Abiotic Stresses on Actinorhizal Plants

Environmental stresses including salinity and heavy metals affect both the growth of actinorhizal plants and beneficial microsymbionts such as Frankia and arbuscular mycorrhizal fungi (AMF) (Evelin et al. 2009; Hanin et al. 2016; Ngom et al. 2016b). Salt tolerance of actinorhizal plants has been studied in many species belonging particularly to families Casuarinaceae (Van der Moezel et al. 1989; Tani and Sasakawa 2003) and Elaeagnaceae (Wen-Hui and Shan 2007; Murata et al. 2012; Maimaiti et al. 2014). These studies showed that, in all these species, plant growth decreased progressively with increased salt concentrations. Similar results were observed in Frankia (Srivastava et al. 2012; Oshone et al. 2017) and AMF strains (Juniper and Abbott 2006; Jahromi et al. 2008) grown in vitro under different levels of salts although, as in plant species, Frankia and AMF isolates varied in their salt tolerance (Evelin et al. 2009; Ngom et al. 2016b). Several studies reported that salinity also affects negatively the establishment of symbioses between actinorhizal plants and Frankia or AMF by reducing root colonization by AMF and plant nodulation by Frankia (Tani and Sasakawa 2003; Mansour et al. 2016; Ngom et al. 2016a). Currently, different molecular methods are used to analyze salt stress on actinorhiza. Because salinity inhibited plant nodulation, Ngom et al. (2016a) investigated the impact of salt stress on the expression of two early symbiotic marker genes: CgNIN (Clavijo et al. 2015) and Cg12 (Svistoonoff et al. 2003), using transgenic plants of C. glauca expressing ProCgNIN:GFP and ProCg12:GFP. This study revealed that salt stress inhibited ProCg12 expression. A qRT-PCR analysis also showed a negative effect of salinity on the expression of a set of plant genes related to carbon and nitrogen metabolisms, as well as to nodule infection by Frankia like Cg12 (Duro et al. 2016). In order to understand the molecular mechanisms of salt tolerance in Frankia strains isolated from Casuarina species, Oshone et al. (2017) used genomic, transcriptomic, and proteomic approaches. Comparative genomic and pangenome analysis revealed that there were 153 shared single copy genes found in the two salt-tolerant strains (CcI6 and Allo2) that were absent in the salt-sensitive (Cci3) and moderately salt-tolerant (CeD) strains. Transcriptome analysis of the highly salt-tolerant strain (CcI6) allowed to identify hundreds of genes that are responsive to salt stress such as genes involved in modulation of membrane

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composition and transport or biosynthesis of compatible solutes. Proteomic profiling confirmed the transcriptome results and revealed additional functions that might be involved in salt stress tolerance (Oshone et al. 2017). Indeed, proteomic analysis allowed to show that a posttranscriptional control seems to be also involved in the regulation of genes under salt stress. In addition to salt stress, the physiological response of Frankia and actinorhizal plants to other unfavorable environmental conditions such as extreme pH and temperature (Tisa et al. 1983; Sayed et al. 2002; Mansour 2003; Gtari et al. 2015), heavy metal toxicity (Richards et al. 2002; Karthikeyan et al. 2009; Bélanger et al. 2015), and chemical and organic stressors (Beauchemin et al. 2012; Mallet and Roy 2014; Baker et al. 2015) has been well documented. However, few molecular studies have been conducted to date to analyze these stresses on actinorhizae. Bioinformatic, qRT-PCR, and proteomic analysis were performed to understand the molecular response of Frankia isolates EuI1c and QA3 to copper (Rehan et al. 2014) and naphthalene (Baker et al. 2015), respectively. Bagnarol et al. (2007) used Myrica seed phenolic extracts and two-dimensional gel electrophoresis to study the molecular mechanisms that control the first steps of actinorhizal symbiosis. Different proteins possibly involved in chaperone biosynthesis, iron transport, nodulation regulation, and oxidative stress were differentially expressed in Frankia strains tested (ACN14a, M16467, and Ea112), suggesting a reorganization of Frankia metabolism for protection against host plant defense (Bagnarol et al. 2007). The same technique of two-dimensional gel electrophoresis was used previously by Hammad et al. (2001) to identify differentially expressed proteins induced in Frankia strain ACN14a by root exudates from its symbiotic host A. glutinosa. Many stress response genes were found in this Frankia strain, among which the sodF gene, was isolated by PCR and genetically well characterized (Hammad et al. 2001; Maréchal et al. 2003).

4.3

Conclusions and Outlook

Molecular research on actinorhiza has been hampered by numerous technical difficulties related to the biology of actinorhizal plants, but many obstacles have fallen in the past 20 years. Genetic transformation protocols are now available for several actinorhizal plant species, and powerful reverse genetics approaches can now be used to demonstrate the function of symbiotic genes; reporter genes can be used to study the regulation of gene expression or as markers of specific cell types. Recent developments related to genetic transformation of Frankia will hopefully allow the development of similarly powerful approaches on the microbial side. The development of genomics tools and particularly the full genome sequencing of several Frankia strains and actinorhizal species is also a major technical achievement offering exciting possibilities to analyze global changes in gene expression associated with actinorhizae. The availability of full genome sequences for several actinorhizal trees also offers the possibility to develop forward genetics approaches using GWAS as it has been done in forest trees (Ingvarsson et al. 2016). On the

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bacterial side, the recent development of gene transfer protocols via mating (C. Pesce, R. Oshone, V. Kleiner, S.G. Hurst IV, and L.S. Tisa, submitted) has overcome a major obstacle in the maturity of this research system. The stability of these Frankia transformants and the expression of exogenous genes in Frankia cells are significant steps in the use of genetics, and efforts are progressing to establish more useful tools. Some improvements are ongoing including the development of a construct that has the reporter mCherry under the F. inefficax strain EuI1c 16S promoter. This construct should allow stable expression of mCherry and would be useful for following the infection pathway within the host plant. Preliminary work on targeted mutagenesis protocols using the CRISPR/Cas9 genome editing tool is showing encouraging progress. A Frankia-specific vector containing the CRISPR/ Cas9 genome editing tool was constructed and is being introduced into Frankia. If successful, we will be able to target specific genes and test for their role in the symbiosis. Like other fields of research, the investigation of molecular mechanisms involved in actinorhizae formation is experiencing a revolution with the possibility to perform very large-scale comparisons with other systems. This flow of new data will certainly lead to major discoveries beyond the field of actinorhiza research but is also a major challenge in the near future as specific models, software implementation, and computational capabilities have still to be developed in order to extract all the relevant biological information. Acknowledgments We gratefully acknowledge support from IRD, CNRS (Project EC2CO), Genoscope, Genopole of Montpellier, and Agence Nationale de la Recherche (AN-06-BLAN0095, BLAN 1708 01, 12-BSV7-0007-02) and United States Department of Agriculture (USDA NIFA 2015-67014-22849, USDA NIFA Hatch 022821, the National University of Quilmes, Argentina (0395/07, 1411/15), The Argentinian National Council for Scientific & Technical Studies (CONICET; PIP 2271 and Bernardo Houssay 2011), and ECOS-SUD (A07B02 and A13B03).

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Sprent JI, Parsons R (2000) Nitrogen fixation in legume and non-legume trees. Field Crop Res 65:183–196 Srivastava A, Singh SS, Mishra AK (2012) Sodium transport and mechanism(s) of sodium tolerance in Frankia strains. J Basic Microbiol Svistoonoff S, Laplaze L, Auguy F, Runions J, Duponnois R, Haseloff J, Franche C, Bogusz D (2003) cg12 expression is specifically linked to infection of root hairs and cortical cells during Casuarina glauca and Allocasuarina verticillata actinorhizal nodule development. Mol PlantMicrobe Interact 16:600–607 Svistoonoff S, Sy MO, Diagne N, Barker DG, Bogusz D, Franche C (2010) Infection-specific activation of the Medicago truncatula ENOD11 early nodulin gene promoter during actinorhizal root nodulation. Mol Plant-Microbe Interact 23:740–747 Svistoonoff S, Hocher V, Gherbi H (2014) Actinorhizal root nodule symbioses: what is signalling telling on the origins of nodulation? Curr Opin Plant Biol 20C:11–18 Svistoonoff S, Benabdoun FM, Nambiar-Veetil M, Imanishi L, Vaissayre V, Cesari S, Diagne N, Hocher V, de Billy F, Bonneau J, Wall L, Ykhlef N, Rosenberg C, Bogusz D, Franche C, Gherbi H (2013) The independent acquisition of plant root nitrogen-fixing symbiosis in Fabids recruited the same genetic pathway for nodule organogenesis. PLoS One 8:e64515 Sy MO, Constans L, Obertello M, Geney C, Laplaze L, Auguy F, Hocher V, Bogusz D, Franche C (2006) Analysis of the expression pattern conferred by the PsEnod12B promoter from the early nodulin gene of Pisum sativum in transgenic actinorhizal trees of the Casuarinaceae family. Plant Soil 281:281–289 Sy MO, Hocher V, Gherbi H, Laplaze L, Auguy F, Bogusz D, Franche C (2007) The cell-cycle promoter cdc2aAt from Arabidopsis thaliana is induced in the lateral roots of the actinorhizal tree Allocasuarina verticillata during the early stages of the symbiotic interaction with Frankia. Physiol Plant 130:409–417 Tani C, Sasakawa H (2003) Salt tolerance of Casuarina equisetifolia and Frankia Ceq1 strain isolated from the root nodules of C. equisetifolia. Soil Sci Plant Nutr 49:215–222 Tisa L, McBride M, Ensign JC (1983) Studies of growth and morphology of Frankia strains EAN1pec, EuI1c, CpI1, and ACN1AG. Can J Bot 61:2768–2773 Tisa LS, Oshone R, Sarkar I, Ktari A, Sen A, Gtari M (2016) Genomic approaches toward understanding the actinorhizal symbiosis: an update on the status of the Frankia genomes. Symbiosis 70:5–16 Tromas A et al (2012) Heart of endosymbioses: transcriptomics reveals a conserved genetic program among arbuscular mycorrhizal, actinorhizal and legume-rhizobial symbioses. PLoS One 7:e44742 Udwary DW et al (2011) Significant natural product biosynthetic potential of actinorhizal symbionts of the genus Frankia, as revealed by comparative genomic and proteomic analyses. Appl Environ Microbiol 77:3617–3625 Valverde C, Wall LG (1999a) Regulation of nodulation in Discaria trinervis (Rhamnaceae)Frankia symbiosis. Can J Bot 77:1302–1310 Valverde C, Wall LG (1999b) Time course of nodule development in the Discaria trinervis (Rhamnaceae)–Frankia symbiosis. New Phytol 141:345–354 Valverde C, Wall LG (2005) Ethylene modulates the susceptibility of the root for nodulation in actinorhizal Discaria trinervis. Physiol Plant 124:121–131 Valverde C, Wall LG, Huss-Danell K (2000) Regulation of nodulation and nodule mass in relation to nitrogenase activity and nitrogen demand in Discaria trinervis (Rhamnaceae) seedlings. Symbiosis 28:49–62 Valverde C, Ferrari A, Gabriel Wall L (2002) Phosphorus and the regulation of nodulation in the actinorhizal symbiosis between Discaria trinervis (Rhamnaceae) and Frankia BCU110501. New Phytol 153:43–51 Valverde C, Ferrari A, Wall LG (2009) Effects of calcium in the nitrogen-fixing symbiosis between actinorhizal Discaria trinervis (Rhamnaceae) and Frankia. Symbiosis 49:151–155

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Molecular and Functional Characterization of Beneficial Bacteria Associated with AMF Spores Monica Agnolucci, Alessandra Turrini, and Manuela Giovannetti

Abstract

In the years to come, a major challenge for agriculture will be the implementation of sustainable intensification of agricultural practice, to ensure sufficient food production for the growing global population and to reduce chemical and energy inputs. This aim may be pursued by promoting the efficient use of beneficial soil microorganisms that play fundamental roles in plant growth and health. Among them, arbuscular mycorrhizal fungi (AMF), and their associated microbiota, can be considered biofertilizers, bioenhancers, and biocontrol agents, showing diverse plant growth-promoting (PGP) properties. Here we focus on approaches for the study of the identity and function of bacteria associated with AMF spores, referred to as spore-associated bacteria (SAB). Culture-independent methods are essential for the identification of their diversity; however, only culture-dependent approaches allow the determination of SAB functional roles, and the selection of the best performing strains, to be tested in laboratory experiments, as well as in the field. The discovery of SAB functional activities, e.g., phosphate solubilization and nitrogen fixation, as well as production of phytohormones, siderophores, and antibiotics, is opening new avenues for their targeted management in agriculture. In this chapter the approaches, techniques, and results relevant to cultureindependent and culture-dependent studies on beneficial SAB will be reviewed. Significant case studies dealing with SAB utilization as inoculants in experimental trials will be discussed, with the aim of prospecting their utilization, individually or in specially designed multifunctional consortia, in sustainable and innovative food production systems.

M. Agnolucci · A. Turrini · M. Giovannetti (*) Department of Agriculture, Food and Environment, University of Pisa, Pisa, Italy e-mail: [email protected] © Springer Nature Singapore Pte Ltd. 2019 D. Reinhardt, A. K. Sharma (eds.), Methods in Rhizosphere Biology Research, Rhizosphere Biology, https://doi.org/10.1007/978-981-13-5767-1_5

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Introduction

In the years to come, one of the major problems to tackle will be represented by food production for a growing global population, while minimizing chemical inputs to the soil and adverse environmental impacts. This objective can be pursued by promoting sustainable methods for intensified agriculture, founded on the efficient use of natural soil resources, such as beneficial microorganisms, that are the fundamental components of soil nutrient flows, playing key roles in the completion of biogeochemical cycles. Among beneficial soil microorganisms, arbuscular mycorrhizal (AM) fungi (AMF) represent a key functional group, facilitating the uptake and transfer of mineral nutrients, such as phosphorus (P), nitrogen (N), sulfur (S), potassium (K), calcium (Ca), copper (Cu), and zinc (Zn), from the soil to the host plants, in exchange for plant carbon, on which they depend as chemoheterotrophic organisms (Smith and Read 2008). AMF are important in agroecosystem processes, as they enhance carbon sequestration and soil aggregation, and plant tolerance to biotic and abiotic stresses (Gianinazzi et al. 2010). Moreover, AMF can also increase the content of beneficial secondary metabolites, an essential property for the production of sustainable high-quality foods (Sbrana et al. 2014). Recent studies reported that the services provided by AMF are often facilitated by the abundant and various microbiota living in association with spores, sporocarps, and extraradical mycelium. Such beneficial microbiota play many plant growthpromoting (PGP) roles, including nitrogen fixation, P solubilization and mineralization, the production of indole acetic acid (IAA), siderophores, and antibiotics (Barea et al. 2002; Rouphael et al. 2015). AMF spores have been identified as a rich source of bacteria (spore-associated bacteria, SAB) to be investigated for their potential PGP activities, with the aim of selecting the best performing strains to be used as biofertilizers and bioenhancers in innovative and sustainable food production systems. The aim of this chapter is to review the developments which contributed to disclose the previously underestimated networks of functional interactions occurring in and around AMF spores. This review will focus on the approaches, techniques, and results that allowed the isolation and selection of SAB strains with specific functional traits.

5.1.1

Arbuscular Mycorrhizal Fungi

AM fungal symbionts belong to the subphylum Glomeromycotina (Spatafora et al. 2016) and show a very low host specificity, establishing mutualistic symbioses (mycorrhiza) with the roots of more than 80% of the species within all major land plant taxa, including the most important food crops, such as cereals, pulses, potatoes, fruit trees, vegetables, and officinal herbs (Smith and Read 2008). AMF are obligate biotrophs, as their life cycle cannot be completed in the absence of host plants. When their spores germinate, AMF produce hyphae able to recognize host roots and to

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differentiate specialized structures on the root surface, the appressoria, which give rise to hyphae growing intercellularly within the root cortex, eventually forming intracellular structures similar to haustoria, the arbuscules. Arbuscules are formed by successive dichotomous hyphal branching and are the key structures of the symbiosis, which are required for nutrient exchange between the two partners: AMF obtain carbon from the host plant and release mineral nutrients absorbed and translocated by the large mycelial network spreading from colonized roots into the surrounding soil (Smith and Read 2008). After reaching their sources of energy and carbon in the host cells, AMF can complete their life cycle producing new spores (Giovannetti 2000) and intraradical vesicles, spore-like storage structures containing lipids. Some AMF species produce spores in the roots, which, in the juvenile stage, are very difficult to distinguish from vesicles. Two types of AM colonization are known: Arum-type and Paris-type (Gallaud 1905, quoted in Smith and Read 2008). The Arum-type is characterized by the spread of fungal symbiont between cortical root cells. Vesicles, when present, are intercellular or intracellular, and arbuscules are terminal on intracellular hyphal branches (Smith and Smith 1997). In the Paris-type intercellular hyphae are not produced, as the fungus spreads directly from cell to cell within the cortex and forms intracellular hyphal coils and intercalary arbuscules along the coils. Most of the experimental works have been carried out on the Arum-type mycorrhizas, which are widely distributed in natural and agricultural ecosystems. The extraradical mycelium (ERM), consisting of a large network of hyphae extending from colonized roots into the soil, represents the key element of the symbiosis, as its structure, extent, and interconnectedness are of fundamental importance for the flow of mineral nutrients absorbed from the soil and translocated to the root cells of host plants. The efficient functioning of such auxiliary absorbing system is determined by the high surface-to-volume ratio of the hyphae, by the hyphal P absorption beyond the P depletion zone around roots, and by the occurrence and differential expression of nutrient transporter genes on ERM hyphae (Karandashov and Bucher 2005; Casieri et al. 2013; Pepe et al. 2017). AMF produce asexual, multinucleate spores, whose phenotypic characteristics, such as shape, color, size, spore walls, subtending hyphae, sporocarp occurrence, and mode of spore germination, are utilized for their morphological identification. AMF spores, whose diameters range from about 50 to 600 μm, develop from extraradical hyphae, either single or aggregated, to form more complex structures, the sporocarps, and live strictly associated with highly diversified microbiota. Some unculturable endobacteria were detected inside the spore cells either by ultrastructural studies (Mosse 1970; MacDonald and Chandler 1981; MacDonald et al. 1982; Bianciotto et al. 1996) or by molecular methods (Naumann et al. 2010; Desirò et al. 2014). Besides these unculturable intracellular organisms, a highly diverse microbial community lives on the spore surface, sometimes sandwiched between the outer and inner spore walls or in the microniches formed by the peridial mycelium surrounding spores and sporocarps (Ames et al. 1989; Walley and Germida 1996; Filippi et al. 1998; Maia and Kimbrough 1998; Artursson and Jansson 2003).

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Bacteria Associated with AMF Spores and Their Functional Roles

SAB were studied by culture-dependent and culture-independent approaches. Molecular studies, such as PCR-denaturing gradient gel electrophoresis (PCR-DGGE) method, allowed the detection of bacteria associated with Funneliformis geosporus, Septoglomus constrictum, and Gigaspora margarita spores (Roesti et al. 2005; Long et al. 2008). We recently identified, by PCR-DGGE and band sequencing, many different bacterial taxa living in close association with the spores of six AMF isolates, belonging to Actinomycetales, Bacillales, Burkholderiales, Pseudomonadales, Rhizobiales, and Mollicutes-related endobacteria (Mre). Interestingly, most of them are known as PGP bacteria, as capable of increasing nutrient availability, by P solubilization, nitrogen fixation, and phytohormone production, and protecting plants against fungal pathogens by the production of antibiotics, siderophores, and hydrolytic enzymes (Agnolucci et al. 2015). With the aim of exploiting PGP bacteria, culture-dependent investigations were carried out, utilizing AMF spores as a source of culturable bacteria, isolated from the spores and spore walls of Glomus versiforme, Rhizophagus clarum NT4, G. margarita, Rhizophagus intraradices, Glomus irregulare, and Funneliformis mosseae (Mayo et al. 1986; Xavier and Germida 2003; Cruz et al. 2008; Bharadwaj et al. 2008b; Lecomte et al. 2011). In a recent study, 374 bacterial strains were isolated in pure culture from R. intraradices spores, with numbers ranging from 5 to 23 CFU per spore (Battini et al. 2016b). Isolated mycorrhizospheric bacteria were characterized for their functional properties, in order to understand how their interaction, either as individual strains or as a consortium, with AMF could affect plant performance. The first functional trait to be assessed was the ability to improve spore germination and boost mycorrhizal activity (Mayo et al. 1986; Xavier and Germida 2003; Giovannetti et al. 2010), which lead to the description of such bacteria as “mycorrhiza helpers” (Frey-Klett et al. 2007). Several studies had previously reported that diverse soil microorganisms affected spore germination and hyphal extension (Mosse 1959; Azcòn 1987, 1989). For example, several Streptomyces species, Pseudomonas sp., and Corynebacterium sp. increased the germination of F. mosseae, Glomus versiforme, and G. margarita spores (Mayo et al. 1986; Mugnier and Mosse 1987; Tylka et al. 1991; CarpenterBoggs et al. 1995). Klebsiella pneumoniae and Trichoderma sp. enhanced hyphal extension of Glomus deserticola and F. mosseae germlings (Will and Sylvia 1990; Calvet et al. 1992), while one bacterium of the Oxalobacteraceae was able to enhance spore germination, germling growth, and root colonization (Pivato et al. 2009). Recent work confirmed that bacterial taxa belonging to Oxalobacteraceae (Burkholderiales) lived strictly associated with hyphae and spores of diverse AMF species and genera (Scheublin et al. 2010; Agnolucci et al. 2015). The mechanism underlying the important functional role of spore germination enhancement was attributed to the capacity of some of the bacterial taxa to degrade chitin, the main component of AMF spore walls, thus facilitating spore germination (Roesti et al.

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2005). Indeed, chitinolytic bacteria were isolated from washed, healthy spores of Glomus macrocarpum and F. mosseae (Ames et al. 1989; Filippi et al. 1998) and from the inner layers of R. intraradices spore walls (Battini et al. 2016b). Besides facilitation of spore germination, the microbiota of the sporosphere may play the role of “mycorrhiza helper” by improving the growth of AMF extraradical mycelium (ERM). For example, Paenibacillus rhizosphaerae, Azospirillum sp., Rhizobium etli, and several Pseudomonas strains significantly improved ERM growth in G. intraradices and R. irregularis in vitro (Bidondo et al. 2011; Ordoñez et al. 2016), while the strains DF57 of Pseudomonas fluorescens and Bc2 of Burkholderia cepacia enhanced mycelial development of Glomus caledonium and G. intraradices in vivo, respectively (Ravnskov and Jakobsen 1999; Ravnskov et al. 2002). Recently, by quantifying the length of AMF hyphae in the soil, Sinorhizobium meliloti TSA41 and Streptomyces sp. W43 N were reported to increase hyphal growth by 24%, compared with hyphal lengths assessed in AMF plants without bacterial inoculation (Battini et al. 2017). The mechanisms underlying this growth promotion could be related to the production of IAA and indole butyric acid (IBA), as the exogenous application of these phytohormones was reported to promote hyphal growth of Diversispora versiformis (Liu et al. 2016). Another fundamental feature of mycorrhizospheric bacteria investigated by many authors was their biocontrol activity against phytopathogens, putatively attributed to their capacity to produce antibiotics (Citernesi et al. 1996; Budi et al. 1999; Li et al. 2007; Bharadwaj et al. 2008a). Actually five Streptomyces isolates, obtained from R. intraradices spores, were molecularly affiliated to strains able to produce the antibiotics chloramphenicol, kirromycin, actinomycin G, and avilamycin A (Battini et al. 2016b). However, also siderophore-producing strains, which in the quoted work represented 66% of all isolates, could play a role in the biocontrol of fungal diseases, due to their ability to inhibit pathogen growth by means of siderophoremediated competition for iron (Davidson 1988; Thomashow et al. 1990; Glick 1995; Arora et al. 2001; Whipps 2001). SAB display other multifunctional PGP activities: they can mediate the uptake of major plant nutrients, such as P and N (Barea et al. 2002). Recent studies reported that highly active P-solubilizing bacteria associated with F. mosseae and R. intraradices spores belong to Streptomyces and Leifsonia (Mohandas et al. 2013) and to S. meliloti (Battini et al. 2016b), respectively. Such bacteria could represent a very important factor in plant nutrition, acting synergistically with AMF to increase P availability, as P is rapidly immobilized and in many soils is unavailable to plant roots. Other studies, utilizing both culture-independent and culturedependent methods, revealed that diverse bacterial species known as N-fixers lived strictly associated with AMF spores and that many strains belonging to Rhizobiales could be isolated, some of which possessing the nifH gene amplicon, confirming the key multifunctional roles played by SAB in mediating the acquisition of major plant nutrients (Bharadwaj et al. 2008b; Agnolucci et al. 2015; Battini et al. 2016b). In the years to come, further research should thoroughly dissect the complex networks of interactions occurring among AMF, associated bacteria, and host plants, in order to reveal the new properties emerging from their possible

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synergies. To this aim, the data on the diversity and composition of AMF-associated bacterial communities obtained by molecular studies should be integrated with those on their functional roles, in the perspective of utilizing the best-performing consortia of AMF symbionts and their associated bacteria in innovative food production systems.

5.2

Approaches, Techniques, and Results

5.2.1

Fungal Material and Spore Collection

Whatever the approach to the study of SAB, the first and indispensable step is represented by spore rinsing, as many and different taxa of generalist bacterial contaminants occur on the surface of spores, either collected from the field or pot cultures. Spores extracted from soil (Gerdermann and Nicolson 1963) were selected under a dissecting microscope, suspended in 1 mL of physiological solution (PS) (9 g L
1 NaCl), rinsed using a vortex mixer at 1500 rpm for 1 min, and then aseptically successively rinsed 15 times in PS. Spores were not rinsed further, as 15 washings were effective for spore surface decontamination.

5.2.2

Culture-Independent Approaches for the Detection of Bacteria Strictly Associated with AMF Spores

5.2.2.1 Techniques Culture-independent approaches are particularly useful when studying SAB, as they are able to overcome the problem of underestimation due to the limitations of cultivation substrates and conditions and of the occurrence of bacteria in viable but non-culturable state. One of the most utilized method is PCR-DGGE analysis of the 16S ribosomal RNA (rRNA) gene, able to obtain the complete fingerprinting of SAB microbiota (Fig. 5.1).

Fig. 5.1 Simplified schematic representation of the detection of the different bacterial species living strictly associated with AMF spores, carried out by using the culture-independent method PCR-DGGE

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The washed spores were homogenized in sterile water, the homogenate was centrifuged at 11,700 g for 20 min, and the supernatant was molecularly analyzed. DNA was extracted using a kit, such as MasterPureTM Yeast DNA Purification kit. Bacterial populations were analyzed by amplification of the V3-V5 of 16S rDNA, utilizing the primers 341 F (CCTACGGGAGGCAGCAG) and 907R (CCGTCAATTCCTTTRAGTTT) (Yu and Morrison, 2004). An additional 40-nucleotide GC-rich tail was added at the primer 341 F 50 end. Amplification was performed in 50 μL, with 10–20 ng of DNA, 5 μL of 10 Gold Buffer (MgCl2-free), 2 mM of MgCl2, 1.25 U of AmpliTaq Gold (Applied Biosystem), 0.2 mM of each dNTPs, and 0.5 μM of each primer. The reactions were performed with a thermocycler with the following cycle parameters: 95  C for 10 min; 94  C for 30s, 55  C for 30s, and 72  C for 60s (for 35 cycles); and 72  C for 10 min. Amplicons of 560 bp were detected by electrophoresis in 1.5% (w/v) agarose gel. For DGGE and fingerprinting analysis, 20 μL of amplicons, supplemented with 20 μL of buffer 2 made with 70% glycerol, 0.05% xylene cyanol, and 0.05% bromophenol blue, were loaded on a 8% polyacrylamide-bisacrylamide (37.5:1) gel with an urea-formamide denaturing gradient ranging from 30 to 65%. A combination of 16S rDNA from several bacterial species was added in the middle and at both ends of each gel as DGGE markers. Gels were run at 80 V and 60  C for 16 h and stained for 30 min in 500 mL of TAE 1 buffer containing 50 μL of SYBR Gold Nucleic Acid Gel Stain. DGGE profiles may be digitally processed and analyzed with BioNumerics software, as reported in Agnolucci et al. (2015), in order to obtain data on the diversity of SAB populations, obtained through clustering and multivariate analyses, determination of richness, dominance, and evenness diversity indices. In addition, the identification of the individual bacterial species was carried out by sequencing the DNA of DGGE bands excised from the gels, using the same primers described above, devoid of the GC-rich tail. Amplicons were purified, quantified, and 50 sequenced. Sequence similarities were determined using the Basic Local Alignment Search Tool (BLASTn). Sequences were aligned with those corresponding to the closest matches from GenBank using MUSCLE as implemented in MEGA software (Edgar 2004a, b), and phylogenetic trees were inferred using the maximum likelihood method based on the Kimura 2-parameter model (Kimura 1980) in MEGA. The confidence of branching was assessed using 1000 bootstrap replicates. The DGGE band sequences were submitted to an official nucleotide archive.

5.2.2.2 Results After PCR-DGGE the profiles obtained from spore homogenates were analyzed. In the case that spores from different AMF species or isolates were investigated, it was possible to compare the banding patterns, analyze them by unweighted pair group method using arithmetic average (UPGMA), and obtain a dendrogram showing the relationships among the different samples, based on similarity and evaluated by the Dice coefficient (Fig. 5.1). If the bands are excised from the DGGE gel and sequenced, it is possible to identify the bacterial species and estimate their relative abundance in the different

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F. coronatus IMA 3

F. mosseae IMA1

F. mosseae AZ225C

R. intraradices IMA5

F. mosseae IN101C

R. intraradices IMA6

Agrobacterium Arthrobacter Geobacillus Bacteroidetes Methylibium Amycolatopsis Uncultered Deltaproteobacteria

Rhizobium Streptomyces Propionibacterium Herbaspirillum M. elongata endobacteria like Massilia Uncultered Bacteroidetes

Sinorhizobium Cupriavidus Pseudomonas Duganella Streptococcus Mre Uncultered Rubrobacter

Bacillus Paenibacillus Ideonella Acidobacteria Lysobacter Uncultered Proteobacteria Uncultered Bacteria

Fig. 5.2 Relative abundance (%) of the microbiota associated with six geographically different AMF isolates belonging to one isolate of F. coronatus, two isolates of R. intraradices, and three isolates of F. mosseae

samples. Figure 5.2 shows the results obtained in a work investigating the microbiota associated with the spores of six different AMF: each isolate was characterized by a diverse bacterial community composition. Species of the genera Arthrobacter and Streptomyces (Actinomycetales) were retrieved, together with members of the orders Burkholderiales, Rhizobiales, Bacillales, and Pseudomonadales and with two different endobacteria related to Mollicutes and Burkholderiaceae (Agnolucci et al. 2015). The high diversity and richness of the bacteria strictly associated with AMF spores have been ascribed to the abundance of nutrients occurring in the sporosphere, a privileged niche where bacteria are able not only to establish and thrive but also to multiply and play multiple key roles, as biofertilizers (phosphatesolubilizing, nitrogen-fixing, and chitinolytic bacteria), biocontrol agents (siderophore- and antibiotic-producing bacteria), and bioenhancers (PGPB).

5.2.3

Culture-Dependent Approaches for the Quantification of Bacteria Associated with AMF Spores

5.2.3.1 Techniques The washed spores (see Sect. 5.2.1) were homogenized and suspended in sterile physiological solution. 100 μL suspension was inoculated onto different microbiological substrates. Spore-forming bacteria were isolated from 1 mL of

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heat-treated (80  C for 10 min) spore suspension. The medium tryptic soy agar (TSA), supplemented with 500 UI L
1 of nystatin and 100 mg L
1 of cycloheximide, was utilized to isolate heterotrophic and spore-forming bacteria.

5.2.3.2 Results SAB abundance was assessed by counting the number of colonies developed after 2 days at 28  C. Then, the selection of bacterial isolates was performed based on phenotypic colony characteristics, i.e., shape, size, edge morphology, surface, and pigment. The isolates should be purified by streaking several times onto the same media utilized for isolation. The pure culture strains can be maintained at
80 C. It is important to mention that from a single spore, it is possible to retrieve 5–23 CFUs (on TSA medium) (Bharadwaj et al. 2008b; Battini et al. 2016b).

5.2.4

Culture-Dependent Approaches for the Detection of SAB Showing Specific Functional Traits

5.2.4.1 Techniques Specific bacterial groups or SAB with particular functional properties were isolated using selective media. For example, Actinobacteria are isolated from Waksman’s agar medium supplemented with 5 mg L
1 of polymyxin and with 100 mg L
1 of cycloheximide and 500 UI L
1 of nystatin to inhibit the growth of gram-negative bacteria and fungi. Chitinolytic bacteria are isolated from minimal medium containing chitin as the only source of carbon (Souza et al. 2009), and putative nitrogen fixers are isolated from Winogradsky agar (Tchan 1984). 100 mg L
1 of cycloheximide and 500 UI L
1 of nystatin were added to inhibit the growth of molds. The bacterial isolates may be further characterized by assessing their PGP activities, such as IAA and siderophore production, P solubilization, and nitrogen fixation ability, and then identified by the sequencing of 16S rDNA (Fig. 5.3). IAA production by SAB isolates was assessed by inoculating the bacteria in 4 mL of Luria–Bertani Broth (LBB), supplemented with 1 mg mL
1 of l-tryptophan, incubated at 20  C in aerobiosis, and centrifuged at 7500 rpm for 10 min. Then, 1 mL of supernatant was added to 2 mL of Salkowski reagent (1.2% FeCl3 in 37% sulfuric acid) and placed in the dark for 30 min. The non-inoculated medium and the medium supplemented with pure IAA represent the negative and positive controls, respectively. Samples developing a red/purple color indicate IAA production. The production of siderophores can be assessed by the overlay chrome azurol S assay (CAS) (Pérez-Miranda et al. 2007). CAS agar is prepared following Louden et al. (2011) using 30.24 g L
1 piperazine-1,4-bis(2-ethanesulfonic acid) (PIPES), 72.9 mg L
1 hexadecyltrimethylammonium bromide (HDTMA), 1 mM FeCl3 6H2O in 10 mM HCl 10 mL, and 0.9 g L
1 bacteriological agar. The bacterial strains, inoculated on TSA, were incubated at 28  C for 2–7 days. Then, 10 mL of CAS agar

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Fig. 5.3 Simplified scheme for isolating and selecting PGP bacterial strains living strictly associated with AMF spores

was overlaid on the bacterial colonies and incubated at 25  C. The strains producing siderophores showed a yellow/orange halo around the colonies, which was measured after 7 days. The capacity of solubilizing organic and inorganic phosphate by SAB is assessed using the National Botanical Research Institute’s phosphate growth medium (NBRIP) (Nautiyal 1999) and phytate-screening medium (PSM) (Jorquera et al. 2008). In the two tests, the bacterial isolates were spotted onto agar plates and grown at 28  C for 7 days. Phytate and phosphate solubilization ability of the relevant bacteria was indicated by halo zones around bacterial colonies that are recorded as well as colony diameter. Bacterial P solubilization capacity is evaluated as phosphate solubilization efficiency (SE), as described by Rokhbakhsh-Zamin et al. (2011). The phosphate solubilization index (PSI) was calculated according to Islam et al. (2007). Putative N-fixers can be screened by PCR amplification of nifH genes. DNA was extracted from microbial cultures grown overnight at 28  C using a kit, such as MasterPureTM Yeast DNA Purification kit. The degenerate primers 19F (50 GCIWTYTAYGGIAARGGIGG-30 ) and 407R (50 -AAICCRCCRCAIACIACRTC30 ) were used to amplify a 390 bp fragment of nifH gene (Ueda et al. 1995). Amplification was carried out in 25 μl, with 10–20 ng of DNA, 1 reaction buffer, 0.2 mM of each dNTPs, 0.5 μM of each primer, and 1.25 U of Takara Ex Taq DNA polymerase. The reaction was carried out in a thermocycler with the following cycles: 94  C 1 min; 94  C 30 s, 56  C 30 s, and 72  C 30 s for 35 cycles; and 72  C 5 min. Amplicons were revealed by electrophoresis in 1.5% (w/v) agarose in TBE 1 buffer gels stained with ethidium bromide 0.5 μg mL
1. The gels were captured as TIFF format files. The selected PGP bacteria were identified by 16S rDNA sequencing. DNA was extracted from liquid cultures grown overnight at 28  C using the MasterPureTM Yeast DNA Purification kit. The amplification of 16S rDNA was carried out using the primers 27f (50 -GAGAGTTTGACTCTGGCTCAG- 30 ) and 1495r (50 CTACGGCTACCTTGTTACGA-30 ) (Lane 1991; Weisburg et al. 1991). PCR was performed in 50 μL, with 10–20 ng of DNA, 1 reaction buffer, 2 mM MgCl2, 1.25 U EuroTaq DNA polymerase, 0.2 mM of each dNTPs, and 0.2 μM of each primer, using a thermocycler with the following cycles: 95  C 2 min; 94  C 1 min and 20s, 54  C 1 min, and 72  C 1 min and 30s for 35 cycles; and 72  C 5 min. PCR amplicons were analyzed, then purified, and sequenced as described above.

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5.2.4.2 Results The number of SAB isolated per spore on TSA medium ranged from 5 to 23 CFUs, comprising on average 1–3 CFUs of spore-forming bacteria, 4–23 CFUs of actinobacteria, 1 CFU of putative N-fixers, and 0.2–1 CFU of chitinolytic bacteria (Bharadwaj et al. 2008b; Battini et al. 2016b). The results obtained from the in vitro screening for PGP traits of strains isolated from TSA and all the other specific media may be expressed (a) as the number or the percentage of bacterial isolates displaying specific PGP traits and (b) as the percentage of bacterial isolates expressing multiple PGP properties. The bacterial isolates producing IAA were discriminated on the basis of the developed levels of intensity in the red/purple color. Accordingly, the radius of the halo of color change allowed the differentiation of variable levels of siderophores produced by SAB. For phytate- and phosphate-solubilizing bacteria, the diameter of the halo zone formed around the colonies differentiated the activity of SAB from low to high (Battini et al. 2016b). Further analyses were carried out on the data obtained, such as the construction of Venn diagrams, to visualize all possible intersections among the relevant functional traits. The sequenced bacterial strains were assigned to species using BLASTn and phylogenetic analyses. Results from BLASTn searches with the 16S rDNA sequences were considered as a match when they showed at least 98% similarity to the query. Affiliation of the sequences with the database 16S rRNA gene sequences may be carried out using neighbor-joining phylogenetic analysis in order to build the relevant phylogenetic trees. Table 5.1 shows the data obtained by the quoted study, with the affiliation of the different SAB strains to the relevant species.

5.3

Discussion

The utilization of culture-independent approaches allowed the detection and identification of specific SAB and the characterization of their diversity, as affected by AMF identity, plant genotype, and environmental conditions. Moreover, SAB molecular identification at the genus/species level represented the first and essential step for proposing their relative contribution to sporosphere communities and their putative roles in this peculiar ecological niche. However, only culture-dependent approaches allow scientists to investigate SAB functional roles, to study their physiological interactions, and to select the best performing strains, among hundreds of isolates, to be further evaluated as biofertilizers and bioenhancers. The regular detection of many Actinobacteria by both methods (cultureindependent and culture-dependent) confirmed their predominance in the mycorrhizosphere (Ames et al. 1989; Filippi et al. 1998) and was correlated with the ability to degrade chitin, a main component of the AMF spore wall, and to hydrolyze biopolymers (Roesti et al. 2005). In particular, species of Arthrobacter and Streptomyces were often retrieved, able to produce a number of enzymes and to biodegrade complex polymers, including chitin and chitosan (Mongodin et al. 2006;

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Table 5.1 Plant growth-promoting traits of bacteria isolated from spores of R. intraradices IMA6

Isolate Sinorhizobium meliloti TSA3 S. meliloti TSA26 S. meliloti TSA41 S. meliloti CH5 S. meliloti CH8 S. meliloti CH17 S. meliloti N23 S. meliloti N28 S. meliloti N29 Streptomyces sp. W43 N Streptomyces sp. W77 Streptomyces sp. W94 Streptomyces sp. W115 Arthrobacter phenanthrenivorans N17 Bacillus pumilus CH10 Fictibacillus barbaricus TSA50 Lysinibacillus fusiformis CH19 Nocardioides albus N13

IAA ++

Siderophore activity


P solubilization SE (%) 115.38

Phytate solubilization halo zone (cm) 0.85

+ ++ ++ +++ +++


++ ++ ++ ++




+ + + + + + ++ ++ + ++ ++

81.82 150 31.25
50 71.43 91.67 84.62 63.64 36.36 54.55 38.46


0.90 0.70 0.25 0.15 0.30 0.65 0.10 0.60 0.80 0.90 1.15 0.50


+ +++

+


69.23


0.25


+




86.67

0.45




++




0.10

Seipke et al. 2012). Such physiological traits were considered essential to AMF beneficial activity, as the partial digestion of AMF outer walls may increase spore germination and germling growth, thus promoting AMF root colonization and symbiosis functioning (Mayo et al. 1986; Carpenter-Boggs et al. 1995; Xavier and Germida 2003; Roestli et al. 2005; Bharadwaj et al. 2008a; Hamdali et al. 2008; Giovannetti et al. 2010). Accordingly, also the presence of SAB taxa affiliated to the Bacillales may represent an important functional trait, as some strains are strong chitin decomposers, producing many kinds of chitinases (Heravi et al. 2014), and may promote mycelial development (Hildebrandt et al. 2006). The isolation and molecular detection of rhizobia from AMF spores, such as Rhizobium and Sinorhizobium, suggest their possible beneficial role as biofertilizers, as they, by nitrogen fixation, can mediate plant acquisition of nitrogen, a major plant nutrient (Bharadwaj et al. 2008b; Agnolucci et al. 2015; Battini et al. 2016b). Accordingly, when spore-associated rhizobial strains were used as inocula, together with AMF, they promoted mycorrhizal functioning by enhancing spore germination, mycelial growth, and mycorrhizal colonization and positively affected plant nutrient uptake (Gopal et al. 2012). Likewise, S. meliloti increased the growth of AMF extraradical mycelium by 19–25% over the levels

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measured in mycorrhizal plants without bacterial inoculation, and they improved plant mineral nutrition (Battini et al. 2017). An important PGP trait of SAB is the ability to solubilize P from mineral phosphate and phytate, as P is a major plant nutrient, occurring at high concentrations in agricultural soils, but being poorly available for plants due to immobilization and precipitation reactions with soil minerals. Moreover, current agriculture is dependent on chemical fertilizers, in particular on phosphate rock P, which is a nonrenewable, finite resource, whose reserves may be depleted in ca.100 years (Cordell et al. 2009). The few works on the occurrence of SAB with P-solubilizing activity reported that strains showing this ability, isolated from F. mosseae spores, belonged to the genera Streptomyces and Leifsonia (Mohandas et al. 2013), while strains isolated from R. intraradices spores belonged to Streptomyces spp., Bacillus pumilus, Lysinibacillus fusiformis, and S. meliloti (Battini et al. 2016b). Such P-mobilizing bacteria, when inoculated together with AMF, could show synergistic activity and enhance P availability to the host plants. Indeed, a recent study reported that some Streptomyces strains facilitated P uptake in maize plants and enhanced the growth of extraradical hyphae, which represent the fungal key structure spreading from mycorrhizal roots, absorbing and translocating P from the surrounding soil to plant roots (Battini et al. 2017). A direct role in the promotion of plant growth may be played by bacteriaproducing phytohormones, mainly IAA, which positively affect many functional activities, such as cell division, elongation, root initiation, and the development of plant root systems (Patten and Glick 2002; Duca et al. 2014). IAA-producing strains were isolated from R. intraradices and F. mosseae spores: most of them were represented by actinobacteria species, followed by S. meliloti, Fictibacillus barbaricus, and Paenibacillus favisporus (Bidondo et al. 2011; Battini et al. 2017). As two of such strains, belonging to the species S. meliloti and P. favisporus, were reported to promote the elongation of AMF extraradical hyphae, the mechanisms underlying such outcome could be ascribed to the alteration of root architecture induced by IAA. The production of siderophores by SAB has been assessed only recently, on R. intraradices spores (Battini et al. 2016b). Such a trait may play an indirect role in the promotion of plant growth, by protecting plants against soil-borne pathogens, as a result of bacterial siderophore-mediated competition for iron (Glick 1995; Whipps 2001). It is important to note that a number of SAB possess multifunctional traits: for example, 17 actinobacterial and 8 chitinolytic strains were able to produce IAA and siderophores and to solubilize P from inorganic and organic forms (Battini et al. 2016b), thus representing good candidates for further tests aimed at evaluating their performance as biocontrol agents, biofertilizers, and bioenhancers. Moreover, recent findings highlighted the ability of some SAB to enhance plant food quality by producing health-promoting phytochemicals (Battini et al. 2016c) and affecting gene expression of key enzymes involved in their biosynthetic pathway (Battini et al. 2016a), in accordance with previous works carried out using PGP rhizobacteria (Copetta et al. 2011; Lingua et al. 2013; Berta et al. 2014; Bona et al. 2015).

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Conclusions and Outlook

In the years to come, a major challenge for agriculture will be the development and implementation of management practices for sustainable intensification of primary production, in order to guarantee enough food crops for the growing global population. Sustainable intensification of agriculture should aim at improving biological soil fertility, which underwent a drastic decline due to the continuous applications of chemical fertilizers and pesticides (Gruhn et al. 2000; FAO 2011). This aim may be pursued by promoting the efficient use of beneficial soil microorganisms that play fundamental roles in biogeochemical cycles and plant nutrition. Among them, the most important group is represented by AMF and their associated bacteria, whose activities enhance the functioning of mycorrhizal symbioses. Culture-independent methods for the study of bacterial communities associated with AMF spores improved our knowledge of their diversity and will contribute to a better understanding of their roles in this peculiar ecological niche. However, only culture-dependent methods allowed the study of the functional roles of SAB, aimed at identifying the most efficient strains, to be further selected as the best performing not only in laboratory experiments but also in the field. The detection of their functional activities, e.g., phosphate solubilization, nitrogen fixation, and production of phytohormones, siderophores, and antibiotics, is opening new avenues for their targeted management in sustainable food production systems. To this aim, the possible synergistic interactions among SAB and among diverse AMF and their SAB should be deeply investigated, in order to understand the functioning of the complex network of microbial interactions and how they affect plant performance. The identification and selection of the most active bacterial strains, inoculated individually or in specially designed multifunctional consortia, will lead to the development of microbial inocula to be used as biofertilizers, bioenhancers, and biocontrol agents in sustainable and innovative food production systems. Acknowledgments This work was funded by a University of Pisa grant (PRA-2015 “Incremento del valore nutraceutico di piante alimentari attraverso l’uso di microrganismi benefici,” Progetti di Ricerca di Ateneo).

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Rokhbakhsh-Zamin F, Sachdev D, Kazemi-Pour N, Engineer A, Pardesi KR, Zinjarde S, Chopade BA (2011) Characterization of plant-growth-promoting traits of Acinetobacter species isolated from rhizosphere of Pennisetum glaucum. J Microbiol Biotechnol 21:556–566. https://doi.org/ 10.4014/jmb.1012.12006 Rouphael Y, Franken P, Schneider C, Schwarz D, Giovannetti M, Agnolucci M, Pascale SD, Bonini P, Colla G (2015) Arbuscular mycorrhizal fungi act as biostimulants in horticultural crops. Sci Hortic 196:91–108. https://doi.org/10.1016/j.scienta.2015.09.002 Sbrana C, Avio L, Giovannetti M (2014) Beneficial mycorrhizal symbionts affecting the production of health-promoting phytochemicals. Electrophoresis 35:1535–1546. https://doi.org/10.1002/ elps.201300568 Scheublin TR, Sanders IR, Keel C, van der Meer JR (2010) Characterisation of microbial communities colonising the hyphal surfaces of arbuscular mycorrhizal fungi. ISME J 4:752–763. https://doi.org/10.1038/ismej.2010.5 Seipke RF, Kaltenpoth M, Hutchings MI (2012) Streptomyces as symbionts: an emerging and widespread theme? FEMS Microbiol Rev 36:862–876. https://doi.org/10.1111/j.1574-6976. 2011.00313.x Smith SE, Read DJ (2008) Mycorrhizal symbiosis. Academic Press, London Smith FA, Smith SE (1997) Structural diversity in (vesicular)–arbuscular mycorrhizal symbioses. New Phytol 137:373–388. https://doi.org/10.1046/j.1469-8137.1997.00848.x Souza CP, Burbano-Rosero EM, Almeida BC, Martins GG, Albertini LS, Rivera ING (2009) Culture medium for isolating chitinolytic bacteria from seawater and plankton. World J Microbiol Biotechnol 25:2079–2082. https://doi.org/10.1007/s11274-009-0098-z Spatafora, J.W., Chang, Y., Benny, G.L., Lazarus, K., Smith, M.E., Berbee, M.L., Bonito, G., . Corradi, N., Grigoriev, I., Gryganskyi, A., James, T.Y., O’Donnell, K., Roberson, R.W., Taylor, T.N., Uehlin, J., Vilgalys, R., White, M.M., Stajich, J.E. (2016). A phylum-level phylogenetic classification of zygomycete fungi based on genome-scale data. Mycologia 108: 1028–1046 https://doi.org/10.1007/s00374-017-1254-5 Tchan YT (1984) Azotobacteraceae. In: Krieg N, Holt JG (eds) Bergey’s manual of systematic bacteriology, vol 1. Williams and Wikins, London, pp 219–225 Thomashow LS, Weller DM, Bonsall RF, Pierson LS (1990) Production of the antibiotic phenazine-1-carboxylic acid by fluorescent Pseudomonas species in the rhizosphere of wheat. Appl Environ Microbiol 56:908–912 Tylka GL, Hussey RS, Roncadori RW (1991) Axenic germination of vesicular–arbuscular mycorrhizal fungi: effects of selected Streptomyces species. Phytopathology 81:754–759 Ueda T, Suga Y, Yahiro N, Matsuguchi T (1995) Phylogeny of sym plasmids of rhizobia by PCR-based sequencing of a nodC segment. J Bacteriol 177:468–472. https://doi.org/10.1128/jb. 177.2.468-472.1995 Walley FL, Germida JJ (1996) Failure to decontaminate Glomus clarum NT4 spores is due to spore wall-associated bacteria. Mycorrhiza 6:43–49. https://doi.org/10.1007/s005720050104 Weisburg WG, Barns SM, Pelletier DA, Lane DJ (1991) 16S ribosomal DNA amplification for phylogenetic study. J Bacteriol 173:697–703. https://doi.org/10.1128/jb.173.2.697-703.1991 Whipps JM (2001) Microbial interactions and biocontrol in the rhizosphere. J Exp Bot 52:487–511. https://doi.org/10.1093/jexbot/52.suppl_1.487 Will ME, Sylvia DM (1990) Interaction of rhizosphere bacteria, fertilizer, and vesicular-arbuscular mycorrhizal fungi with sea oats. Appl Environ Microbiol 56:2073–2079 Xavier LJC, Germida JJ (2003) Bacteria associated with Glomus clarum spores influence mycorrhizal activity. Soil Biol Biochem 35:471–478. https://doi.org/10.1016/S0038-0717(03)00003-8 Yu Z, Morrison M (2004) Comparisons of different hypervariable regions of rrs genes for use in fingerprinting of microbial communities by PCR-denaturing gradient gel electrophoresis. Appl Environ Microbiol 70:4800–4806. https://doi.org/10.1128/AEM.70.8.4800-4806.2004

Part II Plant Pathogens and Microbial Plant Protection

6

Oomycete-Root Interactions Jacob Hargreaves and Pieter van West

Abstract

Oomycetes are descendants of algal-like microorganisms with a natural predisposition to parasitism. They have very specialist adaptations which allow them to infect and kill countless species of plants, many of whom are important food and cash crops. Their asexual progeny, zoospores, show interesting kinetic behavior which may aid their survival in the absence of a plant host; such behavior may actually be applied to facilitate more efficient bioremediation activities. Recent and emergent outbreaks of Phytophthora cinnamomi, P. alni, and P. ramorum have threatened agriculture and horticulture, as well as the wider environment. In response to these and previous outbreaks, methods involving cultural, chemical, and biological control agents have been developed with the aim of stopping outbreaks. In more recent years, an explosion of molecular microbiological research has discovered, for example, RxLR effector proteins, a potential future control target in the farm. Furthermore, other advances are expected with the help of more oomycete genome sequences becoming available and the implementation of the CRISPR/Cas9 gene-editing method. International and interdisciplinary research among scientists should be encouraged, to develop novel control methods and inform legislation and lawmakers to halt the next oomycete that comes around the corner.

J. Hargreaves · P. van West (*) Aberdeen Oomycete Laboratory, Institute of Medical Sciences, University of Aberdeen, Aberdeen, Scotland, UK e-mail: [email protected] © Springer Nature Singapore Pte Ltd. 2019 D. Reinhardt, A. K. Sharma (eds.), Methods in Rhizosphere Biology Research, Rhizosphere Biology, https://doi.org/10.1007/978-981-13-5767-1_6

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Introduction to Oomycetes

Oomycetes are a group of fungal-like microorganisms which share an evolutionary history with brown algae (Beakes et al. 2012). While having various roles within ecosystems, some of which nonpathogenic (Marano et al. 2016), they are widely known for the catastrophic impacts some can have on agriculture. The most infamous incident occurred in the nineteenth century in Ireland, where potato agriculture was almost entirely destroyed by Phytophthora infestans, leading to a large famine (Bourke 1964). Even today, P. infestans causes an estimated global financial loss amounting to €12 billion per annum (Haverkort et al. 2009). Pathogenic oomycetes have developed many highly sophisticated mechanisms to thrive at the expense of their host. Initially, mobile biflagellate zoospores (Morris and Ward 1992) find a host, usually via chemo- or electrotactic mechanisms (Morris et al. 1992; van West et al. 2002), and they encyst and penetrate the host surface via a structure known as appressorium (Meng et al. 2009). Once inside the host, depending on the trophic strategy of the pathogen, they employ various mechanisms to acquire nutrients (Fawke et al. 2015). The basic biological infection mechanism of plant pathogenic oomycetes has been known for a relatively long time (e.g., see reviews by van West et al. 2003; Meng et al. 2009; Fawke et al. 2015). The intention of this review is to cover the most recent advancement in research of oomycetes interacting with plant roots, with the emphasis on the taxonomy of oomycetes and methods of controlling their pathogenic potential on plants.

6.2

Oomycete Taxonomy

6.2.1

Current Taxonomic Placement

Historically the oomycetes were long considered fungi (Lévesque 2011) due to their filamentous growth and trophic strategy, though they exhibited clear differences to “true fungi.” For example, their cell walls contain cellulose, the vegetative hyphae are diploid, and they have many biochemical differences to fungi (Beakes et al. 2012). The development of molecular methods and techniques for fungal and oomycete phylogeny (White et al. 1990; Matsumoto et al. 1999) has helped establish their phylogenetic position on the tree of life. The oomycetes, or Oomycota, are considered either a class or phylum within the kingdom Chromista or Straminipila, respectively. Currently there is still some debate regarding the exact phylogenetic placement of oomycetes. Cavalier-Smith classified the oomycetes into the kingdom Chromista, together with diatoms and brown algae (Cavalier-Smith 1981), classifying them as “pseudo-fungi.” More recently this classification has been questioned; Dick placed the Oomycota within a new kingdom, the Straminipila (Dick 2001), containing members with heterokont flagella whose anterior flagellum is straminipilous. Although the issue is not resolved, the kingdom Straminipila is supported by more contemporary literature, such as that by Beakes and Levesque (Lévesque 2011; Beakes et al. 2012, 2014).

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Fig. 6.1 A Bayesian inference (BI) of phylogeny using the ITS sequences of the main two oomycete clades (the Saprolegniales and Peronosporales), the bootstrap values are given for each branch. red square denotes taxa is a plant pathogen, the pathogenicity of Phytopythium mirpurense has not been demonstrated (represented by a ‘?’), Pythium oligandrum has shown instances of plant pathogenicity as well as a plant-protective mycoparasite (denoted by a yellow square)

The oomycetes themselves are then split further into three groups: the saprolegnian and the peronosporalean clades and the more basal oomycetes (Beakes et al. 2012). In terms of pathogenicity, the saprolegnian clades are generally associated with freshwater animal diseases (van West 2006) and the peronosporalean with plant diseases (Kamoun et al. 2015; Green et al. 2015; Zitnick-Anderson and Nelson 2015). The more basal oomycetes tend to be living in marine environments, infecting, for example, algae and crustaceans, although some are found in terrestrial and freshwater environments (Beakes et al. 2012). A Bayesian inference (BI) of phylogeny using the ITS sequences of the main oomycete clades is shown in Fig. 6.1.

6.2.2

Peronosporales

The major genera of the peronosporalean clade include Phytophthora and Pythium, both of which are associated with plant disease (Kamoun et al. 2015) and contain many root pathogens. Recently a new genus located between Phytophthora and Pythium was introduced in 2010, named Phytopythium (Abad et al. 2010). It was originally classified as Pythium clade K, but phylogenetic analyses showed that it was a genus of its own with features resembling both Pythium and Phytophthora (Abad et al. 2010). More recently, the genus Phytopythium has been completely

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circumscribed by de Cock et al. using analysis of nuclear ribosomal DNA (LSU and SSU) and mitochondrial DNA encoding cytochrome oxidase subunit 1 (COI) (de Cock et al. 2015).

6.2.3

Oomycete Barcoding and Phylogenetic Analysis

The establishment of one, or several suitable region(s) in the genome to be utilized as a barcode, be it for species identification or phylogenetic analysis, is dependent on several important factors. It must be universal in all species, have suitable regions for universal (or mostly universal) primers, and if possible have a high copy number in the genome. An addition, the chosen barcode region must also be relatively conserved, if one wants to carry out analysis on organisms that diverged tens of millions of years ago compared to organisms that diverged only a million years ago. The most common region used as a barcode of oomycetes (and other taxa) is the internal transcribed spacer (ITS) region of the ribosomal genes (Robideau et al. 2011). This region is universal among eukaryotes and evolves rather rapidly, which is why it is favored in identifying and phylogenetically placing species (Matsumoto et al. 1999). For identifying and phylogenetically placing genera and higher taxonomic levels, the large ribosomal subunit (LSU) RNA region and small ribosomal subunit (SSU) RNA region are favored, because they are far more conserved, and are therefore better suited to address the relationships between more distantly related taxa (de Cock et al. 2015). The cytochrome-c-oxidase genes cox1 and cox2 have also been adopted quite widely in the barcoding of oomycetes (Choi et al. 2015). A very useful concise description of methods for molecular phylogenetic analysis can be found in a paper written by Telle et al. (2011) who used molecular phylogenetic analysis to reveal cryptic species of the downy mildews.

6.3

Zoospore Root Targeting

Zoospores are the vegetative biflagellate progeny of oomycetes that are produced asexually from structures called sporangia. They have an important role in plant pathogenicity and have several features which make them ideal for locating and attaching to the host. The production of zoospores is dependent on particular stimuli, in particular wet conditions and temperatures below 12
C. Hence, zoospore formation can be induced by what is known as cold shock (Ribiero 1983). In the case of Phytophthora palmivora, zoospore release was noted after only 5 minutes of cold shock (Walker and van West 2007). Their two flagella (one tinsel, one whiplash) allow them to move and steer themselves in a liquid environment. A study investigating the zoospores of several Pythium and Phytophthora species found that they could move at velocities of 50–250 μm/s (Appiah et al. 2005). Their pathogenicity is bolstered by their ability to detect and respond to various external stimuli such as electrical fields (van West et al. 2002; Appiah et al. 2005) and certain

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chemical compounds (Machlis 1969; Morris and Ward 1992). These features allow plant pathogenic zoospores to find their host and to attach to the root surface, which happens very quickly with production of adhesins, once a suitable host is found (Bartnicki-Garcia 1973).

6.3.1

Bioconvection and Chemotaxis

In addition to the aforementioned chemotaxis and electrotaxis, some zoospores also have an interesting auto-aggregatory behavior which, at high enough zoospore densities, is triggered in some cases even in an absence of an external chemical stimulus, for example, with Achlya (Thomas and Peterson 1990). Work with Achlya (Thomas and Peterson 1990), Pythium, and Phytophthora (Reid et al. 1995) zoospores has shown that if an exogenous attractant is provided to a sample of zoospores, they swim toward the source; this congregating action then causes many zoospores to join the swarm by following other zoospores via a hypothesized, zoospore produced auto-attractant, and not from the initial chemical stimulus. This native behavior indicates that this auto-aggregation is beneficial to zoospore survival (Thomas and Peterson 1990). There is speculation that this behavior might aid the survival of pathogenic zoospores in the absence of host roots, as some zoospores auto-aggregating could persist by remobilizing resources from other zoospores in the aggregate (Reid et al. 1995). Another theory put forward to explain this auto-aggregation behavior is bioconvection; Ochiai et al. found that Phytophthora citricola zoospores do not aggregate by chemo- or phototactic means and do not show auto-attraction. Instead they found that the zoospore complex structures were being formed through bioconvection (the motion of large numbers of small organisms in a fluid) (Ochiai et al. 2011). Similar observations were described by Savory et al. (Savory et al. 2014) who studied the auto-aggregation process of zoospores from P. infestans. They found that zoospores of the late blight pathogen required both chemotaxis and bioconvection in specific, time-separated roles to display the auto-aggregatory behavior (Savory et al. 2014). This discovery was made using mathematical modelling, a novel methodology in this context. Figure 6.2 shows the combined model of auto-aggregating P. infestans zoospores; for more information see their paper (Savory et al. 2014).

6.4

Impact of Recent Oomycete Disease Outbreaks

Despite the existence of a large number of saprotrophic oomycetes (Marano et al. 2016), oomycetes were originally probably “born to kill” because early dividing oomycete genera tend to be parasites, many of whom are obligate parasites, indicating they have been “hard wired” for parasitism (Beakes et al. 2012). A target of many oomycetes are plants; indeed oomycetes have caused devastating outbreaks, which have destroyed industries and led to the starvation of millions (Bourke 1964).

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Fig. 6.2 A comparison of zoospore behaviour over time (top left to bottom right), showing the aggregation of zoospores (aggregates appear as white circular dots in solution) from individual plumes, merging into denser plumes over time (read top left through to bottom right) (a). On the right is the simulated model of zoospore aggregation over time, factoring in both bioconvection and chemotaxis (b), resulting in a model which behaves in the same manner as the actual zoospores in (a). Colours blue to red represent low to high zoospore densities. (Taken with permission from Savory et al. 2014) Table 6.1 The top 10 oomycete pathogens in molecular plant pathology, as voted by 65 scientists in the oomycete plant pathology community Rank 1 ¼2 ¼2

Species Phytophthora infestans Hyaloperonospora arabidopsidis Phytophthora ramorum

4 5

Phytophthora sojae Phytophthora capsici

6 7 ¼8

Plasmopara viticola Phytophthora cinnamomi Phytophthora parasitica

¼8 10

Pythium ultimum Albugo candida

Common disease name(s) Late blight Downy mildew Sudden oak death; Ramorum disease Stem and root rot Blight; stem and fruit rot; various others Downy mildew Root rot; dieback Root and stem rot; various others Damping off; root rot White rust

No. of papers (2005–2014) 1230 137 378 276 541 326 315 142 319 65

Taken from Kamoun et al. (2015)

In 2015, a review of the top oomycete pathogens in molecular plant pathology was compiled (Kamoun et al. 2015). The list shown in Table 6.1 shows the top ten, as voted by the oomycete plant pathogen community. A few examples from the list will be explored in greater detail here.

6.4.1

Phytophthora cinnamomi

P. cinnamomi Rands was first described in Sumatra in 1922; it was isolated from cinnamon trees (hence the name). From Southeast Asia (its suspected region of origin) it has spread globally (Robin and Desprez-Loustau 1998; Hardham 2005),

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and it now poses significant problem in Europe (Brasier 1996), Australia (Hardy et al. 2001), and the Americas (Hardham 2005). In addition to its global reach, it also infects thousands of plant species; this includes important crop plants such as avocado, pineapple, peach, chestnut, and macadamia (Hardham 2005). P. cinnamomi is not an obligate pathogen, showing the ability to live saprotrophically (Zentmyer and Mircetich 1966; Hardham 2005); however, this ability doesn’t appear to be ubiquitous (Collins et al. 2012) and is probably dependent on moisture, with a demonstrated 6-year survival in moist conditions (Zentmyer and Mircetich 1966). Because of the large potential agricultural and ecological costs of this species, various control mechanisms have been developed. Aryantha et al. (2000) found that applying chicken manure to compost substrate before the planting of Lupinus albus seedlings promoted the growth of endospore-forming bacteria, a factor strongly associated with seedling survival when exposed to P. cinnamomi. Another treatment includes applying phosphite salts to plants. This has two different benefits as it directly acts on the pathogen and also causes a protective challenge response to the treated plant (Guest and Grant 1991). This treatment has been discussed at length by Hardy et al. (2001).

6.4.2

Phytophthora alni

P. alni was first discovered in Britain in 1993 (Gibbs 1995), and by 2004 it could be found across Europe (Jung and Blaschke 2004) as the cause of a new devastating disease infecting alder (Alnus spp.), among other species. Causing distinctive bleeding cankers and destroying roots, it normally kills saplings in one season and can kill, or at least significantly damage, mature trees after a few years (Černý and Strnadová 2010). Being such a prolific killer of alder, it can cause huge losses of trees and changes in local ecosystems, as seen in the Czech Republic, where in only 6 years from being first found in the country, it has now spread along hundreds of kilometers of alder stands (Černý and Strnadová 2010). When discovered it was found to be an unknown Phytophthora with certain interesting features, such as a seeming similarity to Phytophthora cambivora (Brasier et al. 2004). The major problem with this assessment was that the unknown Phytophthora (now P. alni) was very specifically pathogenic on alder while P. cambivora was not (Brasier and Kirk 2001). Eventually it was shown that this new Phytophthora species was formed via an interspecific hybridization event between a P. cambivora-like species and an unknown taxon related to Phytophthora fragariae (Brasier et al. 1999). Subsequently, the species P. alni was split into three subspecies: P. alni subsp. alni (Paa), P. alni subsp. uniformis (Pau), and P. alni subsp. multiformis (Pam) (Brasier et al. 2004). Others have since then developed molecular methods to differentiate between the subspecies (Ioos et al. 2005, 2006). Recent findings by Husson et al. (2015) have helped confirm the ancestral status of Paa (being a hybrid of Pau and Pam); they have also recommended that each subspecies be reclassified as individual species, but this has not been widely adopted.

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Attempts to model the impact of climate change to P. alni have been carried out by Aguayo et al. (Aguayo et al. 2014), who modelled the impact of P. alni-induced alder decline in two specific, climactically different regions in France. Their results were modelled on the fact that low winter temperatures and high summer temperatures are unfavorable to the disease, which indicate a mixed outlook for prevalence of P. alni alder disease, depending on geographical location within Europe.

6.4.3

Phytophthora ramorum

P. ramorum was first discovered in 1993 as a new twig blight of Rhododendron species. Observed in Germany and the Netherlands, it was first described in detail in 2001 (Werres et al. 2001). Experiments showed that while P. ramorum is very pathogenic to rhododendron twigs (Werres et al. 2001), there are even more susceptible plant species (Davidson et al. 2003). While considered to be a pathogen of aerial plant systems, pathogenicity via root systems of rhododendrons has also been shown (Parke and Lewis 2007). Within several years after its description, it was already the cause of extensive mortality of oak trees (Quercus spp.) in California, known as sudden oak death (Rizzo et al. 2002). The first outbreak of P. ramorum in the UK (in a garden center) occurred in 2002, causing dieback of Viburnum tinus plants (Lane et al. 2003). By 2003 23 plant species were found to be naturally infected by P. ramorum in 12 plant families, with a pathogen distribution across Europe, Canada, and the western USA (Davidson et al. 2003). More recently, P. ramorum has been the cause of a significant disease outbreak of juvenile Japanese larch (Larix kaempferi) in the UK (sudden larch death), and by 2010 it had spread over 1900 hectares in British larch plantations (Brasier and Webber 2010). Quickly after the emergence of P. ramorum, molecular protocols were developed to detect P. ramorum rapidly, such as a dual detection and quantification tool using real-time PCR with specific P. ramorum primers (Hayden et al. 2004). Other methods to allow for potential detection of P. ramorum (among other species) in the field have been developed (Tomlinson et al. 2005). Molecular analysis using amplified fragment length polymorphism PCR (AFLP-PCR) has indicated that P. ramorum consists of two distinct clusters: a single clonal lineage in North America (NA1) and an array of closely related AFLP types in Europe (EU1) (Ivors et al. 2004). This was expanded on and refined again by Ivors et al. (Ivors et al. 2006), who used microsatellite markers to find a new novel clade in the USA (NA2). Their study also revealed that the EU1 clade was now also detectable in some US nurseries, emphasizing the importance of the nursery trade in the spread of P. ramorum. A fourth lineage of P. ramorum has since been discovered in the UK, coined EU2 (Van Poucke et al. 2012); it is still endemic to the British Isles. P. ramorum is a major problem to the nursery plants industry, and the protocol for nurseries with P. ramorum is now strictly regulated by the USA (December 10, 2012, Federal Order (DA-2012-53) and January 10, 2014, Federal Order (DA-2014-02)) and the European Union (Decision 2002/757/EC).

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Phytophthora austrocedrae, an Emerging Plant Pathogen

A disease of Austrocedrus (Austrocedrus chilensis), otherwise known as Chilean cedar or ciprés de la cordillera, has been studied for many years with an unknown cause. Suggestions included that the disease was being caused by an oomycete (Havrylenko et al. 1989). A Phytophthora species was eventually isolated and described as Phytophthora austrocedrae (Greslebin et al. 2007). In 2010 a new disease of native juniper (Juniperus communis) in northern England (UK) was discovered, with dead and dying juniper found throughout an approximate 14 ha area, with common symptoms including foliage reddening and browning over the crown of the trees and scattered dieback of shoots and branches. It was found that the responsible agent was P. austrocedrae (Green et al. 2012). In 2013 a quantitative method based on real-time PCR was developed for the detection of P. austrocedrae in response to its emergence in the UK (Mulholland et al. 2013), and by 2015 the provision of Koch’s postulates was met (Green et al. 2015). They also found 19 juniper sites with instances of P. austrocedrae scattered across northern England and Scotland, with the authors speculating the instance of moving and standing water to be positively associated with dissemination of P. austrocedrae (Green et al. 2015). According to the British Forestry Commission, this pathogen has now been found across Great Britain in many multiple sites, both in the environment and in plant nurseries.

6.4.5

The Nursery Plant Trade

The global trading of live plant material across wide geographical distances has become a common practice in recent years, and the creation of exotic garden landscapes with imported plants is not a new phenomenon. Improvements in trade and transport mechanisms (Dehnen-Schmutz et al. 2007) have allowed the plantrelated industry to grow, in particular since the 1950s (Brasier 2008). While major problems with invasive plant species, such as Japanese knotweed (Fallopia japonica) in the UK (Hollingsworth and Bailey 2000), are recognized, there is also an overlooked, and potentially catastrophic, transfer of many new microbial species associated with this trade. For comparison, imagine collecting thousands of animal species (many with thousands of members) from the Amazon rainforest and releasing them in the Bornean rainforest, an event that could potentially lead to many extinction events. While this example is not exactly translatable to microbial communities, this is in a similar form being done thousands, if not millions of times every year by horticulturalists who unwittingly introduce microbes associated with imported plants into native soils. Oomycetes are a natural component of the soil microflora, and it has already been shown that nursery trade can be the source of oomycete outbreaks (Brasier 2008). As mentioned previously, there are now strict regulations in both the EU and the USA on importation of plants. The USA does not allow plant importation with soil material, but in the EU, this is not precluded, thus increasing the chances that a

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plant pathogen could be introduced with the soil of a symptomless plant. Such a case has recently been reported by Migliorini et al. (Migliorini et al. 2015), who sampled plants imported from non-EU countries and found Phytophthora in 70% of tested symptomless potted plants. The conclusion of their study was that the EU needs more rigid legislation regarding trade of live plants in pots. These results are echoed from an EU wide study where Phytophthora was found in 91.5% of 732 nurseries tested across 18 EU nations (Jung et al. 2016), and they support the introduction of certain biocontrol strategies (Jung et al. 2016) and improvement of management strategies, as discussed recently by Parke and Grünwald (2012). With increasing private online trade, there have been concerns about Internet sales of plants that could promote the spread and introduction of pathogenic oomycetes, as well as other microbial plant pathogens (Giltrap et al. 2009). These concerns are justified, as shown by a study on plant import in Germany from various non-EU countries, which found that plants sold online are extremely likely not to comply with phytosanitary requirements and for many (68%) customs declarations was missing or incorrect (Kaminski et al. 2012). Furthermore, investigations into the dispersal mode of alien plant species in Poland found that those bought online were transported at significantly greater distances than those bought in shops (Lenda et al. 2014). With our ever greater reliance of the Internet, it appears likely that this problem will persist, if not increase, in the future if no adequate measures are put in place.

6.5

Control of Oomycete Disease

Since the dawn of agriculture, mankind has faced a constant struggle against nature to control an array of crop pests, be it of plant, animal, or microbial origin, including oomycetes. One of the earliest, active preventative measures taken to protect grapes from downy mildew was the treatment of plants with Bordeaux mixture (a mixture of copper(II) sulfate and slaked lime) to prevent infection by Plasmopara viticola (Pickering 1907). Today, control methods are still being developed, be they traditional or cultural, chemical, biological, or a combination of those methods.

6.5.1

Indirect Control Methods

As many of these methods are not new, they will not be covered in much detail; for more detail please refer to Chap. 5 of Phytophthora Diseases Worldwide (Erwin and Ribiero 1996). Phytosanitation or phytosanitary practices are defined as the measures of clean and hygienic practices pertaining to plants. The development and encouragement of phytosanitary practices is not a recent phenomenon; such methods were certainly being seriously developed in the 1950s (Baker 1957). For control of oomycete disease, phytosanitary practices have been shown successful to a limited degree; despite this, in more recent times, phytosanitary methods are still being developed

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and applied (Pérez-Jiménez et al. 2009). A very useful protocol for phytosanitary strategies associated with Phytophthora ramorum has been made (Landis 2013). Solarization is a very simple technique, which relies on transparent polyethylene sheets to trap radiant solar energy to increase soil temperatures and kill unwanted soil pathogens (Katan et al. 1976). The benefit of such a treatment is that it doesn’t rely on chemical treatments, is nonhazardous, and is generally cheaper than other control methods (Katan et al. 1976). This technique was shown to be effective against root rot in cucumber (Cucumis sativus) caused by Pythium ultimum in a trial in Denmark (Christensen and Thinggaard 1999). A similar experiment using solarization against Phytophthora fragariae causing root rot of strawberry and red raspberry was also promising (Pinkerton et al. 2002). However certain plant pathogenic oomycetes appear to be resistant to solarization techniques, as demonstrated in Phytophthora capsici (French-Monar et al. 2007).

6.5.2

Chemical Control Methods

As previously mentioned, Bordeaux mixture was one of the first chemical treatments used to prevent the infection and spread of oomycetes. Since then, there have been many companies developing new chemical treatments, either fungicidal agents or chemicals designed to trigger induced resistance in plants (Machinandiarena et al. 2012). One compound, which is used to control several Phytophthora species, is phosphite (Smillie et al. 1989; Miller et al. 2006). Phosphite is prepared by hydrating a phosphite salt (such as potassium phosphite) and spraying the resultant solution onto the plant. It was found that in potato this triggers an induced resistance through activation of the jasmonic and salicylic acid pathways (Machinandiarena et al. 2012). Interestingly, in sufficient concentrations, phosphite has been shown to directly reduce growth of three Phytophthora species (P. cinnamomi, P. nicotianae, and P. palmivora) (Smillie et al. 1989). Many fungicides have been developed to either kill oomycetes or prevent sporulation and subsequent entry into the host, and experiments have demonstrated their efficacy on major plant disease causing genera, such as Pythium and Phytophthora (Cohen et al. 1995; Rebollar-Alviter et al. 2005). The Pennsylvania State University has a web page that is dedicated to the diagnostics and treatment of root-rot pathogens with a sizable list of fungicidal agents (https://extension.psu.edu/fungalroot-rots-and-chemical-fungicide-use). More recently, partially due to fungicide resistance and environmental concerns of certain fungicidal agents, there have been attempts to develop new, novel chemical treatments (Silva-Castro et al. 2017). For example, presenting P. cinnamomi with a mixture of copper nanoparticles and copper oxychloride reduced its growth by 76%, while beneficial fungi, Trichoderma harzianum and Rhizobium spp., were unaffected (Banik and Pérez-de-Luque 2017). Similar work by Silva-Castro et al. (2017) tested the fungicidal activity of chitosan oligomers, propolis, and nanosilver on well-known plant pathogens of forests. They found that synergistic applications of chitosan oligomers and propolis or silver nanoparticles were especially effective against certain fungi, while for oomycete pathogens Phytophthora alni subsp. alni

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and Phytophthora cambivora, individual treatments of propolis or chitosan oligomers, respectively, were better at inhibiting growth (Silva-Castro et al. 2017).

6.5.3

Biological Control Methods

Biocontrol agents (BCAs) have been widely studied as control methods of a host of pathogens, including oomycetes. There are a number of potential mechanisms that may make a microorganism an effective BCA, competition for limited nutrients, production of antimicrobials, and parasitism (Baker 1987); they can also produce lytic enzymes and stimulate plant immune response (Meena et al. 2017). For example, it was shown that treatment of apple seedlings infected by Phytophthora cactorum (causing root and crown rots), with Trichoderma and Gliocladium spp., after 14 days significantly increased the plant weight and decreased root damage compared to the control (Smith et al. 1990). The bacterium Bacillus subtilis has also been shown to act as a BCA against Phytophthora and Pythium; however, the authors found that the pathogen Botrytis fuckeliana grew resistance against B. subtilis over a period of 4 weeks to levels of being ineffective, warning that BCAs could also fail as treatment methods; they didn’t test the long-term effectiveness with any oomycete pathogen (Li et al. 1998). To increase the efficacy, Ezziyyani et al. (2007) used a combinatorial approach and treated soils containing the pepper (Capsicum annuum) killing oomycete Phytophthora capsici; they found they could reduce P. capsici in the soil by 75%. Work by Timmusk et al. (2009) has also shown the importance of experimenting with a plant system rather than solely in vitro; they used the bacterium Paenibacillus polymyxa to protect Arabidopsis thaliana plants against Phytophthora palmivora and Pythium aphanidermatum, finding that the bacteria’s biofilm formation around the root system of Arabidopsis was crucial in providing protection. Rather than just concentrating on one or several BCAs, there has been research concentrating on whole microbial communities and soil compositions to act as disease suppressors against a range of microbial plant pathogens (Noble and Coventry 2005). The suppressive effects of composts on soil-borne diseases, which included oomycetes like Pythium and Phytophthora, were investigated by Noble and Conventry (2005), who found that compost concentrations above 20% to be a very important factor in suppression of soil-borne pathogens including oomycetes. Subsequent work on compost and compost tea also concluded that the composts suppressed growth of soil-borne diseases, as found by Noble and Coventry (2005), and that sterilized compost does not continue to repress the aforementioned diseases (St. Martin and Brathwaite 2012). Obviously the soil communities are providing a large proportion of the protection against these soil-borne diseases, but with such a complex system as compost, having to take into account the chemical and physical characteristics, there needs to be more investigation and study done (St. Martin and Brathwaite 2012). A very useful book chapter that was recently published on BCAs and the wider role of beneficial microbes can be read by those who are interested (Meena et al. 2017).

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Oomycetes in Bioremediation

Despite the obvious harm done by oomycete plant pathogens, there are instances where oomycetes can be utilized to provide ecological benefits. Example would be the usage of Lagenidium giganteum as a potential biocontrol agent against mosquitos (Kerwin 2007) and Pythium oligandrum in suppressing another oomycete, Pythium ultimum in sugar beet seedlings (Martin and Hancock 1987). However more recently, studies have been conducted showing the potential application of eukaryotic zoospores as facilitators of bioremediation in concert with species of appropriate bacteria. Sungthong et al. (2015) investigated the development of zoospores of Pythium aphanidermatum in the presence of pollutants (polycyclic aromatic hydrocarbons (PAHs)) and PAH-degrading bacteria. They found that the zoospores and bacteria did not interact antagonistically, and the zoospores settled at the water/PAH interface, creating mycelial networks which, along with the movement of the zoospores, enhanced the bioavailability of PAHs. Later studies found that the thrust force created by the motile zoospores in a mobilization assay could mobilize and distribute stationary Mycobacterium gilvum VM552 and slightly motile Pseudomonas putida G7, both PAH-degrading bacteria (Sungthong et al. 2016).

6.7

Methods for Functional Characterization of Oomycete Genes

With a molecular understanding of the mechanisms of oomycete pathogenesis, we can find “chinks in the armor” of the infection/disease process and create treatments to most effectively stop the disease in its tracks. An important step in the development of functional analysis of oomycete genes was the creation of a successful transformation protocol for P. infestans (for initial developed protocol, please see Judelson et al. (1991)). Another big step in the history of molecular biology of oomycetes was the development of double-stranded RNA-mediated transient gene silencing, again in P. infestans (Whisson et al. 2005). This method was faster than the traditional methods of gene silencing and, while only transitory, allowed fast identification and confirmation of gene function. Below are described the important advancements in oomycete molecular biology to date. More recently, the CRISPR/Cas9 technique was successfully implemented in an oomycete (Phytophthora sojae) (Fang and Tyler 2016), which demonstrated its efficacy by replacing the Avr4/6 gene with a reporter gene. The following year they published a detailed protocol of the CRISPR/Cas9 system in P. sojae, with optimized transformation methods for other Phytophthora spp. including P. capsici and P. parasitica (Fang et al. 2017).

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Effector Proteins

Effector proteins are proteins that are secreted by a disease-causing pathogen, where they function to help the pathogen in the infection process. Some effectors play a role in the intercellular spaces between pathogen and host, whereas others are translocated into a host cell, influencing and manipulating host innate immunity (Whisson et al. 2007, Wawra et al. 2012, Anderson et al. 2015, Fawke et al. 2015). It has been shown that plants have an array of disease resistance (R) proteins that can recognize pathogen effector proteins; these R proteins can then elicit a response, such as hypersensitive response and programmed cell death to stop the pathogens in its tracks (Martin et al. 2003). Such pathogen effectors that can be recognized by an R protein are termed avirulence (Avr) proteins, and they have been found in oomycetes (Whisson et al. 2007). For a long time, despite the knowledge of oomycete Avr genes, the mechanism of entry was not yet understood until Rehmany et al. found that an array of Avr proteins in the plant pathogen Peronospora parasitica had a conserved, four amino acid motif, called the RxLR motif (x denoting variable amino acid). They speculated that because of the degree of conservation, the motif must be functionally important, even suggesting that the RxLR motif may play a role in translocating these effector proteins (Rehmany et al. 2005). In 2007 it was shown that the RxLR containing effector protein Avr3a was indeed translocated into the host cell (Whisson et al. 2007). More recent work on the Avr3a effector protein have indicated that the RxLR sequence is actually cleaved before secretion of the protein and thus it is unlikely to interact with the host and play a role in the translocation process itself (Wawra et al. 2017), as was initially thought to be the case (Kale et al. 2010). It is therefore more likely that the RxLR motif in Avr3a is important for intracellular processing, possibly directing the effector protein to the haustoria where it is secreted (Wawra et al. 2017).

6.7.2

Genome Sequencing

The genome sequence of two Phytophthora species, P. sojae and P. ramorum, was published together in a 2006 groundbreaking paper (Tyler et al. 2006); these were the first oomycete genomes sequenced. They identified ~ 19,000 genes in the P. sojae genome and ~ 16,000 in the P. ramorum genome. At this time P. ramorum was an emergent pathogen in California and Europe so for the purposes of disease tracking the group identified ~ 13,700 single nucleotide polymorphisms (SNPs) for development of genetic markers to carry out population genetics analysis (Tyler et al. 2006). This and many other features were discovered thanks to the construction of the genome sequences; a review paper published about this genome sequencing project in more detail can be read (Govers and Gijzen 2006). The genome sequence of Phytophthora infestans was published 3 years later in 2009 (Haas et al. 2009). In 2010 the genome sequence of the plant pathogen Pythium ultimum was published (Lévesque et al. 2010); one of the most surprising results

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from the Pythium genome was that it does not have any classical RxLR effectors. Another interesting discovery was that P. ultimum had a small gene family of cadherins, which are involved in cell adhesion; it was the first report of these proteins in a non-metazoan genome (Lévesque et al. 2010) Around the time that the first Phytophthora genomes were being published, there was, understandably, complementary development of annotation databases to aid the identification of genes, one of the first was VMD (Tripathy et al. 2006). The annotation database used by Lévesque et al. on the genome of P. ultimum was MAKER (Cantarel et al. 2007), they also used PANTHER (Protein ANalysis THrough Evolutionary Relationships) system to find significant differences in gene families (the newest version can be found here (Mi et al. 2017)), and finally they used PHRINGE (Phylogenetic Resources for the Interpretation of Genomes) to reconstruct evolutionary relationships among all oomycete protein families (http:// genomeprojectsolutions.com/PHRINGE_pipeline.html). After the genome publications comes the more detailed analysis, often comparing multiple species’ genomes, which is providing new and very interesting results. Spanu and Kämper (2010) looked at patterns in the published genomes of fungi and oomycetes, remarking at the convergence shown in developing biotrophy: the loss of plant cell wall-degrading enzymes, the expansion and diversification of protein effector families in the genome, and the expansion of transposable elements. Richards et al. demonstrated using comparative genomics that there has been extensive horizontal gene transfer (HGT) between oomycetes and fungi; the HGT genes are widely varied in function, but predicted HGTs include genes associated with resisting plant defense mechanisms and effector proteins (Richards et al. 2011). A good review on these genome analyses in the oomycetes has been written by Judelson (Judelson 2012).

6.8

Conclusions

Research carried out on oomycetes is varied among a plethora of scientific fields and is hugely important. Despite decades of research, oomycetes still pose serious problems to agriculture, horticulture, and the wider environment. The globalized world we live in today presents many opportunities for the next plucky emergent oomycete pathogen. Our opinion is that, moving further ahead, there needs to be: (a) A better global cooperation and integration of research efforts, most importantly of interdisciplinary fields to develop novel pathogen control practices. (b) Opening of clear and frank dialogue with lawmakers when the current legislation is not sufficient to prevent (as much as possible) introduction of pathogens and/or major outbreaks. Acknowledgments Our work is supported by the BBSRC (JH, PvW), NERC (PvW), The South Atlantic Environmental Research Institute (SAERI) (JH), and the University of Aberdeen (PvW).

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Exploitation of Rhizosphere Microbiome Services Valentina Riva, Elisa Terzaghi, Lorenzo Vergani, Francesca Mapelli, Elisabetta Zanardini, Cristiana Morosini, Giuseppe Raspa, Antonio Di Guardo, and Sara Borin

Abstract

The rhizosphere is a soil hot spot where, due to a tight plant-bacteria interaction, plants recruit a beneficial microbiome, enhancing its density and activity. Rhizosphere microbial communities have the potential to provide several services, and their management and “engineering” can be exploited to set up agro-environmental biotechnologies. In this chapter, after a brief overview of the array of services that we can obtain from rhizosphere beneficial microbiome, two case studies are presented: (i) the exploitation of plant growth-promoting bacteria to increase plant tolerance to drought, potentially able to improve crop yield in arid and semiarid lands, and (ii) the exploitation of plant biostimulation effects over degrading microbial populations in the rhizosphere, sustaining phyto-rhizoremediation approaches in PCB-contaminated soils. In each case study, experimental settings, in vitro and in vivo tests, and the result evaluation and modeling are reported together with a discussion of the critical issues.

Valentina Riva and Elisa Terzaghi are contributed equally to the work. V. Riva · L. Vergani · F. Mapelli · S. Borin (*) DeFENS, University of Milan, Milan, Italy e-mail: [email protected] E. Terzaghi · E. Zanardini · C. Morosini · A. Di Guardo DiSAT, University of Insubria, Como, Italy G. Raspa DCEME, Sapienza University of Rome, Rome, Italy © Springer Nature Singapore Pte Ltd. 2019 D. Reinhardt, A. K. Sharma (eds.), Methods in Rhizosphere Biology Research, Rhizosphere Biology, https://doi.org/10.1007/978-981-13-5767-1_7

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Introduction: The Rhizosphere Microbiome and Its Services

The rhizosphere is considered a favorable soil niche where microbial density and activity are enhanced, since the organic compounds released through rhizodeposition constitute a carbon and energy source for the soil dwelling microbiome, contributing to the phenomenon known as the “rhizosphere effect”. The microbial communities inhabiting the rhizosphere usually exhibit a lower phylogenetic diversity than those present in the surrounding non-vegetated soil, due to a selection action mainly driven by the specific profile of the plant rhizodepositions, sources of nutrients and signaling molecules (Bulgarelli et al. 2013; Niu et al. 2017). Plants exploit, therefore, the rhizosphere effect to establish a close association with a core community of beneficial bacteria recruited from the root surrounding soil (Lundberg et al. 2012; Niu et al. 2017). Since both plants and bacteria depend on their tight interactions for survival and maintenance of their fitness, this association can be considered as a meta-organism (Berendsen et al. 2012; Philippot et al. 2013; Thijs et al. 2016). In particular, plant growth-promoting rhizobacteria (PGPR) within the rhizosphere community provide several important services to the plant, such as the enhancement of nutrient bioavailability, the stimulation of root development through hormone regulation, and the protection against pathogens and abiotic stresses (Mendes et al. 2013; Vergani et al. 2017). Given the importance of plant cultivation for the increasing human population, in particular in a contest of climate change and fast soil consumption, these services are also of paramount interest to our society. Beneficial plant-microbe partnerships can be exploited to reduce the water footprint in agriculture, to enhance soil fertility, and to protect crops from diseases, leading to more sustainable agricultural practices as a consequence of water saving and of a reduction in the release of agrochemicals. In addition, PGPR abilities can be useful to sustain the growth of vegetation in stressful environmental conditions like drought, salinity, and soil phytotoxicity, thus helping to counteract the loss of arable land due to desertification and soil degradation, allowing environmental restoration and the reuse of marginal lands (Weyens et al. 2009). The positive plant-microbe relationship established in the rhizosphere could, moreover, sustain an additional service, which is phyto-rhizoremediation which constitutes a promising sustainable approach for the in situ remediation of polluted soils and sediments. It relies on the stimulation by the plant of the degrading microbes in its rhizosphere, in a complex interaction involving roots, root exudates, rhizosphere soils, and microbial communities. A pivotal aspect of rhizoremediation is represented by the occurrence of PGPR associated to plants growing in polluted soils, which can boost their growth and provide various benefits to plants, in turn enhancing the biostimulation of microbial pollutant degraders (Vergani et al. 2017). Both fungi and bacteria have been studied and demonstrated to have an essential role in the rhizosphere and in the services that we can exploit from the rhizosphere. Due to the differences in the physiology of these groups of microbial organisms, the mechanisms of association with the plant and growth promotion are completely different. In this chapter we will focus in particular on the bacterial fraction of the rhizosphere microbiota.

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PGPR as Biofertilizers

Rhizosphere bacteria can act as biofertilizers by increasing the bioavailability of key nutrients including nitrogen, phosphorus, and iron that can be a limiting factor for the plant growth (reviewed by Bulgarelli et al. 2013). This ability can be exploited to minimize the fertilizer input in sustainable agriculture. Plants need nitrogen to be in the form of ammonia or nitrate, so they depend on diazotrophic bacteria able to catalyze the reduction of atmospheric dinitrogen. The most known of these bacteria, such as rhizobia, establish a close association with plants through the formation of nodules on roots, and free-living diazotrophic bacteria in the rhizosphere can provide part of the available nitrogen to a broad range of plants by establishing looser and less specific biocoenosis. Phosphorus is also a mineral nutrient essential for the growth of plants that can only take it up in its soluble forms (H2PO4 and HPO42 ). This bioavailable fraction, however, constitutes approximately only the 0.1% of the total phosphorus content in soils, and many plant species rely on symbiotic associations with fungi such as arbuscular mycorrhizae (AM) and ectomycorrhizae for their phosphorous uptake. For this reason, phosphate must be supplemented with chemical fertilization to increase the biomass of cultivated plants. However, the majority of this phosphorus rapidly becomes unavailable for the plant due to immobilization in the soil. The ability to solubilize both inorganic and organic phosphate forms is common among rhizosphere-associated bacteria, through the release of organic acids or the secretion of extracellular phosphatases. Many bacteria also produce organic compounds called siderophores that chelate Fe3+ and increase its availability for plant uptake after the reduction of Fe3+ to Fe2+ or directly as siderophore-Fe3+ complex. The growth-promoting influence of siderophoreproducing bacteria is particularly important when the plant is growing under ironlimiting conditions, such as alkaline soils. The use of PGPR as biofertilizer inocula in experimental studies has proved their ability to increase plant biomass, revealing their potential application to increase cultivated plant productivity and reduce the release of chemical fertilizers (Weyens et al. 2009; Mendes et al. 2013).

7.1.2

PGPR as Biostimulants and for Stress Mitigation

PGPRs promote the growth of associated plants also by interfering with phytohormone production, having the effect of improving plant nutrition and increasing tolerance toward environmental stress. Among the phytohormones produced by bacteria, the most studied is the auxin indole-3-acetic acid (IAA) related to the promotion of root elongation and lateral root development. The effectiveness of IAA-producing bacteria in promoting the development of the root apparatus has been observed by different bacterial strains on several plant species. This represents a significant advantage for the plant, which can explore an increased area of soil for water and nutrient uptake. Nonetheless, this beneficial effect seems to depend on the auxin concentration, since high level of this hormone may inhibit root growth (Weyens et al. 2009). Rhizosphere bacteria can also modulate the production of the

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plant hormone ethylene, through the activity of the enzyme 1-aminocyclopropane-1carboxylic acid (ACC) deaminase. This enzyme hydrolyzes the immediate precursor of ethylene in plants, thereby lowering the rate of ethylene biosynthesis that is induced by biotic and abiotic stress and leads to an inhibition of plant development. In this way, PGPR bacteria help the plant to counteract environmental stressful conditions without affecting the small burst of ethylene, which is believed to activate key plant defense responses (Mendes et al. 2013; Vergani et al. 2017). Also the production of gibberellins by PGPR has been described to sustain the plant in counteracting salinity stress (Leitão and Enguita 2016). Besides interfering with plant homeostasis, several other mechanisms have been associated to the capacity of PGPR to alleviate environmental stress in plants, in particular improving adaptation to saline and drought stresses (Qin et al. 2016).

7.1.3

PGPR for the Protection from Plant Pathogens

PGPR can have indirect promotion mechanisms by protecting the plant against pathogens, in turn improving its health status. Pathogen protection can occur in several ways, such as the production of antimicrobial compounds and lithic enzymes, the induction of systemic resistance in plants, and the competition for plant-derived nutrient that can hinder plant pathogens leading to the development of disease-suppressive soils. A large body of literature is available on this topic (reviewed, among other, by Berendsen et al. 2012; Bulgarelli et al. 2013; Hu et al. 2016; Rahman et al. 2018), since the exploitation of PGPR for biological control of plant diseases and crop protection is very promising in sustainable agriculture practices, potentially able to decrease pesticide release in the environment and the consequent risks related to the emerging of antimicrobial-resistant populations and genes (Chowdhary and Meis 2018).

7.2

Plant-Bacteria Interactions for the Rhizoremediation of Polluted Soils

Thanks to microbial degradation abilities, detoxification occurs naturally in polluted soils, a process called natural attenuation. Rhizoremediation is a specific type of phytoremediation that involves both plants and their associated rhizosphere microbiome, and it has been described as an effective strategy for the detoxification of organic contaminants from impacted soils. It relies on the stimulation of the pollutant degrading microbes by the plant, which provides root exudates acting as nutrients or triggering the degradation of organic pollutants through co-metabolism, while, in turn, PGPR bacteria protect the plant from the stress derived from soil toxicity and promote its growth through the mechanisms described above (Vergani et al. 2017). However, the efficiency of rhizoremediation is strongly dependent on the complexity of soil contamination, the plant species, and the establishment of beneficial interactions between the plant and the soil degrading microbiome (Vergani et al. 2017; Thijs et al. 2016). Hence, the natural attenuation process can

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be exploited by engineering the rhizosphere microbiome with interventions on the plant host, nutrient supply, soil chemical and physical parameters, and with the inoculation of competitive bacterial strains, in order to develop more efficient and site-specific rhizoremediation strategies for the cleanup of contaminated soils (Thijs et al. 2016). In the following chapters, two different case studies will be described, which exploit the beneficial rhizosphere microbiome to obtain two services of relevant importance for restoration and sustainable water management in arid and semiarid lands and remediation of contaminated soils.

7.3

Case Study 1: Improvement of Plant Drought Tolerance

Drought is one of the major environmental stresses that limit plant growth and agricultural productivity around the world. Because of the ongoing global warming and climate changes, it is projected that the land area affected by drought will increase by twofold and water resources will decline by 30% by 2050 (Falkenmark 2013). Finding efficient strategies for the mitigation of drought stress is therefore increasingly gaining importance.

7.3.1

Microbial Mechanisms Responsible of the Improvement of Plant Drought Tolerance

Plants may cope with water scarcity through different physiological strategies; nevertheless, drought is among the most destructive abiotic stresses, since its multidimensional action affects the whole plant levels. Furthermore, most of the approaches suggested for controlling the negative effects of drought stress in plants, such as breeding for tolerant varieties and genetic engineering, are time-consuming and cost-intensive. In this scenario, plant-associated microbes with the capacity to improve drought tolerance in crop plants have received considerable attention in the last years. It has been reported that mycorrhizal fungi and PGPR recruited by the plant in the rhizosphere can contribute to alleviate abiotic stresses of the host improving plant physiological processes associated with drought resistance. Many research works characterized the activity of isolated microbial strains, identifying several mechanisms putatively responsible of the promotion activity, briefly overviewed in the following paragraph (see recent reviews by Ngumbi and Kloepper 2016; Soussi et al. 2016; Kumar and Verma 2018). Several bacterial strains have been isolated from plant rhizosphere and characterized in vivo for their capacity to improve drought tolerance to a variety of plants. Some PGPR strains were reported to be able to promote root growth and to alter root architecture, with the consequences to increase the root surface area, in turn leading to improve water and nutrient uptake. PGPR demonstrated, moreover, the capacity to promote also shoot growth, hence potentially decreasing the yield losses due to the plant response to drought stress, which inhibits shoot growth aiming to

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decrease water loss by evapotranspiration. Other PGPR drought protection mechanisms involve a strict plant-bacteria relationship, inducing plant cells to increase the content of osmoprotectant compounds (e.g., proline, free amino acids, sugars) or ROS (reactive oxygen species)-scavenging enzymes, both conferring higher tolerance toward drought-induced damages. The most studied and widespread mechanisms hypothesized to explain PGPR activity in increasing drought tolerance in plants are, nevertheless, the control of plant growth regulators. Lowering the concentration of ethylene, which is overproduced in response to stressful conditions, by 1-aminocyclopropane-1-carboxylate (ACC) deaminase, is considered to be one of the major mechanisms employed by PGP bacteria to favor plant growth under stress conditions. Other metabolites produced by PGP bacteria and potentially involved in supporting plant growth under stress conditions include phytohormones that modulate root development (i.e., indole-3-acetic acid), osmolytes that contribute to reduce cell dehydration, siderophores, and volatile compounds (Vurukonda et al. 2016). Additional mechanisms of PGPR growth promotion are directed to alleviate other environmental challenges that the plant is experiencing in dry lands. The production of exopolysaccharides (EPS), which protect roots from mechanical stress determined by dry soil compactness, can indeed play an important role under water deficit. Salt stress is, moreover, strictly related to drought, since high salt concentrations in the soil are very frequent in arid and semiarid regions. Salinity stress is one of the most common abiotic stress factors in modern agriculture, and PGP microbes can have several direct and indirect mechanisms of growth promotion (reviewed by Kumar and Verma 2018).

7.3.2

Approaches, Techniques, and Results

7.3.2.1 PGPR Isolation Sources Several recent ecological studies have found that microbial symbionts can confer habitat-specific stress tolerance to host plants, suggesting that the basis for the stress tolerance-enhancing effects of microbial symbionts is the coevolution of plant and microbes under harsh environmental conditions. Thus, it is a good strategy to look for plant-beneficial microorganisms that confer resistance to a specific environmental stress from the environments where that stress is a regular phenomenon. Water availability and drought gradients in soil were described to shape the structure of plant-associated bacterial communities. It was observed that the microbial diversity decreases along with water shortage and the relative abundance of the different bacteria change: under arid conditions, a higher proportion of Gram-positive bacteria, especially represented by the phylum Actinobacteria and the genus Bacillus, was observed by metataxonomic analyses (PCR fingerprinting and 16S rRNA sequencing) (Kavamura et al. 2013; Köberl et al. 2011). Recent advances in sequencing technologies and metagenomics have enabled us to explore and compare the microbial diversity that is associated with extreme environments and gain insight into how microorganisms survive in these harsh ecological niches (Lebre et al. 2017). Bacteria that colonize the rhizosphere of plants in dry environments may have undergone a selective pressure in order to survive. In addition, these bacteria may confer a certain

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level of tolerance to plants, as well as other functions such as plant growth promotion and soil maintenance due to their function and strategy in the ecosystem. Marasco and co-workers (2012) showed a variation in the distribution of bacteria into the endosphere and rhizosphere compared to uncultivated soil associated with Capsicum annuum plants under desert farming conditions, indicating that the enrichment of specific bacterial taxa is given by the plant itself. Moreover, most of the bacteria (88%) isolated from the root system of pepper plants exhibited multiple PGP activities and stress resistance capabilities, indicating that they can be active and hence express their PGP features in vivo under water stress conditions. Likewise, bacterial communities isolated from Salicornia plants grown under hypersaline ecosystems in Tunisia showed resistance to high-temperature, osmotic, and saline stresses and were able to perform different plant growth-promoting activities (Mapelli et al. 2013). Plants adapted to extreme environmental features, thus, represent ideal sites for discovering novel biotechnological agents which could be exploited to sustain other plants, including agricultural crops, to counteract water shortage in arid land agriculture.

7.3.2.2 Autochthonous or Allochthonous PGPR? One main question is if a PGPR strain isolated from a plant species would be equally efficient in promoting the growth when inoculated to a different species. This issue has important practical and economic consequences related to the formulation of microbial products to be applied on specific or broad range of crop plants. Different research works showed the cross-compatibility of PGPR strains, allowing their exploitation as biofertilizers tailored for arid lands. Rhodococcus sp., a Grampositive bacterium isolated from the rhizosphere of olive tree growing in south Tunisia, was able to promote root fresh biomass of tomato plants (Marasco et al. 2013); rhizospheric and endophytic Pseudomonas sp. isolated from the native halotolerant coastal plant Suaeda salsa were observed to be responsible for strengthening growth and salt stress responses of rice plants (Yuan et al. 2016); Pseudomonas putida MTCC5279 isolated from the desert regions of Rajasthan was able to increase drought resistance in chickpea (Tiwari et al. 2016). These evidences lead to consider that the nature of the plant-bacteria interaction in dry environments can have a limited level of specificity and that PGPR isolates may determine resistance to water stress in plants other than the one of the original isolation, increasing their application potential. It is, therefore, apparent that in relation to plant resistance to water stress, a feature of primary evolutionary importance for all plants, a crosscompatibility between PGPR and different plant models exists at least on a short term (Marasco et al. 2013). On the contrary, when PGPR isolates were applied on the same plant species but growing in different environmental contexts, the results were contradictory, and in some cases the strains showed different levels of promotion efficacy. Some strains were reported to promote plant growth under both irrigated and drought conditions (Gagné-Bourque et al. 2016), while others were specifically able to enhance plant growth only under stress and resulted ineffective in plant growth promotion under optimal irrigation conditions (Rolli et al. 2014; Chen et al. 2017). These findings

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suggest that the promotion activity of bacteria may be either stress-dependent or stress-independent.

7.3.2.3 In Vivo and In Vitro Testing of Drought Tolerance Promotion Isolated bacterial strains can be tested in vitro for traits that could be associated to plant growth promotion (PGP) (Marasco et al. 2012). Plate and tube assays are available and generally adopted to select the strains having PGP potential (Table 7.1): the production of hormones (the auxin indole acetic acid), ACC deaminase enzyme, bioavailable nutrients (ammonia, soluble phosphate, iron chelating siderophores), and other compounds (EPS) is routinely tested by most of the authors. Besides potential PGP traits, the strains could also be characterized in vitro for the capacity to tolerate abiotic conditions typical of arid soils, i.e., water and osmotic stress, high salt concentrations, and temperature extremes (Table 7.1). In vitro tests are easy to be performed and are useful to select the most promising strains among large collections of isolates. The PGP effect is nevertheless the result of a strict relation between bacteria and plant; therefore other complex factors drive the efficacy of a strain. In vivo experiments under drought conditions are, for this reason, required. According to in vitro PGP tests and the ability of the isolates to cope with several abiotic stresses (Table 7.1), the most promising strains should be further tested in pot experiments for their potential in growth promotion and plant drought tolerance improvement. Using surface-sterilized seeds, an established concentration of bacteria is supplied to plants by seed biopriming or by liquid inocula provided to seedlings. During the period of growth under controlled conditions, water-deficit stress is Table 7.1 In vitro screening of plant growth-promoting activities and stress resistance capabilities In vitro screening of plant growth-promoting activities Production of indole-3-acetic acid Solubilize insoluble phosphate compounds Ammonia synthesis Protease activity ACC deaminase activity Atmospheric nitrogen fixation ability Production of exopolysaccharides Siderophore production In vitro screening of stress resistance capabilities Resistance to salt

Resistance to high temperature

Tolerance to osmotic stress

Methods Bric et al. (1991) Ahmad et al. (2008) Cappuccino and Sherman (1992) Nielsen and Sørensen (1997) Belimov et al. (2005) Penrose and Glick (2003) Santaella et al. (2008) Schwyn and Neilands (1987) Methods Adding 5, 8, and 10% sodium chloride to culture media and incubating the plate at 30  C for 7 days Verifying growth at 4, 42, and 50  C in solid media placed in incubators for 7 days Adding 10–20% polyethylene glycol (PEG) to the original liquid media

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imposed to plants withholding water for a certain time (from 7 to 18 days depending on plant variety) or maintaining soil moisture at constant low levels (Zolla et al. 2013). When plants are harvested, different parameters are measured to evaluate the potential of PGP bacteria to improve drought tolerance in plants. In particular, the parameters to take into account are (i) vegetative parameters including shoot and root lengths, shoot and root dry weight, and number of leaves and nodes and (ii) productive parameters like fruits yield per plant and fruits quality. In order to evaluate how the inoculated strain promoted plant growth and alleviate water stress, it is also possible to measure leaf physiological parameters, possibly using not destructive methods, including net photosynthesis, evapotranspiration, and stomatal conductance. Chen et al. (2017) for instance, measured free proline, soluble sugars, malondialdehyde (MDA), and total chlorophyll from leaves of wheat plants inoculated with the PGP strain LTYR-11ZT. The proline concentration in leaves, which usually increases drastically under drought stress, was reduced with the inoculation of the PGP strain: this decrease could be indicative of a less damage in wheat plants in the presence of the bacterium. Similarly, wheat plants colonized with LTYR-11ZT strain had significantly lower MDA contents compared with non-inoculated control under drought stress: MDA level can reflect the degree of cell membrane damage, so the bacteria inoculation can help in overcoming this kind of damage. About soluble sugars, which drastically increase under water-deficit conditions, it is known that they play a role in drought tolerance maintaining cell osmotic turgor: the presence of the PGP bacterium further increased the concentration of sugars allowing better osmotic adjustment in the host plants. Finally, chlorophyll concentration is considered as an important indicator of drought tolerance in plants: this study revealed that chlorophyll level was increased in treated plants under stress conditions, suggesting that the PGP bacterium helped plants adapt to drought stress. Another approach was used by Gagné-Bourque and co-workers (2016) with Brachypodium plants inoculated with Bacillus subtilis strain: they set up a DNA methylation assay, and they showed an increase in the abundance of methyl transferases involved in the maintenance and regulation of DNA methylation, suggesting that this bacterium could potentially act at the epigenetic level to increase drought stress tolerance in plants.

7.3.2.4 Study of Plant-Bacteria Relationship To understand and compare the “holistic” responses of the bacteria and plant system under water-deficit conditions, a powerful tool is the study of their metatranscriptome, consisting of the entire set of transcripts that are expressed with in a meta-organism in a particular developmental stage or under different environmental conditions. Using quantitative real-time PCR, upregulation of drought-response genes DREB2B-like, DHN3-like, and LEA-14-A-like in the aerial parts of Brachypodium was identified when plants were inoculated with a PGPR Bacillus subtilis strain (Gagné-Bourque et al. 2016). Using microarray analysis, a set of drought signaling response genes were discovered to be downregulated under drought stress in Arabidopsis thaliana plants colonized by a PGPR Pseudomonas chlororaphis strains compared to non-inoculated plants

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(Cho et al. 2013). Yuan et al. (2016), using PICRUSt (Phylogenetic Investigation of Communities by Reconstruction of Unobserved States) tool for predicting S. salsa-associated microbiome functions under salinity stress, discovered an enrichment of diverse two-component systems and ATP-binding cassette transporters, which may provide a selective advantage for bacterial adaptation to adverse conditions. More recent in situ applications of “omics” technologies provided key insights into the responses of xerotolerant microorganisms in their natural xeric environments. The microarray-based GeoChip technology, which encompasses an array of functional genes that are involved in metabolism and stress responses, has been efficaciously applied to directly measure xeric stress responses (Chan et al. 2013). A critical step in the interaction between beneficial bacteria and the host plants is the efficient root colonization by inoculated bacteria. To play an effective role in plant growth promotion, a bacterial strain must be able to respond to the plant rhizodepositions and colonize its root apparatus entering in association/competition with the resident rhizosphere microbial community. The potential ability of PGP isolates to efficiently colonize plant root system can be tested in vivo by performing an adhesion assay. To evaluate the colonization pattern of the test PGP strain, one of the most exploited strategies is to manipulate it in order to obtain a gfp-tagged strain. To have the possibility to perform long-term adhesion test, the tagging should be chromosomal rather than plasmidic, in order to minimize the appearance of not fluorescent revertants. Plant roots are exposed for a period of time (i.e., 16 h) to the bacterial suspension of the fluorescent mutant and then analyzed by confocal microscopy (Mapelli et al. 2013; Chen et al. 2017). For strains reluctant to genetic manipulation, a culture-dependent monitoring of colonization could be performed, but the colonization in this case can result overestimated due to the presence of a cultivable endogenous plant microbiota difficult to be distinguished from the inoculated PGP strain. Inoculants could be, in alternative, quantified in the rhizosphere metagenome by quantitative real-time PCR (qPCR) assay, basing on sequence signatures previously identified in the strain genome (Gagné-Bourque et al. 2016). The abovementioned methods could also be useful to detect the vertical transmission of the PGP strains initially recruited from the rhizosphere, evaluating their presence into the seeds and in the offspring tissues. Vertical transmission of PGPR to the plant progeny is very interesting as it enables a plant with an established endophytic community to pass bacteria with beneficial characteristic to the next generation and ensures the presence of a beneficial microbiota at an early stage of seedling growth. Bacteria having the potential to establish as endophytes have other qualities that make them of particular interest, like their advantage of being relatively protected from the competitive high-stress environment of the soil, since they live in the interior parts of the plants. As compared to rhizospheric bacteria, endophytic inocula showed positive results in the promotion of plant growth and stress tolerance induction in plant under harsh environmental condition (Naveed et al. 2014); however, it has not been resolved whether plants benefit more from an endophyte than from a rhizospheric bacterium. It was

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proved that endophytes might exhibit phytopathogenic effects under certain conditions, and rhizosphere bacteria might also be able to colonize the internal parts of the plant (Hardoim et al. 2015). This is because microbial population dynamics are affected by a variety of factors, such as plant species, soil type, and biotic/abiotic factors.

7.3.3

Discussion

7.3.3.1 Scaling Up from Laboratory to the Field: The Importance of the Residing Soil Microbiota Many factors, known and unknown, influence the interaction between plants, soil, and microbiota. The relationship is complex and dynamic, and this is the reason why bacterial inoculants often perform well in controlled laboratory experiments and then fail to give a beneficial effect in natural agricultural settings. One possible explanation is that laboratory experiments are often conducted using sterilized soil substrates, while the microbial inoculants face competition with the native soil microbiota when inoculated to natural soil. Plant inoculation with selected bacterial strains in natural soil conditions could fail to achieve expected outcomes because of the competition with native microbial communities and the limited colonization efficiency. Moreover, it is well known that the behavior of microorganisms as pure cultures could be different from their behavior in a microbial community. It has been hypothesized that it is better to leverage on the natural community rather than trying to transplant microorganisms (East 2013). Zolla et al. (2013) unraveled indeed the importance of the soil microbiome as a whole in alleviating drought stress, against the convention of a single bacterial application, demonstrating the ability of a sympatric soil microbiome to increase Arabidopsis growth under water-deficit conditions. 7.3.3.2 Synthetic PGPR Communities Many different PGPR strains have been characterized up to now, discovering very different mechanisms of plant growth promotion and protection toward drought. A possible approach to exploit this high functional diversity is to inoculate plants with a mixture of strains having different putative beneficial traits. The combination on inoculants may not necessarily produce additive or synergistic effects but could, instead, generate a competitive situation in the rhizosphere without benefitting the plant. To better understand this complexity, a novel strategy was recently proposed to reconstruct and transfer into the rhizosphere the functional microbial groups instead of individual isolates. Synthetic communities are indeed a powerful tool to investigate fundamental principles in natural systems as they are more similar to natural conditions compared to pure cultures, but at the same time they have a reduced complexity and higher controllability. Niu and co-workers (2017) inoculated maize axenic plants with seven strains representing three of the four most dominant phyla found in maize roots and by tracked by a selective culturing method the abundance of each strain during root colonization in maize seedlings.

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This method allowed to study the dynamics and function of root bacterial assemblages, leading to identify key species necessary for plant protection.

7.4

Case Study 2: Rhizoremediation of PCB Polluted Soil

Polychlorinated biphenyls (PCBs) are a class of xenobiotic organic compounds which were massively produced for a variety of industrial uses (as dielectric fluid in capacitors and transformers) and to a lesser extent in construction materials (such as paints, caulking, and lighting ballasts) in many countries. The production lasted for many decades until it was banned by the 1980s because of increasing concerns about their environmental and human health risks. PCBs are highly persistent in the environment and tend to accumulate in the food chain, posing adverse effects to human health and the environment also due to their chronic toxicity (IARC 2015). Although measures were taken to reduce their emissions, resulting in a substantial decrease of their concentrations, PCB are still present in the environment and can be released from reservoir compartments including contaminated soil. Nowadays, several remediation solutions for PCB-contaminated soil are available based on physical methods and thermal and chemical treatments (Gomes et al. 2013); however, a greater interest is increasingly addressed to bioremediation technologies, such as phyto-rhizoremediation, representing a sustainable alternative to more expensive and disruptive traditional remediation options. Phyto-rhizoremediation involves two main actors (plants and microorganisms) and encompasses a set of processes such as contaminant root uptake from soil and translocation to steam and leaves (phytoextraction), plant enzymatic transformation (phytodegradation), and rhizoremediation (van Aken et al. 2010; Gomes et al. 2013; Passatore et al. 2014). Rhizoremediation is based on the plant enhancement of the microbial degrading activity in the root zone (rhizosphere), and it is the most important removal process for hydrophobic organic chemicals such as PCBs (Vergani et al. 2017). Many studies with different degrees of complexity have been conducted to investigate the potential of plant-microbe interactions in the remediation of PCB-contaminated soils. Since an exhaustive review of current rhizoremediation publications is beyond the scope of this book chapter, the main objective will be to (1) outline the main experimental and modeling approaches used to investigate and predict the role of rhizoremediation in terms of contaminant concentration reduction in soil; (2) describe different type of techniques involved in rhizoremediation experiments including sampling, analytical, and microbiological techniques; (3) summarize the main results of rhizoremediation experiments in term of PCB half-lives; (4) discuss the main pitfalls of rhizoremediation experiments performed so far; and (5) suggest some guidelines to carry out experiments in order to provide more accurate, comparable, and useful data.

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Experimental Approaches and Techniques

7.4.1.1 Type of Experiments Different types of rhizoremediation experiments are available in the literature. They range from short-term laboratory studies, including in vitro (Kučerová et al. 2000; Dudášová et al. 2014) and hydroponic (Chu et al. 2006) tests, growth chamber (Mehmannavaz et al. 2002; Chekol et al. 2004), and microcosm experiments (Kurzawova et al. 2012; Meggo and Schnorr 2013; Liang et al. 2015) to midterm greenhouses (Dzantor et al. 2000; Javorská et al. 2009; Shen et al. 2009; Li et al. 2011) and long-term field trials (Mackova et al. 2009; Teng et al. 2010; Tu et al. 2011; Ancona et al. 2017), including forensic field studies (Leigh et al. 2006). A complete list of PCBs rhizoremediation experiments performed so far can be found in the following reviews: van Aken et al. (2010), Gomes et al. (2013), Passatore et al. (2014), Vergani et al. (2017), and Terzaghi et al. (2018). Laboratory and greenhouse experiments generally investigated the role of plantmicroorganism interactions in reducing PCB concentrations in spiked soil or soil collected from contaminated sites. These experiments allowed to study rhizoremediation (1) under controlled conditions, i.e., independent from seasonal meteorological variability and nutrient and water availability, and (2) in a more homogeneous soil in terms of contamination type and levels (well mixed) and characteristics (texture, organic carbon content, etc.). However, soils at contaminated sites may be characterized by scarcity of nutrients and lack of microbial diversity, together with a high spatial heterogeneity of contaminant concentrations and soil properties; moreover, they often contain mixtures of organic and inorganic chemicals at concentrations whose range can be phytotoxic and inhibit plant development (root growth); in addition, weathered PCBs are often less bioavailable, especially compared to experiments in which soils are spiked with fresh chemicals; finally, laboratory and greenhouse experiments are usually conducted for a short period of time and maximizing the root biomass/soil weight ratio, which in field could be achieved after many years of root exploration. These conditions, together with other plant stressors and physical restrictions, i.e., temperature variability and diverse water availability in fields, plant pathogens, herbivores, and weed species, as well as when the depth of contamination is higher than the rooting zone, may lead to significantly different results from those obtained using a promising plant species (selected from laboratory or greenhouse experiments) in a field-level remediation trial (Gerhardt et al. 2009). This highlights the need to perform more realistic laboratory and greenhouse experiments and to translate and validate their results in field (Schwitzguébel 2017). Some efforts were also devoted to assess directly in field the rhizoremediation capability of native species (forensic field studies) already adapted to grow in contaminated soil and under environmental stress; however, the high heterogeneity of soil and the absence of controlled conditions could be relevant confounding factors in determining the true rhizoremediation effect.

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7.4.1.2 Experimental Design The success and robustness of a rhizoremediation experiment imply a carefully planned experimental design, i.e., the definition of (1) the number and type of treatments, (2) the type of controls, (3) the number of replicates, and (4) the duration of the trials. A single treatment (planted) is generally considered for field studies (Whitfield Åslund et al. 2007; Ficko et al. 2010, 2011a), except for few studies which evaluate two levels of contamination (Kacálková and Tlustoš 2011). On the contrary, in laboratory and greenhouse experiments, a higher number of treatments can be compared in parallel, for example: (1) different plant species (Dzantor et al. 2000); (2) two plant species individually grown or in consociation (Li et al. 2013); (3) one plant species added or not added with bacteria (Mehmannavaz et al. 2002; Teng et al. 2010; Xu et al. 2010), fungi (Teng et al. 2010; Lu et al. 2014; Qin et al. 2014), earthworms (Lu et al. 2014), or amendments (Dzantor et al. 2000; Shen et al. 2009); (4) one plant species grown in a single or mixed congener contaminated soil (Meggo and Schnoor 2013); and (5) undergoing different cycles of flooding and draining conditions of soil (Chen et al. 2014). Every experimental design should include adequate controls such as an unplanted control, to understand the importance of volatilization, infiltration, (naturally growing) plant uptake, and baseline biodegradation, in reducing chemical concentration in soil; a sterile control (whenever possible), to differentiate the potential of baseline biodegradation from volatilization; an unplanted control + everything else which is added to the soil (bacteria, fungi, earthworms, amendments, nutrients, etc.) to evaluate their exclusive effects on concentration reduction; and a planted control with clean soil, to monitor cross-contamination (especially in microcosm/greenhouse conditions). While the unplanted control is generally set up, only few studies considered sterile (Li et al. 2011; Slater et al. 2011) and not contaminated control (Meggo and Schnoor 2013). Chemical concentration in controls and treatments needs to be monitored simultaneously to evaluate if a statistically significant difference exists. Moreover, the same fertilization, irradiation, irrigation conditions, as well as pest and weed control precautions must be applied to both treatments and controls. However, water regime needs to be carefully adjusted in the different treatments and controls by ensuring a similar moisture level rather than by dispensing an equal volume of water to avoid the occurrence of saturated and anoxic conditions which may differ among the treatments. For each treatment and control, a suitable number of replicates should be set up to allow a statistical analysis of the results, and in general a high enough number of replicates (3–5) should ensure more accurate results; moreover, a randomized arrangement of pot and plot replicates should be chosen to guarantee that differences in soil concentrations are due to treatment instead of soil characteristics, moisture, and light exposure dependent of different positions in a greenhouse or in field. Additional information can be found in books of statistical methods for biological experiment (Sparks 2000). Experiments are generally performed in triplicates except for some studies that consider a lower (Mehmannavaz et al. 2002) or higher number of replicates (Teng et al. 2010; Tu et al. 2011); in some works, the random collection

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of soil sample in the same replicate considered allows only the evaluation of intrapot variability (Liang et al. 2014). Moreover, in some papers the number of biological replicates is not explicitly declared, but just the number of analytical replicates is shown (Ancona et al. 2017). Experiment duration can range from few days for in vitro and hydroponic test to up to 270 days for growth chamber and microcosm experiments (Mehmannavaz et al. 2002); greenhouse experiments can last from 60 to 180 days (Li et al. 2011; Lu et al. 2014) while field trial from 90 to 730 days (Xu et al. 2010; Tu et al. 2011). Experimental time, especially for field trials, should be properly chosen to observe significant results. Some studies have demonstrated that a decrease in soil concentration is generally only observed after many consecutive vegetative seasons because the high heterogeneity of soil could mask small but potentially significant changes in soil concentrations obtained after only one or two growing seasons (Ficko et al. 2011a, b). The experiment protocol should also include a detailed description of fertilization, irradiation, irrigation, and pest/weed control conditions which depends on plant species, environmental conditions and growth chamber, microcosm, pots, and field plots characteristics. For more information, refer to specific paper.

7.4.1.3 Selection of Plant Species Plants have a limited ability to uptake PCB from soil and metabolize them efficiently, except for few species of Cucurbitaceae family (zucchini, pumpkin) (Hülster et al. 1994; Whitfield Åslund et al. 2007, 2008) and some oil-containing plant such as carrots (Topp et al. 1986), highlighting that neither phytoextraction nor phytodegradation can be considered relevant processes in the phytoremediation of PCBs. On the contrary, plant-microbe beneficial interactions play an important role in influencing PCB degradation in rhizosphere soil (rhizoremediation) because of (1) bacterial plant growth-promoting activity, (2) stimulation of microbial growth and PCB-degrading bacteria activity through the release of plant secondary metabolites, (3) increase of PCB bioavailability by root exudates that have bio-surfactant properties, and (4) presence among root exudates of compounds which sustain PCB degradation acting as co-metabolites (Vergani et al. 2017). There is not a single plant species generally recognized and used for the remediation of contaminated soil, but a long list of plant species, including both crop and spontaneous species (herbaceous) as well as shrubs and woody ones, has been tested in PCB phytoremediation experiments. The most employed herbaceous plant species belong to genera of Medicago (M. sativa, M. polymorpha), Panicum (P. virgatum, P. variegatum, P. clandestinum), Festuca (F. arundinacea), Phalaris (P. arundinacea), Cucurbita (C. pepo, C. pepo spp. Pepo, C. maxima, C. moschata), Brassica (B. napus, B. nigra), Nicotiana (N. tabacum), and Solanum (S. nigrum), while considering the woody ones, these are genera of Salix (S. caprea, S. alaxensis) and Populus (P. deltoids x nigra DN34, P. generosa x P. nigra). Other species can be found in Gomes et al. 2013; Vergani et al. 2017; Terzaghi et al. 2018. The selection of plant species is a key point in the success of phytoremediation procedures. Their ability in enhancing PCB degradation in soil has to be supported

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by other characteristics such as (1) tolerance to high concentrations of contaminants (including co-contaminants), (2) resistance to scarcity of nutrients, and (3) fast growth, especially high root biomass production. Native plant species should be preferred, being already well adapted to climatic conditions and soil characteristics of the site to be remediated, as well as to other stressors such as water availability and pests (Gerhardt et al. 2009); moreover, they eliminate ecological risks due to the introduction of a non-native species to an ecosystem.

7.4.1.4 Techniques Rhizoremediation experiments comprise different techniques, including sampling, analytical, and microbiological techniques. Besides microbiological techniques, already mentioned in the first case study, an overview of sampling and analytical techniques generally adopted is reported below. Sampling strategies encompass the selection of appropriate sampling methods and the definition of compartments to be sampled. Soil collected in a contaminated site or manually spiked tends to be heterogeneous; therefore, soil homogenization techniques must be adopted to reduce concentration variability and ensure similar initial conditions in all pots/reactors at the beginning of the experiment or to analyze a sample that is representative of the whole pot/reactor concentration. Typically adopted homogenization techniques range from simple quartering techniques (Meggo and Schnoor 2013; Liang et al. 2014) to more complex procedure such as “one-dimensional Japanese slab-cake” (Low et al. 2010) or other incremental sampling techniques (ITRC 2012). The collected samples may include above- and belowground plant biomass (leaves, stems, and roots), in addition to bulk soil and/or rhizosphere soil, i.e., the soil firmly adhering to the roots (Xu et al. 2010). Different types of containers are used for samples collected for chemical and microbiological analysis: while sterile plastic bags are suitable for microbiological analysis, glass jars must be used for chemical analysis to reduce chemical adsorption on container walls and/or contamination by plasticizer used in the plastic polymer, e.g., phthalates. After sampling, samples should be quickly transported on ice, to minimize microbial activity, to laboratory and stored until analyses at 4  C (for direct microbiological analysis) or 20  C (for chemical or molecular analysis). Appropriate field, transport, and equipment blanks should be collected to evaluate the analytical quality (cross-contamination) of sample collection and transport (EPA 2011). To evaluate the efficacy of the phytoremediation strategy, sampling of soil and plant materials should be carried at different times of the trial to evaluate the significance of concentration changes. Different analytical techniques are used to determine PCB concentration reduction in soil, and accumulation in plant. PCB extraction is performed with ultrasonic bath (Liang et al. 2014), Soxtec extractor (Li et al. 2013), or accelerated solvent extractor (Dzantor et al. 2000) using different solvent mixtures, e.g., hexane/acetone 1/1/, while sample purification is achieved by means of sulfuric acid (Meggo and Schnoor 2013), Florisil (Li et al. 2013), or silica gel (Liang et al. 2014). PCB quantification is carried out by GC-ECD (Chekol et al. 2004; Slater et al. 2011), GC-MS, or GC-MS/MS (Lu et al. 2014; Liang et al. 2014) with internal or the less

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accurate external standard calibration (Teng et al. 2010; Tu et al. 2011). Laboratory blanks should also be analyzed to assess the performance of the analytical procedure. A number of reference texts on analytical chemistry for PCBs are available to provide guidance (Erickson 1997; O’Sullivan and Sandau 2014).

7.4.2

Modeling Approaches

In the last two decades, three main types of multimedia fate models were developed to predict the chemical fate in the environment including also the contribution of the plants: (1) “Forest models” – they do not usually include root and stem compartments since they are mainly applied to organic compounds such as PCBs for which root uptake and translocation into stem are not important (Log KOW (octanol-water partition coefficient) > (3) and they have as main goal estimation of forest filter effect and air-canopy exchanges (Wania and McLachlan 2001; Nizzetto and Perlinger 2012). (2) “Crop models” – they include also fruit compartment (in addition to roots, stems, and leaves), and they aim to estimate the plant concentration after the application of a pesticide and/or the calculation of human intake of hydrophobic compounds and their metabolites taken up from soil by plants roots (Legind et al. 2011; Fantke et al. 2011; Trapp 2015). (3) “Phytoremediation models” – they are mainly applied to more polar compounds and metals, rather than to hydrophobic compounds, and they were developed to simulate chemical plant uptake, phytovolatilization, and transport in the vadose zone and aquifers (Ouyang 2002, 2008; Manzoni et al. 2011; Canales-Pastrana and Paredes 2013). These models generally contain a simplified soil compartment parameterization and do not include, among the chemical loss fluxes from soil, the enhanced biodegradation process due to the plant-microbe interactions; moreover, the existing phytoremediation models generally require a large amount of inputs (i.e., plant physiological parameters) that are often difficult to obtain through measurements and increase the model complexity (Canales-Pastrana and Paredes 2013). Recently, a full dynamic model (SoilPlusVeg model) based on fugacity was developed to predict the environmental fate of organic chemicals in an air-vegetation-litter-soil system (Terzaghi et al. 2017). In this model, the soil compartment is subdivided into several 5-mm-thick layers to better reconstruct the chemical fate through the soil depth, and it is parameterized considering different parameters such as organic carbon (OC) content, seasonal dissolved organic carbon (DOC) fluxes, organic matter (OM) degradation rate, texture, initial water conditions, etc. Moreover, the chemical loss fluxes (volatilization, runoff, infiltration, diffusion, root uptake, and biodegradation) could also account for the enhanced biodegradation process due to plant-microbe interaction considering literature-

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derived rhizoremediation half-life (see sect. 7.4.3) and their dependence on soil temperature and water content (Walker 1974). Such a type of model could be a powerful tool to assess the phytoremediation performance in a contaminated site and estimate the time requested to achieve regulatory thresholds, considering all loss processes that an organic chemical can undergo. It could also help in evaluating the performance of phytoremediation under different scenarios (changing climate, water availability, etc.), considering both spatial and temporal variability of those parameters that may affect chemical biodegradation and therefore predict the fluctuations of the degradative potential of the soil microbial community over the whole year.

7.4.3

Laboratory, Greenhouse, and Field Experiment Results: Rhizoremediation Half-Lives

Rhizoremediation experiment performed in the past decades produced many encouraging and promising results, although some unsuccessful attempts were obtained especially at a field-scale (Whitfield Åslund et al. 2007; Ficko et al. 2010). However, the failure of phytoremediation in field was critically evaluated by Gerhardt et al. 2009 to avoid the spread of negative opinions that could affect this promising remediation strategy. The efficacy of a rhizoremediation treatment can be expressed by the calculation of the chemical half-life, i.e., the time needed for a contaminant concentration to decrease by half compared to its initial concentration. This calculation requires the knowledge of chemical concentration in the control at the beginning of the experiment and in both the control and the treatment at the end of the experiment, to differentiate volatilization, infiltration, and baseline degradation from plant uptake and enhanced biodegradation (Terzaghi et al. 2018). Recently, an attempt to estimate half-lives from rhizoremediation experiments, for now on called rhizoremediation half-lives (HLrhizo) for PCB, was made (Terzaghi et al. 2018) carefully selecting from the literature some properly conducted studies and considering a first-order degradation kinetic (Fig. 7.1). A great number of studies found in the literature were discarded because their experimental design or methods and techniques adopted do not provide usable data (see sect. 7.4.4.1). The obtained HLrhizo ranged from few months (~ 2) for mono-PCBs to some years (~ 12) for deca-PCBs and were up to a factor of 5 lower than generic PCB halflives (HLgen) reported in the literature (not derived from rhizoremediation experiments) (Paasivirta and Sinkkonen 2009) highlighting the importance of plant-microbe interaction in improving PCB degradation in soil. The calculated HLrhizo represent average values considering studies conducted with different experimental approaches (laboratory, greenhouse, and field), plant species, conditions, and techniques; therefore, their variability (RSD, relative standard deviation) could range from 17% for mono-PCBs to 91% for penta-PCBs. Moreover, while many values (up to ~ 40) were available for low chlorinated PCBs, especially tri-, tetra-, and penta-PCBs, data about higher chlorinated congeners were totally

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missing in the literature, and therefore their HLrhizo were tentatively assumed as half the corresponding HLgen (see Terzaghi et al. 2018 for more details). However, some greenhouse experiments were recently performed to investigate the rhizoremediation of higher chlorinated PCBs using a soil contaminated by different PCB mixture, including Fenclor DK (a technical grade deca-PCB (PCB 209) mixture), collected in a National Priority Site (SIN) locate in Northern Italy (Di Guardo et al. 2017). Once the results will be available, a true HLrhizo (not assumed) may be calculated also for high chlorinated congeners. Although some uncertainties due to the variety of the experimental protocols of the different studies are considered, the HLrhizo values proposed by Terzaghi et al. (2018) could be used as selected input parameters in multimedia fate models (see Sect. 7.4.2) that aim to predict PCB fate in a contaminated soil and the time requested to achieve regulatory threshold in a PCB-contaminated site, allowing to draw up its remediation plan.

7.4.4

Discussion

7.4.4.1 Problems and Pitfalls of Current Rhizoremediation Experiments Rhizoremediation experiments could provide useful data, i.e., the rhizoremediation half-lives, to be used in PCB fate assessment in a contaminated soil subjected to a rhizoremediation treatment. However, often the data produced, given the variety of

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approaches, techniques, and constraints, need a careful assessment in order to provide comparable, accurate, and useful data for fate estimation. Recently, Terzaghi et al. (2018) highlighted potential weaknesses and limitations of the rhizoremediation experiments performed so far, including different aspects of the experimental design, sampling techniques, and analytical and microbiological techniques such as (1) the type of the tested chemicals (single congeners vs. mixtures), (2) the type of contamination (spiked soil vs. aged), (3) the type of the experimental facility (laboratory, greenhouse, field), (4) the duration, (5) the setup of appropriate controls, (6) the planning of an adequate number of biological replicates, (7) the adoption of soil homogenization techniques, (8) the employment of suitable analytical techniques, and (9) the evaluation of the “degrading potential” of the soil and its enrichment during the experiment. Since each of these aspects can have different consequences on the calculation of HLrhizo (over- or underestimation), some guidance can be provided to assist researchers in the setup of suitable experiment to obtain comparable PCB half-lives. In the literature, few studies about single PCB congeners are available (Li et al. 2011; Meggo and Schnoor 2013, Tu et al. 2017), while many studies deal with mixtures (e.g., Aroclor, Delor, etc.) (Mehmannavaz et al. 2002; Chekol et al. 2004; Mackova et al. 2009; Qin et al. 2014). This reflects in an overestimation of HLrhizo for single congeners, especially the low chlorinated ones which could additionally derive from the degradation of higher chlorinated congeners (increasing loss of -Cl) that compose the mixtures. Moreover, the selected PCB mixtures are typically low chlorinated (Aroclor 1242, Delor 103, etc.) resulting in a lack of HLrhizo for high chlorinated PCBs (octa, nona, and deca). Rhizoremediation experiments (mainly laboratory and greenhouse trials) are often performed with fresh spiked soil (Dzantor et al. 2000; Chekol et al. 2004) rather than with aged contaminated soil leading to an underestimation of HLrhizo since spiked PCB are generally more bioavailable to PCB-degrading microorganisms. In addition, few field studies are conducted, and pot experiments (laboratory and greenhouse) do not generally reproduce field conditions (see Sect. 7.4.1.1) obtaining a not statistically significant concentration reduction between the unplanted control and the planted treatment in field experiments, when using a plant species that showed promising results in pots (Ficko et al. 2010; Whitfield Åslund et al. 2007). Furthermore, long-term studies are uncommon, although a significant decrease in PCB soil concentration could be appreciated after several plant growing cycles (Ficko et al. 2011a, b; Whitfield Åslund et al. 2007). Many studies did not consider all the appropriate controls (see Sect. 7.4.1.2) and did not provide measured initial control concentration, but they considered just nominal concentration (Chekol et al. 2004; Li et al. 2011) leading to an underestimation of HLrhizo. In other cases, concentration in control is not measured in parallel to concentration in treatments but just at the beginning of the experiment (Meggo and Schnoor 2013; Shen et al. 2009) allowing the calculation of an overall chemical half-life including volatilization, infiltration, plant uptake, and baseline biodegradation in addition to enhanced biodegradation due to plant-microbe interaction. Similarly, the number of biological replicates is often inadequate to interpret results with statistical methods (see Sect. 7.4.1.2) and guarantee robust data.

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Soil homogenization techniques (see Sect. 7.4.1.4) to reduce PCB concentration variability and therefore observed small changes in concentrations, are uncommon especially at the end of the experiment; sometimes only some subsamples which may be not representative of the whole pot (Meggo and Schnoor 2013; Mehmannavaz et al. 2002) are collected; some researchers try to separate bulk soil from rhizosphere soil (Xu et al. 2010; Ancona et al. 2017), although determining their boundary is very difficult, because in the rhizosphere plant-microbe interaction is stronger and higher degradation rates are expected; therefore, when calculating and comparing HLrhizo, the type of soil measured needs to be considered in order to avoid misunderstanding. Concerning the analytical techniques, single congener analysis should be performed with GC-MS systems and mass-labeled internal standard, instead of using GC-ECD and external standard calibration to obtain single PCB half-lives. PCB congener identification with GC-ECD is more difficult especially if a dual ECD is not used and external standard quantification does not guarantee accurate results. Moreover, other soil parameters are scarcely monitored (OC fraction, DOC concentration, texture, pH, temperature, and redox potential) although they may influence PCB concentration reduction. On the microbiological point of view, the evaluation of the presence and enrichment of degrading microorganism in soil is important to attribute concentration reduction to the concentration and diversity of degrading microorganisms rather than different remediation approaches. Molecular methods, especially based on metagenomics, represent a powerful tool to study the degrading potential of rhizosphere microbiota and discover new enzymes putatively involved in contaminant degradation. In the case of aerobic PCB degradation, however, few bacterial strains have been isolated having the capacity of PCB mineralization. As a consequence, there is a general lack of data needed for (i) the proper annotation of genes coding for new enzymes involved in PCB mineralization and (ii) the evaluation of the degrading potential of the rhizosphere microbiota through the quantification of marker genes in the soil metagenome (Vergani et al. 2017).

7.4.4.2 Overcoming Unsuccessful Rhizoremediation in Field One of the most important recent challenges in rhizoremediation is the translation of laboratory and greenhouse experiment results to field. The two main causes behind the unsuccessful results in field have been recently mentioned in the literature (Gerhardt et al. 2009): (1) the lack of plant stress factors (variations in temperature, nutrients, precipitations, pathogens attacks, and weed species competition) in the laboratory and greenhouse experiment with respect to field and (2) the inadequacy of techniques adopted to observe a significant contaminant concentration reduction. As for drought stress, PGPR constitute a promising tool to alleviate plant stress in contaminated soil. Research and exploitation of beneficial microbes could help to overcome the problems related to a poor development of plants in highly contaminated area, through a PGPR-enhanced phytoremediation. The finding and application of rhizobacteria that at the same time, in parallel to degrading ability, can

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also act in the biocontrol of pathogenic organisms, the increase in nutrients uptake by plants, or decrease the effect of environmental stress, are particularly promising (Narasimhan 2003). Due to the high heterogeneity of soil, reliable monitoring methods are required to detect small but significant decrease in concentrations. Evaluating rhizoremediation efficiency averaging many data points (composite sample) in field could mask concentration reduction at one point of the field plot; therefore, it could be useful to collect samples at the same point over time to follow its changes in concentrations (Gerhardt et al. 2009) in that point, in addition to the average across the field. Moreover, the acceptable level of significance for statistical analysis should be raised from 5% (P  0.05) to 10% (P  0.1), and longer time frame for field trials (almost 3 years) should be considered to avoid misleading results.

7.5

Conclusions and Outlook

The rhizosphere was demonstrated to constitute a hot spot for beneficial microorganisms, which can be exploited in agro-environmental biotechnologies to obtain key services, in the frame of the development of a bioeconomy-based society. Two different services have been presented, i.e., (1) the promotion of a sustainable agriculture practice in arid lands, thus contributing to face global warming challenges by improving drought tolerance and water use efficiency in crop plants, and (2) soil bioremediation from toxic organic ubiquitous pollutants like PCBs, contributing to restore contaminated lands and decrease human and ecological exposure to toxic agents. A large body of knowledge about microbiological, molecular, chemical, and modeling aspects of the presented approaches is available, especially at the laboratory and microcosm level. We have in culture many bacterial strains which exhibit a good potential to improve drought tolerance to plants or degrade organic pollutants. We need nevertheless a better understanding of what makes a beneficial PGPB and/or a degrading strain to interact in the rhizosphere with the plant and the residing microbiota in a multipartite ecological relationship. This would provide an important insight to better select and handle microbes. Profound knowledge is, moreover, needed on the survival and persistence of inoculated microbes in the field and on their effects on the native microbial communities. It is also necessary to understand the abiotic context dependency of inoculation or biostimulation approaches. The transfer of laboratory-based knowledge to the field is currently the bottleneck that we need to overcome to set up effective agro-environmental biotechnologies relying on the functional potential of rhizosphere microbiome. Acknowledgments We thank the EU Horizon 2020 MADFORWATER project (GA No. 688320, www.madforwater.eu) and the collaboration of the “Caffaro Working Group”: Stefano Armiraglio, Simone Anelli, Vanna M. Sale, and Paolo Nastasio and the funding agency Ente Regionale per i Servizi all’Agricoltura e alle Foreste (ERSAF). DSAT of University of Insubria is also acknowledged for funding part of ET’s salary.

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8

Methods for Detecting Biocontrol and Plant Growth-Promoting Traits in Rhizobacteria Gustavo Santoyo, Juan M. Sánchez-Yáñez, and Sergio de los Santos-Villalobos

Abstract

The field of agriculture requires new strategies to control the damage caused by phytopathogens and to effectively promote plant growth and production. Until recently, one of the best alternatives was the application of microorganisms exhibiting biocontrol and plant growth-promoting traits. Therefore, to select the best microorganisms, it is essential to analyse these beneficial activities based on reliable search methods and to ensure the greatest extent possible and their successful application in the field. In this chapter, we have summarized and compared different methods to detect direct and indirect activities that promote plant growth, with an emphasis on rhizobacteria. Moreover, we have compiled a detailed description of the methods that could be of interest for analysing bacterial isolates that exhibit potential plant growth-promoting activities.

8.1

Introduction

The human population has doubled over the past 50 years, and it is expected that by the year 2050, the population would be close to 10 billion; it has been proposed that this is the maximum capacity the Earth can hold (Wilson 2003). To further aggravate the situation, the areas of arable land are reducing drastically, i.e. soil degradation has been reported to affect 1.9 billion hectares of agricultural land worldwide, which G. Santoyo (*) · J. M. Sánchez-Yáñez Instituto de Investigaciones Químico Biológicas, Universidad Michoacana de San Nicolás de Hidalgo, Morelia, Michoacán, Mexico e-mail: [email protected] S. de los Santos-Villalobos CONACYT-Instituto Tecnológico de Sonora, Ciudad Obregón, Sonora, Mexico © Springer Nature Singapore Pte Ltd. 2019 D. Reinhardt, A. K. Sharma (eds.), Methods in Rhizosphere Biology Research, Rhizosphere Biology, https://doi.org/10.1007/978-981-13-5767-1_8

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is increasing rapidly at a rate of 5–7 million hectares each year (Mabit et al. 2014), propelling us to design sustainable new strategies to increase crop production, without harming the environment and animal health. One strategy that has been proposed by several researchers is the generation of transgenic plants that can express bacterial genes with insecticidal activity to generate fruits of larger size or those with a longer shelf life. Although, this is a promising strategy, certain sectors of the population (and some academies) have rejected its use for various reasons, many of them without scientific justification (Domingo 2016). Thus, an alternative way to improve crop yield and to control diseases caused by phytopathogens is the use of plant growth-promoting rhizobacteria (PGPR) (Kloepper et al. 1980; Santoyo et al. 2012). Therefore, the use of PGPR is a sustainable and viable strategy to improve plant production to feed the human population and has been widely accepted by various sectors of the population and regions of the world (Glick 2014). Currently, a registry list is of more than 150 microbial strains as bio-inoculants that are applied to agricultural crops; however, these strains are not enough to cover the worldwide demand (Parnell et al. 2016). Unfortunately, many PGPR with high potential are not commercialized, even in the regions from where these strains are isolated (Bashan et al. 2014). The success of a bio-inoculating product, whether for biocontrol of phytopathogens or biopromotion of growth or production of agricultural crops, begins with the isolation and search for beneficial characteristics of the potential microorganisms (Höfte and Altier 2010). Moreover, the microorganism needs to go through multiple stages from the collection of the sample from the field to the laboratory, for characterization of the mechanisms of its biocontrol and plant growth-promoting activities based on reliable methods, which sometimes require multiple tests to determine a certain activity, and from the laboratory to its application in the field with a robust statistic, allowing a good projection during its inoculation in the greenhouse or field to term it as a biocontrol agent (Parnell et al. 2016; de los Santos-Villalobos et al. 2018). This procedure will ensure that these microbial products gain the confidence of the producer for its continued commercialization and effective application over the years.

8.1.1

The Rhizosphere as a Reservoir of Plant Growth-Promoting Bacteria

The rhizosphere is the portion of the soil that is influenced by root exudates (Brink 2016). Root exudates include molecules, such as amino acids, vitamins, and sugars, which serve as nutrients for the microbial communities (Badri et al. 2009). In this ‘small’ ecosystem, bacteria are present in abundance because they utilize the nutrients from the rhizosphere and can interact and stimulate the growth of the plant by direct or indirect mechanisms. Several reviews have described these functions with detailed activities (Glick 2012; Santoyo et al. 2012; OrozcoMosqueda et al. 2018). Direct activities include functions that facilitate the availability of nutrients to the plants or that produce and modulate their hormones.

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Indirect mechanisms include plant protection against phytopathogens, e.g. by acting as biocontrol agents. Although it should be noted that certain activities can be classified under both mechanisms, e.g. production of siderophores, some volatiles, and 1-aminocyclopropane-1-carboxylate (ACC) deaminase activity (Glick 2012). Figure 8.1 shows a summary of direct and indirect mechanisms exhibited by rhizospheric bacteria to promote plant growth. Multiple studies have shown that the rhizosphere is a reservoir or source of promising plant growth-promoting bacteria (Ahemad and Kibret 2014). It has also been proposed that some types of soil may contain between 108 and 109 bacterial cells per gram of soil, including diversity of thousands of species (Schoenborn et al. 2004). Obviously, the diversity and abundance of bacterial species may vary depending on the type of soil and whether it presents any stress (e.g. nutrient and water shortages, salinity, and metal contamination) for the bacterial life. It has been suggested that abundance in stressed soils can drop to 104 cells per gram of soil (Timmusk et al. 2011). Owing to this abundance and diversity of the bacteria in the rhizosphere, it is rather appealing to analyse different rhizospheric environments worldwide, with different environmental conditions, to search for PGPR that are viable options as bio-inoculants that enhance local, regional, or global agriculture. In fact, several PGPR species have managed to pass the laboratory barrier and have been commercialized; some of the bacterial genera include Rhizobium, Streptomyces, Serratia, Pantoea, Burkholderia, Azospirillum, and Agrobacterium. However, the vast majority of the species mostly used include the genera Bacillus and Pseudomonas. This is probably due to the fact that Bacillus can sporulate, facilitating its storage, encapsulation, and subsequent application in the field (Villarreal-Delgado et al. 2018). In the case of Pseudomonas, which do not sporulate, it is possibly used owing to its abundance in ‘suppressor’ soils and because it contains an abundant resource of antimicrobial metabolites, phytohormones, and volatile compounds, which can activate a range of direct and indirect mechanisms for plant growth promotion (Santoyo et al. 2012; Hernández-León et al. 2015).

8.2

Direct Methods to Promote Plant Growth

8.2.1

Production of Phytohormones

Phytohormones are signal molecules that play important roles as regulators during the different stages of plant growth and development. Likewise, biotic and abiotic factors can be a signal for plants to regulate their internal hormonal levels for reducing the stressful effects (Verma et al. 2016). However, in addition to plants, several species of rhizobacteria also produce phytohormones and exhibit effects similar to those of phytohormones produced by plants.

8.2.1.1 Indoleacetic Acid (IAA) One of the most studied plant hormones produced by rhizobacteria is the auxin IAA. IAA participates in processes such as seed germination, controls vegetative

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Fig. 8.1 Direct and indirect mechanisms exhibited by rhizospheric bacteria to promote plant growth. (a) Plant hormones are produced by rhizobacteria, including the auxin indoleacetic acid (IAA), gibberellins (GAs), and cytokinins (CKs), which can exert different types of regulation during the growth and development of the plant. (b) Ethylene, a plant regulator of the response to various types of stress. Ethylene levels can be reduced by the action of the bacterial enzyme 1-aminocyclopropane-1-carboxylate (ACC) deaminase by hydrolysing the precursor ACC into α-ketobutyrate and ammonia. (c) Rhizobacteria can facilitate the acquisition of essential nutrients

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growth processes, and formation of lateral and adventitious roots; moreover, it can mediate processes of response to light and gravidity, affects photosynthesis, production of metabolites, and resistance to different types of stress (Fahad et al. 2015). Different genera of PGPR have been studied as producers of IAA, such as Aeromonas (Halda-Alija 2003), Azotobacter (Ahmad et al. 2008), Bacillus (Swain et al. 2007), Bradyrhizobium (Fukuhara et al. 1994), Burkholderia (Halda-Alija 2003), Pseudomonas (Hernández-León et al. 2015), and Rhizobium (Ghosh et al. 2008). IAA produced by rhizobacteria can promote or decrease plant growth, which is dependent on the levels of IAA produced and the internal IAA levels of the specific plant (Glick 2012). Owing to the aforementioned reasons, detecting IAA synthesis in rhizobacteria as a potential biopromotors of plant growth is important. One of the detection methods of this phytohormone is the Salkowski’s colorimetric technique (Gordon and Weber 1951). Currently, there are several versions of the technique. Although the method is not as accurate because it can detect other compounds (indolepyruvic acid and indoleacetamide), it has been widely used as it is very simple, fast, and cheap for analysing a high number of bacterial isolates. The Salkowski’s reagent, which contains perchloric acid (or sulphuric acid), oxidizes the indole groups present in IAA to yield pink colour of different shades. Colour changes can be detected at the wavelength of 530 nm, which is the wavelength at which highest absorbance of IAA occurs. This method always requires the implementation of a standard curve with pure IAA. Furthermore, it is required that the bacterial cultures are supplemented with tryptophan as a precursor of IAA synthesis in bacteria, which is the main route of IAA synthesis (Ghosh et al. 2008). Other more accurate methods for detecting IAA, as well as other hormones in bacteria, are high-performance liquid chromatography (HPLC) or gas chromatography-mass spectrometry (GC-MS) (Fett et al. 1987). These methods are very precise but are time-consuming when analysing few samples and require expensive equipment and materials; therefore, they cannot be used routinely for the screening of high number of rhizobacterial strains. Thus, it is recommended to use colorimetric methods (such as Salkowski’s) for primary screening, followed by HPLC or GC-MS for complete and precise IAA quantification (de los SantosVillalobos et al. 2013).

ä Fig. 8.1 (continued) in plants, such as iron (Fe), phosphorus (P), and nitrogen (N). (d) Antagonistic rhizobacteria synthesize an array of antimicrobial compounds and enzymes with lytic activity to inhibit the growth of various plant pathogens, thereby reducing disease symptoms in crops. (e) Rhizobacteria that produce siderophores (Fe-chelating compounds) can facilitate the acquisition of Fe to the plant, and at the same time, render the plant less available to potential phytopathogens. Volatile organic compounds emitted by rhizobacteria can also promote plant growth and exhibit inhibitory action against fungal pathogens. For example, dual activities have been reported for dimethylhexadecylamine (DMHDA)

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8.2.1.2 Gibberellins (GAs) and Cytokinins (CKs) GAs are plant hormones that regulate a wide variety of developmental phenomena in plants, including cell elongation, primarily the stem, and germination of seeds. However, it was not first discovered in plants but in the fungus Gibberella, and, hence, the name was given as gibberellin (Phinney 1983). In the case of CKs, their role has been documented in multiple aspects of plant growth and development, such as cell division, shoot initiation and growth, leaf senescence, apical dominance, nutrient uptake, and root nodulation (Sasaki et al. 2014). GAs and CKs have also been studied in PGPR that produce GAs and/or CKs. According to Glick (2012), our knowledge on the role of GAs and CKs produced by bacteria is based on physiological studies in growing plants supplemented with exogenous addition of purified hormones for growth. One of the most effective techniques to detect the production of GAs and CKs, among other phytohormones, is HPLC. Other techniques such as Sephadex G-25 and Dowex-50 column chromatography and thin-layer chromatography (TLC) have also been used to detect, isolate, and characterize GAs and CKs (Karadeniz et al. 2006). A new version of TLC is high-performance thin-layer chromatography (HPTLC), which allows automaton of different steps of the conventional TLC technique. Moreover, HPTLC provides an increased resolution that allows more precise quantitative measurements (Reich and Schibli 2006).

8.2.2

ACC Deaminase Activity

Ethylene is one of the most studied plant hormones because it can regulate various aspects of plant development, including induction of germination. In addition, ethylene participates as a regulator of the response to various types of stress, including stress due to abiotic type factors such as floods, drought, and temperature. Furthermore, it mediates the responses to stress due to phytopathogen attack, mechanical damage and injuries, and growth in soils contaminated with heavy metals (Fujita et al. 2006). All these stresses can induce an increase in ethylene production by plants, leading to their senescence, chlorosis, and abscission. Therefore, beneficial bacteria that interact with plants can alleviate different types of stress because they contain the enzyme ACC deaminase that hydrolyses ACC (the immediate precursor of ethylene in all higher plants) to α-ketobutyrate and ammonia. These compounds are easy to assimilate, and, therefore, an increase in ethylene levels in the plant is prevented. ACC deaminase was first described in soil bacteria by Honma and Shimomura (1978), and since then, many studies have reported that the application of PGPR with ACC deaminase activity to seeds and plant roots promote the plant’s growth as well as crop production, even under adverse conditions (Glick 2012). A method for evaluating the bacterial activity of ACC deaminase, which has been widely explored in numerous studies, is described in detail in the article by Penrose and Glick (2003). This method involves spectrophotometry routinely used in

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laboratories and is based on measuring the activity of ACC deaminase as a modification of the method of Honma and Shimomura (1978) by measuring the amount of α-ketobutyrate produced when ACC deaminase hydrolyses ACC. The number of μmol of α-ketobutyrate produced by this reaction is determined by comparing the absorbance at 540 nm of a sample with a standard curve of α-ketobutyrate. Furthermore, Penrose and Glick (2003) also proposed a gnotobiotic root elongation assay in canola (Brassica campestris) plants to evaluate growth promotion by bacteria exhibiting ACC deaminase activity.

8.2.3

Acquisition of Nutrients

A vast majority of the world’s arable land is not suitable or does not contain the nutrients necessary for good agricultural development and production. Therefore, chemical fertilizers are used excessively. In multiple reports, it has been documented that plants uptake only a low percentage (20–50%) of the fertilizer applied and the rest is volatized, which produces greenhouse gases, and leached, resulting the contamination of the aquatic mantles. In addition, the excessive use of fertilizers decreases the soil quality (Tilman et al. 2002). Therefore, using rhizobacteria that facilitate the acquisition of nutrients in plants, such as iron (Fe), phosphorus (P), and nitrogen (N), is one of the most studied direct mechanisms of plant growth promotion, and its application to crops is now essential.

8.2.3.1 Biological Nitrogen Fixation N is important for living organisms, including those that play a key role in the nitrogen cycle, because it is a part of various biological molecules (e.g. proteins and nucleic acids). Majority of the N enters the ecosystems, including the agricultural ecosystem, by biological fixation of atmospheric N2 (Levy-Booth et al. 2014). Atmospheric N2 is relatively inert, instead of ‘fixed’ forms such as ammonia (NH3) or nitrate (NO3 ) (Hoffman et al. 2014). It has been recognized for decades that nitrogen fixation (conversion of N2 to NH3) is important and often a limiting factor in agricultural production. To counteract this limitation, nitrogen fertilizers are usually applied to poor soils, although their negative consequences to the environment are known. The alternative to this problem is using PGPR with the ability to fix atmospheric nitrogen. PGPR of the genera Bradyrhizobium, Mesorhizobium, Sinorhizobium, and Rhizobium are part of the rhizobial order, known as rhizobia. Rhizobia are diazotrophic (freeliving) microorganisms present in the root nodules of leguminous plants, and their distinguishing characteristic is the ability to fix atmospheric nitrogen in symbiosis with leguminous plants (Mus et al. 2016). Thus, most studies on nitrogen fixers have been performed in symbiotic legume-rhizobia. Detecting nitrogen-fixing bacteria in the rhizosphere is not an easy process. Usually, there is no, as far as we know, selective medium for nitrogen-fixing bacteria. Rhizobia proliferate in low salt media (NaCl2); however, highly diverse non-nitrogen-fixing bacteria also proliferate in the media. One of the previous

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methods to determine if the bacterial strain can fix nitrogen is the detection of the nifH gene (encoding one component of the nitrogenase enzyme, which is the only family of enzymes responsible for the reduction of atmospheric nitrogen to ammonia) by PCR (Widmer et al. 1999; Zehr et al. 1998). Once nifH has been detected, these isolates could be termed as potential nitrogen-fixing bacteria. For decades, this method has been the most used strategy for identifying nitrogen fixers, although other techniques have been proposed (Burris and Wilson 1957; Lopez-Lozano et al. 2016). The most commonly and widely used method to assess N2 fixation (an indirect method) is the acetylene reduction assay as it measures ethylene formation from acetylene by nitrogenase. More recently, fragments of nifH have been detected in tissue mRNA samples of sugarcane and potato plants (Terakado-Tonooka et al. 2008; Thaweenut et al. 2011), confirming the functionality of nifH in endophytic bacteria, such as Azospirillum, Herbaspirillum, Azoarcus, Acetobacter, and Burkholderia, and not only in the root nodules of leguminous plants. This could lead to the potential isolation of these bacteria and their use as inoculants in non-legume crops. Undoubtedly, a more efficient method for screening nitrogen fixers is necessary that would benefit the scientific community and would promote studying nitrogen-fixing microorganisms in various environments such as soil, lakes, and forests.

8.2.3.2 Solubilization of Phosphates P is an essential macronutrient for plant growth as it is involved in photosynthesis, biological oxidation, nutrient uptake, and cell division (Meena et al. 2016). However, despite its abundance in soils, P is insoluble and not available to plants because of the soil pH. Therefore, like other nutrients, such as N, agricultural soils worldwide are supplemented with inorganic P in the form of chemical fertilizers to improve crop production. The use of phosphorus-solubilizing bacteria as inoculants increases P uptake by plants. Strains of the genera Pseudomonas, Bacillus, and Rhizobium are among the most efficient phosphate solubilizers. The main mechanism for the solubilization of mineral phosphate is the production of organic acids and acid phosphatases, which participate in the mineralization of organic phosphorus in the soil (Rodríguez y Fraga 1999). Isolating phosphate-solubilizing rhizobacteria is relatively easy, fast, and cheap by using Petri dishes containing Pikovskaya (PVK) agar (Pikovskaya 1948). The medium (PVK) contains tricalcium phosphate (TCP) as a source of phosphates (Nautiyal 1999). The inoculated Petri dishes are incubated for a few days (approximately 7 days) with potential P-solubilizing bacteria. Post incubation, colonies with a clear halo are marked as positive for phosphate solubilization. Phosphatesolubilizing bacteria can also be analysed in liquid medium, and the content of soluble-P is estimated colorimetrically (Jackson 2005). Moreover, other methods and protocols with some variants have been proposed for qualitative screening of phosphate-solubilizing bacteria based on visual observation, with similar efficiencies (Nautiyal 1999; Mehta and Nautiyal 2001).

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Indirect Methods to Promote Plant Growth

8.3.1

In Vitro Antagonism

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The antagonistic action of rhizobacteria and biological control of diseases caused by phytopathogens is recognized as an indirect mechanism to promote and improve the growth and health of plants. Therefore, detecting these activities in rhizobacteria is a desirable characteristic in bio-inoculants (Santoyo et al. 2012, 2016). Detecting rhizobacteria with antagonistic activity (antagonists) can be performed through confrontation tests in Petri dishes, either using other bacteria or phytopathogenic fungi. Trials may include the confrontation of one antagonist, one pathogen; two antagonists, one pathogen; three antagonist, one pathogen; and four or six antagonists, one pathogen in the inocula. Inoculation of the bacterial antagonist may also vary by inoculating either a drop containing a certain amount of CFU/mL or inoculation using a microbiological loop. To detect the action of diffusible compounds in the medium, culture supernatants are usually purified in liquid media (after purification of the bacterial cells). Likewise, to evaluate the production of organic volatiles, divided Petri dishes are used where there is no physical contact between the organisms (Hernández-León et al. 2015). These trials are highly valuable and have been continually described in recent publications worldwide to search for new antagonists. Of course, the use of controls without inoculation is essential to obtain reliable results. Recently, Hernández-León et al. (2015) screened for bacterial antagonists isolated from the rhizosphere of plants of Medicago truncatula, where they inoculated up to 100 different bacterial isolates around an agar plug containing a phytopathogenic filament fungus in a single Petri dish. However, special care must be taken for fast-growing isolates, such as some strains of Bacillus licheniformis, which can cover the entire Petri dish within a few hours. The result was isolation and characterization of four PGPR strains with direct and indirect activities that promote plant growth.

8.3.2

Synthesis of Antibiotics and Lytic Enzymes

The use of antagonistic rhizobacteria affords the ability to synthesize an array of antimicrobial compounds and enzymes that exhibit lytic activity to inhibit, restrict, or eliminate the growth of various phytopathogens (Liu et al. 2017). Bacteria of the genera Pseudomonas and Bacillus are leading in such studies, and antibiotics, such as 2,4-diacetylphloroglucinol, phenazine-1-carboxyclic acid, phenazine-1carboxamide, pyoluteorin,pyrrolnitrin, oomycinA, viscosinamide, butyrolactones, kanosamine, zwittermicin-A, aerugine, rhamnolipids, cepaciamide A, ecomycins, pseudomonic acid, azomycin, and cepafungins, produced by these PGPR have been reported. In case of lytic enzymes, chitinases, lipopeptides, cellulases, and glucanases with antagonist activity have been reported (Villarreal-Delgado et al. 2018). These antibiotics and lytic enzymes possess antagonistic activity against specific phytopathogens, viruses, insects, and helminths and exhibit phytotoxic,

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antioxidant, cytotoxic, and antitumour action (Fernando et al. 2005; MartínezAbsalón, et al. 2014). To detect the synthesis of antibiotics in rhizobacteria, techniques such as HPLC are usually used. However, one can prescreen for genes that code for various antibiotics using primer combinations for PCR. Zhang et al. (2006) performed this technique by using 30 primers to amplify biosynthetic genes, encoding phenazine-1-carboxylic acid, 2,4-diacetylphloroglucinol, pyoluteorin, pyrrolnitrin, and zwittermicin A, in strains of Pseudomonas and Bacillus. Further evaluation of the strains revealed the production of these compounds and was confirmed by HPLC. To detect lytic activities, Petri dishes with specific media can be used. For example, trypticase soy agar (TSA) medium is used for detecting proteases; the formation of a transparent halo around the colony is indicative of proteolytic activity (Cattelan et al. 1999). Cellulase, chitinase, and glucanase activities can be detected by measuring the degradation of specific substrates, such as chitin and cellulose, and are analysed by spectrophotometric methods (de los SantosVillalobos et al. 2012). The released by-products can also be compared to control standards. The use of specific inhibitors, such as allosamidin, a chitinase-specific inhibitor, can yield more specific results of the action of such specific lytic enzymes (Martínez-Absalón et al. 2014). Some other compounds such as lipopeptides of the iturin family exhibit antagonist activity against diverse phytopathogenic fungi. Some members of the iturin family, such as mycosubtilin, also exhibit haemolytic activity (Maget-Dana and Peypoux 1994; Leclère et al. 2005). Other lipopeptides, such as surfactin, produced by Bacillus strains also show haemolytic activity (Hsieh et al. 2004). Haemolytic activities can be easily detected by inoculating the rhizospheric bacteria on Petri dishes containing blood agar, and haemolytic halos (β-haemolysis) are observed around the colonies after few hours of incubation (Hernández-Salmerón et al. 2014). Thus, formation of haemolytic halos can be a pre-indication of lipopeptide production by potential antifungal rhizospheric strains, but further analysis are required to confirm the presence or type of lipopeptide and its potential risks to human health.

8.3.3

Competition in the Rhizosphere

The rhizosphere is a highly competitive microenvironment as the microorganisms that inhabit it compete to obtain the nutrients that exude the roots of the plants and occupy the best spaces and niches. Usually, the success of bacteria inoculated in the rhizosphere to improve plant health through mechanisms of biocontrol and promotion of plant growth is, in large part, attributed to being efficient colonizers of the rhizosphere (Bloemberg and Lugtenberg 2001). Various methods, such as promoter-trapping technology, in vivo expression technology (IVET), are important to identify genes whose expression is high in the rhizosphere (Rainey 1999). Other methods proposed by Scher et al. (1984) have been relevant to evaluate the ability of rhizosphere and rhizoplane colonization by plant growth-promoting bacteria in non-sterile soils. Such methods include

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inoculation of a certain amount of CFU/mL of the bacteria in corn seeds (an ideal plant for this purpose), which germinates and roots grow through the container. Colonization of the inoculated bacteria in the different zones of the roots, measured as CFU, can enable detection of competitive bacteria in a non-sterile environment (rhizoplane and rhizosphere). However, further analyses using random amplified polymorphic DNA (RAPD) and fluorescent markers and determining resistance to antibiotics are required (Dekkers et al. 2000). Furthermore, this method has enable the evaluation of the best mixtures of bacterial inoculants that are highly competitive and do not exhibit antagonism between different species (Rojas-Solís et al. 2016).

8.4

Compounds with Dual (Direct and Indirect) Activity

8.4.1

Siderophores

Fe is one of the most abundant elements on earth; however, it is not bioavailable in all forms. For example, ferric ion (Fe+3) is the most abundant but also the least assimilable form by plants (Taylor and Konhauser, 2011). Therefore, Fe deficiency in agricultural crops is a global problem. To compensate for this deficiency, various organisms, including beneficial bacteria and plants, produce siderophores that allow them to acquire Fe in different organic and inorganic forms present in the soil (Saha et al. 2016). Other important role of siderophores produced by rhizobacteria, besides increasing plant growth, is biocontrol. Klopper et al. (1980) proposed that the synthesis of siderophores and Fe chelation in the rhizosphere by bacteria of the genus Pseudomonas is a mechanism that restricts the availability of this nutrient to potential pathogens, thereby inhibiting their growth. Therefore, detecting the synthesis of siderophores in rhizobacteria is a desirable characteristic in PGPR. The production of siderophores can be easily detected through the growth of bacterial colonies in plates with CAS medium (chrome azurol sulphonate) (Schwyn and Neilands 1987). If siderophores are secreted, they will chelate Fe in the media and form a Fe-siderophore complex, thereby changing the medium colour from blue to yellow-orange tones. This colour change is because siderophores exhibit a greater affinity for Fe than that for chromogen. The synthesis and production of siderophores for purification purposes in liquid media are also feasible and very useful to determine its specific role and to differentiate its effect from other metabolites (Elad and Baker 1985). The presence of siderophores excreted in the medium and subsequent purification can be determined using a spectrophotometer by measuring the absorption peak at 408 nm, and the concentration of the compounds can be determined using the maximum absorption value and molar absorption coefficient according to the method of Meyer and Abdallah (1978). Other recent methods with some variations useful for screening siderophore-producing strains have been proposed by using 96-well microplates (Arora and Verma 2017).

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Volatile Organic Compounds (VOCs)

Similar to VOCs in organisms of the three different domains of life, the volatiles produced by rhizobacteria also play different and important roles in communication (Bitas et al. 2013; Gershenzon and Dudareva 2007; Stotzky and Schenk 1976). Ryu et al. (2003) demonstrated the role of volatile compounds, primarily 2,3-butanediol and acetoin, produced by beneficial bacteria in promoting the growth of Arabidopsis thaliana. One of the characteristics of their study was the use of divided Petri dishes that allowed differentiating the effects of compounds that are diffusible in the medium versus volatile compounds produced by the bacteria. Following this study, multiple studies demonstrated the role through which VOCs produced by rhizobacteria promote the growth and health of the plants. VOCs have been assigned different functions, including direct and indirect mechanisms for plant growth promotion. For example, the compounds hydrocyanic acid (HCN), dimethyl disulphide (DMSD), and N,N-dimethyl hexadecylamine (DMHDA) are potent inhibitors of the growth of phytopathogenic mycelium and simultaneously present activities to promote plant growth (Orozco-Mosqueda et al. 2013; Rojas-Solís et al. 2018). Likewise, the roles of rhizobacterial volatiles as initiators of defence responses in plants have been reported (Ryu et al. 2004). Most emitted compounds are species-specific, but some volatile blends overlap in rhizobacteria like Pseudomonas and Stenotrophomonas (Rojas-Solís et al. 2018; Kai et al. 2009). Analysing the antagonistic activity or promoting the growth of plants is relatively easy through the system of divided Petri dishes; however, for determining and differentiating the profile of volatile bacterial compounds, GC-MS is primarily performed. Although GC-MS is a powerful method to quantify and identify volatiles, there are also some limitations, as suggested by Kai et al. (2009). For example, the compounds can only be considered identified if they exhibit a mass spectrum that coincides with the library-employed identical Kovats indices on two columns of different polarity. Likewise, the identification of new compounds has to be elucidated by other methods, such as nuclear magnetic resonance analysis (Kai et al. 2009).

8.5

Conclusions and Outlook

In this book chapter, we have discussed some mechanisms that promote plant growth directly and indirectly and some screening methods most commonly reported in literature to detect potential PGPR. Some PGPR can be cultured relatively easily and economically, with screenings on Petri dishes using a specific media to detect such activities (i.e. siderophores, proteases, or solubilization of phosphates), for rapid detection of the plant beneficial activities exhibited by potential rhizospheric bacteria. Other methods are more elaborate or require expensive equipment; therefore, cheaper, faster, and more effective options are proposed as prescreening methods. It is also clear that other techniques or molecular tools were outside the scope of this review, such as metagenomics, transcriptomics, or metabolomics. These tools

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are currently very relevant to detect genes related to plant growth promoter mechanisms and for massive sequencing analysis of ribosomal genes (16S/18S) of a hidden diversity that cannot be discovered by microbial culture methods (Nesme et al. 2016). There are also other mechanisms that were not detailed or were mentioned superficially in this work (owing to word count limit), such as systemic resistance induced by rhizobacteria. However, multiple studies can be referred to for further information. Finally, this chapter intends to inform students and young researchers who are interested in embarking on this fascinating area of research on beneficial bacteria, their mechanisms of growth stimulation in plants, and biocontrol of potential phytopathogens. Although this area is fairly justified by its agronomic and, therefore, economic importance, it is still relevant in the basic aspect of discovering basic mechanisms of plant-bacteria-pathogen communication and interaction. Without a doubt, both of these aspects of basic and applied research are unexplored oceans waiting for new sailors. Acknowledgements G.S. thanks the Coordinación de la InvestigaciónCientífica of the Universidad Michoacana de San Nicolás de Hidalgo for the financial support to research projects.

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A Split-Root Method to Study Systemic and Heritable Traits Induced by Trichoderma in Tomato Plants M. B. Rubio, H. A. de Medeiros, M. E. Morán-Diez, P. Castillo, R. Hermosa, and E. Monte

Abstract

The split-root methodology constitutes an excellent tool to study local versus systemic plant-induced responses. In the most common approach, two different organisms coinfect the two separated root halves of a same plant. Split-root plants have been used to study the biocontrol potential of fungi and bacteria by the induction of systemic defenses in the plant against bacterial, fungal, oomycete, and nematode diseases and insect pests. In our particular case study, we applied this methodology to demonstrate the systemic and heritable effects induced by the biocontrol strain Trichoderma atroviride T11 in tomato plants which were tested against the root-knot nematode (RKN) Meloidogyne javanica (Mj), a major tomato pathogen worldwide. This approach allows Trichoderma and the root pathogen to be kept separate for the analysis of the T11 effects on the penetration, development, and reproduction of Mj in tomato roots upon activating plant systemic responses. The method also enables the plant green mass and nematode infection parameters to be determined and the gene expression analysis related to systemic responses and heritable traits, in terms of defense and growth, induced by T11 when plants are infected with Mj in the progeny of split-root plants.

M. B. Rubio · H. A. de Medeiros · R. Hermosa · E. Monte (*) Department of Microbiology and Genetics, Spanish-Portuguese Institute for Agricultural Research (CIALE), University of Salamanca, Salamanca, Spain e-mail: [email protected] M. E. Morán-Diez Department of Microbiology and Genetics, Spanish-Portuguese Institute for Agricultural Research (CIALE), University of Salamanca, Salamanca, Spain Phytopathology Department, University of Viçosa, Viçosa-MG, Brazil P. Castillo Institute for Sustainable Agriculture, Spanish National Research Council (CSIC), Córdoba, Spain © Springer Nature Singapore Pte Ltd. 2019 D. Reinhardt, A. K. Sharma (eds.), Methods in Rhizosphere Biology Research, Rhizosphere Biology, https://doi.org/10.1007/978-981-13-5767-1_9

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Introduction

Trichoderma is a genus of filamentous fungi that includes species used as biological control agents against a wide range of plant pathogens under different agriculture models (Harman et al. 2010). The most common mechanisms effectuating the biocontrol behavior of Trichoderma include mycoparasitism, competition for nutrients, and/or antibiosis (Lorito et al. 2010). In addition, some Trichoderma strains exert beneficial effects on plants in terms of increased percentages and rates of seed germination, nutrient uptake, growth promotion, and defense stimulation against biotic and abiotic stresses (Shoresh et al. 2010; Hermosa et al. 2012; Rubio et al. 2014, 2017). Trichoderma biocontrol activity against the root-knot nematode (RKN) Meloidogyne javanica (Mj) has been observed in greenhouse experiments (Rao et al. 1998; Sharon et al. 2001; Goswami et al. 2006; Pedroche et al. 2009; Sokhandani et al. 2016). This RKN is a highly adapted obligate biotrophic species that infects numerous plant species from many different families worldwide (Abad et al. 2009). Infection by Mj begins with penetration of the root tissues by a secondstage juvenile (J2) at the root elongation zone. The nematode then migrates in the apoplast, ultimately reaching the vascular cylinder where the formation of giant cells (GC) is induced. Thereafter, the J2 undergoes three molts, developing into an adult. Female adults remain sedentary, producing large egg masses and root galls, while adult males migrate to the soil. Secretions from RKN contain effectors that act as primary signaling molecules at the plant-nematode interface. The oxidative response of the host to nematode infection is a key component of the plant’s defense strategy, although antioxidant enzymes secreted by RKN allow the nematode to survive (Abad et al. 2003; Gheysen and Mitchum 2011). Split-root plants have been widely used for a variety of experimental purposes. This system displays several features that make it an excellent tool to study local versus systemic plant-induced responses. Table 9.1 lists research studies that describe the use of this system to explore systemic mechanisms triggered by biological control organisms. This kind of methodology can be applied under a variety of experimental conditions: (i) to evaluate the systemic effects of two populations of pathogenic species (Harris and Ferris 1991; Umesh et al. 1994; LaMondia 2003; Aryal et al. 2011); (ii) to study the regulation of mycorrhization and nodulation (Sargent et al. 1987; George et al. 1992; Vierheilig et al. 2000; Catford et al. 2003); (iii) to determine systemic effects of applying plant elicitors such as lipopolysaccharides, root exudates, or phytohormones on pathogens (Reitz et al. 2000; Vierheilig et al. 2003; He and Wolyn 2005; Soler et al. 2013); (iv) to analyze as to how soil amendments affect microbe communities or plant resistance (Lievens et al. 2001; Cohen et al. 2005; Yogev et al. 2010; Eizenberg et al. 2017); and (v) to study the effect of abiotic stresses (water irrigation, salinity, mineral uptake dynamics, fertilization, etc.) on plant fitness (Li et al. 2016; Lima et al. 2016; Redwan et al. 2017; Xin et al. 2017). However, split-root approaches have their limitations since data replicates are often scarce due to the own peculiarity of this approach when it is applied under in vivo conditions. In addition to being

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Table 9.1 List of research studies where authors describe the use of a split-root methodology to test the plant-induced responses triggered by an inducer organism against a responder-pathogenic organism (including in both cases species of different taxonomic groups: fungi, bacteria, nematodes, or oomycetes) Plant system Arabidopsis Asparagus Banana

Barley

Carnation Chickpea Cotton Cucumber

Eggplant

Grapevine

Grass (Ammophila arenaria) Medicago Olive Pigeon pea

Tested organisms (inducer/responder) Fusarium oxysporum1*/Meloidogyne incognita3 F. oxysporum*/F. oxysporum f. sp. asparagi1 Rhizophagus irregularis (Glomus intraradices)1 (AMF)/Radopholus similis3 and Pratylenchus coffeae3 F. oxysporum* and Fusarium diversisporum1*/ R. similis R. irregularis, Funneliformis mosseae (Glomus mosseae)1, and Gigaspora rosea1 (AMFs)/ Gaeumannomyces graminis var. tritici1 Pseudomonas fluorescens2/Fusarium graminearum1 P. fluorescens/Pythium ultimum4 F. mosseae/G. graminis var. tritici Pochonia chlamydosporia1, Pochonia rubescens1, and Lecanicillium lecanii1/G. graminis var. tritici F. oxysporum*/F. oxysporum f. sp. dianthi F. oxysporum*/F. oxysporum f. sp. ciceri Bacillus vallismortis2 and Glomus versiforme1 (AMF)/Verticillium dahliae1 Pseudomonas corrugata1 and Pseudomonas aureofaciens2 (PGPRs)/Pythium aphanidermatum4 Trichoderma hamatum1/Phytophthora capsici4 Pseudomonas putida2 and Serratia marcescens2 (PGPRs)/Fusarium oxysporum f. sp. cucumerinum Bacillus amyloliquefaciens2 (PGPR)/Fusarium oxysporum f. sp. cucumerinum Paenibacillus alvei2/V. dahliae F. oxysporum*/V. dahliae Arthrobacter spp.2 and Blastobotrys (Arxula) spp.1/ V. dahliae1 R. irregularis/Xiphinema index3 M. incognita*/Meloidogyne arenaria3 Glomus spp.1 and Scutellospora castanea1 (AMF)/ Pratylenchus penetrans3 Catenaria spp.1/Tylenchorhynchus ventralis3 F. mosseae1/Aphanomyces euteiches4 P. fluorescens/V. dahliae Bacillus cereus2 and P. aeruginosa/Fusarium udum1

Literature Martinuz et al. (2015) He et al. (2002) Elsen et al. (2008)

Vu et al. (2006) Castellanos-Morales et al. (2011, 2012) Henkes et al. (2011) Jousset et al. (2011) Khaosaad et al. (2007) Monfort et al. (2005) Postma and Luttikholt (1996) Kaur and Singh (2007) Zhang et al. (2012) Chen et al. (2000) Khan et al. (2004) Liu et al. (1995) Liu et al. (2017) Antonopoulos et al. (2008) Pantelides et al. (2009) Papasotiriou et al. (2013) Hao et al. (2012) McKenry and Anwar (2007) De la Peña et al. (2006) Piskiewicz et al. (2009) Zhang and Franken (2014) Cabanas et al. (2017) Dutta et al. (2008) (continued)

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Table 9.1 (continued) Plant system Potato Rice

Soybean Tea Tomato

Tested organisms (inducer/responder) Bacillus sphaericus2 and Agrobacterium radiobacter2/Globodera pallida3 Paenibacillus polymyxa2 (PGPR)/F. oxysporum f. sp. niveum Serendipita (Piriformospora) indica/Lissorhoptrus oryzophilus5 Sinorhizobium fredii2/Heterodera glycines3 P. aeruginosa/Fomes lamoensis1 and Ustulina zonata1 Bacillus subtilis2/M. incognita Bacillus fortis2 and B. subtilis/F. oxysporum f. sp. lycopersici M. incognita*/M. incognita P. fluorescens and P. putida/F. oxysporum f. sp. lycopersici F. mosseae/Phytophthora parasitica4 F. oxysporum*/M. incognita Trichoderma atroviride1/Meloidogyne javanica3 P. polymyxa/M. incognita Trichoderma spp.1/M. incognita

Watermelon

F. oxysporum* and Rhizobium etli2/M. incognita Trichoderma koningiopsis1/F. oxysporum f. sp. radicis-lycopersici F. mosseae and R. irregularis/P. parasitica Trichoderma harzianum1/M. javanica B. subtilis, Bacillus atrophaeus2, and Burkholderia cepacia2 (PGPR)/F. oxysporum f. sp. lycopersici P. aeruginosa, P. fluorescens/Rhizoctonia solani1, and M. javanica F. mosseae/M. incognita and P. penetrans G. versiforme/Ralstonia solanacearum1 Fusarium spp.*/F. oxysporum f. sp. niveum

Wheat

Sterile red fungus1 (SRF)/G. graminis var. tritici

Literature Hasky-Guenther et al. (1998) Ling et al. (2011) Cosme et al. (2016) Tian et al. (2014) Mishra et al. (2014) Adam et al. (2014) Akram et al. (2013) Anwar and McKenry (2008) Boukerma et al. (2017) Cordier et al. (1998) Dababat and Sikora (2007) Medeiros et al. (2017) Khan et al. (2012) Martinez-Medina et al. (2017) Martinuz et al. (2012) Moreno et al. (2009) Pozo et al. (2002) Selim et al. (2014) Shanmugam and Kanoujia (2011) Siddiqui and Shaukat (2002a, b, 2004) Vos et al. (2012) Zhu and Yao (2004) Larkin et al. (1996); Larkin and Fravel (1999) Aberra et al. (1998)

Notes: AMF, arbuscular mycorrhizal fungus; PGPR, plant growth-promoting rhizobacteria; * nonpathogenic isolate; 1, fungal species; 2, bacterial species; 3, nematode species; 4, oomycete species; 5, insect species

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cumbersome, split root is difficult to implement, and many plant species with soft tissues are not easy to handle (Kassaw and Frugoli 2012). Recent studies using split roots have served to demonstrate that Trichoderma induces systemic resistance toward Mj in tomato, without the need for the organisms to be in direct contact, and that plants adapt their salicylic acid (SA)- and jasmonic acid (JA)-regulated Trichoderma-dependent protection according to the stage of RKN infection (Martínez-Medina et al. 2017; Medeiros et al. 2017). Moreover, the split-root system has demonstrated its usefulness in heredity studies since the progeny of Trichoderma-primed split-root tomato plants displayed increased size and inherited resistance to RKN without costs to fitness, in which the production of auxin-induced reactive oxygen species is involved (Medeiros et al. 2017). In this chapter, we expand the methods described in this work that allow Trichoderma and a target root pathogen to be used separately for the analysis of the effects of a biocontrol agent on the penetration and reproduction of a RKN nematode in tomato roots upon activating both plant systemic and heritable responses.

9.2

Approaches, Techniques, and Results

9.2.1

Experimental Design

The overall experimental design is illustrated in Fig. 9.1 in which the major steps of this study using a split-root system are highlighted. Figure 9.1 shows Trichoderma and/or Meloidogyne inoculations, seed collection to procure a first generation (F1) of plants, the sampling time points for carrying out the phenotypic analysis, the determination of the nematode infection parameters in tomato and its progeny, and analysis of defense marker genes in the offspring.

9.2.2

Tomato Split-Root System and Inoculations with Ta and Mj

Tomato seeds (Solanum lycopersicum “Marmande”) were surface-sterilized and rinsed thoroughly with sterile distilled water, sown in 3 L pots containing commercial organic substrate Tref (Jiffy, Castillo Arnedo, Navarrete, Spain) previously autoclaved during 2 successive days. The pots were maintained in a greenhouse and watered as needed. After 5 weeks, each tomato plant was transferred into two adjacent pots containing an autoclaved mixture of Tref and vermiculite (3:1, v/v) with half of the entire root system planted into each pot. Trichoderma atroviride IMI 352941 (International Mycological Institute, CABI Bioscience, Egham, UK), named as T11 or Ta, and the RKN Meloidogyne javanica (Mj) isolated from infected tomato roots in Córdoba (Spain) were the fungus and the nematode used in the plant root inoculations, respectively. T11 was grown on potato dextrose agar (PDA) medium, and conidia of 7-day-old PDA plates were harvested and then adjusted to 1x107 conidia mL
1 with sterile distilled water. For Mj inoculum preparation, nematode eggs were filtered through a 5-μm pore-size

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F0 w

w

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Plants from the five treatments were grown in the greenhouse until fruit production

w

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w Ta+Mj

15 dai with Mj (300 units of J2)

30 dai with Mj (500 units of J2)

Stain of roots with fuchsin acid and determination of: -NTNR -Number of nematodes in J2, J3, J4 and adult stages

Determination of: -NG -NEM -NEM/NG ratio

F0

Seeds

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After 14 days Determination of green mass parameters

After 15 days Gene expression analysis by qPCR

After 21 days Substrate infection with 1,000 Mj eggs 30 dai with Mj

Determination of: -NG, NEM and NEM/NG -Green mass parameters

Fig. 9.1 Experimental design of a split-root system with adult tomato plants, the fungus Trichoderma atroviride T11 and the nematode Meloidogyne javanica

membrane, exposed to 0.01% (w/v) mercury (I) chloride (Hg2Cl2) for 10 min, followed by a 0.7% streptomycin solution, and rinsed with sterile distilled water. Eggs were then collected from the membrane and placed in sieves with a 25-μm pore

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size and soaked in 0.01 M 2-(N-morpholino)ethanesulfonic acid (MES) buffer under aseptic dark conditions for 3 days; J2 s were collected after hatching. One week after transplanting the tomato plants into the split-root system, one half of the roots of each tomato plant was inoculated with 1 mL of the T11 conidial suspension (1  107 spores/mL) forming the following treatment groups: w/Ta and Ta/Mj and 1 mL of the T11 conidial suspension (1  107 spores/mL) plus 500 J2 s, w/Ta + Mj treatment group. Five days later, Mj infection was performed with 500 or 300 J2 s in treatments w/Mj and Ta/Mj. In total, the split-root experimental design consisted of five treatments: (i) the two root systems treated with sterile water (w/w) as the control condition; (ii) one half of the root system inoculated with sterile water and the other half with Ta (w/Ta); (iii) one half of the root system treated with sterile water and the other half with Mj (w/Mj); (iv) one half of the root system inoculated with Ta and the other half inoculated with Mj (Ta/Mj); and (v) one half of the root system inoculated with Ta plus Mj (w/Ta + Mj).

9.2.3

Greenhouse Experiments and Sample Collection

Greenhouse assays were carried out with the split-root experimental design as described above. Three root samples from three different replicate plants (F0) of w/Mj, Ta/Mj, and w/Ta + Mj treatments were collected at 30 days after inoculation (dai) with 500 J2 s Mj to estimate the number of galls (NG) and the number of egg masses (NEM). The average values of NG and NEM corresponded to six plants from two independent experiments. Three plants (F0) of every treatment indicated above (w/w, w/Ta, w/Mj, Ta/Mj, and w/Ta + Mj) were maintained in the greenhouse until they bore fruits, and two fruits per plant were subsequently collected. Then, a seed pool from 12 fruits was made for each of the five treatments from two independent experiments. Seeds were washed, disinfected, and sown into 500 mL pots containing autoclaved Tref substrate. Three seeds were sown per pot (14 pots in total), comprising a total of 42 F1 plants per treatment. At 14 days, five F1 plants (coming from different pots) per treatment were used to calculate the green mass index. Then, at 15 days, another ten F1 plants (again coming from different pots) per treatment were taken and pooled. The expression of defense and phytohormone cross talk marker genes was analyzed by qPCR. At 21 days, the Tref substrate of five pots containing one remaining plant each was infected with 1000 Mj eggs. At 51 days (30 dai with Mj), the five F1 plants per treatment were used to determine the NG, the NEM, and the NEM/NG ratio and to calculate the green mass geometric parameters. The average values of NG and NEM of the F1 plants corresponded to ten plants coming from two independent experiments. An additional greenhouse assay was carried out in duplicate with the split-root experimental design described above using 300 specimens of J2 in the Mj inoculations. Root samples from F0 plants of the treatments w/Mj, Ta/Mj, and w/Ta + Mj were used to determine the number of total nematodes inside the roots

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(NTNR) and the developmental stages of Mj at 15 dai. Data were recorded as average values of eight root samples from two independent experiments.

9.2.4

Nematode Measurement Assay

Root samples were washed, dissected into 1- to 2-cm segments, and placed in 50 mL of tap water. Then root tissue pieces were disinfected with 1.5% sodium hypochlorite and rinsed with tap water. The material was then drained and transferred to a beaker containing 30 mL of water to which 1 mL of stain (3.5 g acid fuchsin, 250 mL acetic acid, and 750 mL distilled water) had been added. This solution was then heated to boiling for approximately 30 sec. After cooling to room temperature and rinsing with running tap water, the root material was placed in 20–30 mL of glycerine acidified with a few drops of 5 N HC1, heated to boiling, and cooled. The root segments were then pressed between microscope glass slides for observation, and the NG and NEM were counted (Bybd et al. 1983; Silva et al. 2009). These data were used to calculate the NEM/NG ratio. Moreover, root tissues from F0 plants were observed microscopically to determine the NTNR and the developmental stage of the nematodes (Abad et al. 2009). The developmental stages identified were vermiform J2, swollen J2, third-stage juveniles (J3), fourth-stage juveniles (J4), young females without eggs, females with egg masses, and mature males. J3 and J4 were inactive, lacked stylets, and contained partially developed pharynx. J3, J4, and young females were observed still enclosed in J2, J3, and J4 cuticles, respectively.

9.2.5

Green Mass Determination Assay

The green mass index was calculated using the height and width of the canopy as geometric variables. The height of the canopy was measured perpendicular to the soil, and since the width of the canopy varied with plant height, three 1/3 heights were defined to measure the mean width parallel to the soil. Thus, the green mass index was calculated in a similar way as previously described (Sánchez-Hermosilla et al. 2012) using the following equation: Green mass index ðcm2 Þ ¼

Total height ðcmÞ  ½width 1 ðcmÞ þ width 2 ðcmÞ þ width 3 ðcmÞ 3

Then, green mass percentages were calculated considering the control plants represented 100% (w/w ¼ 100%). Data from each treatment were recorded as average values of twenty 14-day-old-F1 plants and twenty F1 plants at 30 dai with Mj from two independent experiments.

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Gene Expression Analysis by Quantitative Real-Time PCR (qPCR)

cDNA was synthesized from 1 μg of total RNA, which was extracted from ten pooled root samples of three different F0 plants or ten pooled F1 plants per treatment using TRIZOL® (Invitrogen Life Technologies), purified with RNeasy MinElute Cleanup kit (Qiagen), and then used for reverse transcription with an oligo (dT) primer with the Transcriptor First Strand cDNA Synthesis kit (Takara Inc.), following the manufacturer’s protocol. cDNA from two independent experiments was pooled. Each cDNA was obtained from ten different F1 plants per treatment, and three technical replicates were analyzed by real-time PCR using an ABI PRISM 7000 Sequence Detection System (Applied Biosystems) with Brilliant SYBR Green QPCR Master Mix (Roche) in a total volume of 10 μl. All reactions were performed under previously described conditions (Medeiros et al. 2017). Ct values were calculated using the Applied Biosystems software, and transcript abundance was calculated in Microsoft Excel from Ct values and normalized to the actin gene signal. The relative expression levels of defense marker genes were analyzed by qPCR using specific primers (Medeiros et al. 2017) and the 2
ΔΔCT method (Livak and Schmittgen 2001).

9.2.7

Results

Nematode parameters (NG, NEM, NTNR, and the number of nematodes in the different developmental stages) from the F0 plants of the w/Mj, Ta/Mj, and w/Ta + Mj treatments, reported by Medeiros et al. (2017), revealed that in tomato Ta exerts a systemic protective effect against Mj. Particularly, the number of adult nematodes and NG developed in tomato roots was significantly reduced by Ta. Trichoderma-induced systemic defense was accompanied by increased levels of SA-dependent defense genes and oxidative burst in the split-root plants challenged with Ta. Two-week-old F1 plants of the five split-root tests showed different developmental degrees as calculated by the green mass values (Fig. 9.2): w/w (100%), w/Ta (130%), w/Mj (108%), Ta/Mj (22%), and w/Ta + Mj (125%). These results were indicative of a heritable tomato growth-promoting effect triggered by Ta in F0 plants. When these same F1 plants were inoculated with Mj, those derived from the Ta/Mj treatment resisted the RKN attack, measured by lower NG, NEM, and NEM/NG values than those from the untreated control, and also showed less growth and increased expression of SA- and JA-dependent genes. However, F1 plants from the w/Ta treatment were the highest but also showed significantly increased defense against Mj in terms of lower NG, NEM, and NEM/NEG values. These plants also displayed increased expression levels of genes related to ROS production.

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Fig. 9.2 Method followed to calculate the green mass values in tomato plants

9.3

width 1

1/3 height

width 2

1/3 height Total height

width 3

1/3 height

Discussion

The split-root methodology can be employed with plants of different species, ages, and sizes (Table 9.1). The major constraint of this approach may be due to the low flexibility and the small size of roots that can hinder the experimental procedure, for example, in the case of woody plant roots and liverwort rhizoids, respectively. Plants with a dominant taproot system (i.e., primary, swollen, or tubercular) are unable to be equitably separated in two pots. Nevertheless, there are references of split-root method application in cotton (Zhang et al. 2012), tea (Mishra et al. 2014), Arabidopsis (Martinuz et al. 2015), and woody plants such as grapevine (McKenry and Anwar 2007; Hao et al. 2012) or olive (Cabanas et al. 2017). Regarding the small size of roots, in some plants like Arabidopsis, this methodology has been successfully applied in in vitro assays using fungi against nematodes (Martinuz et al. 2015). As shown in Table 9.1, split-root methodology can be applied not only to Trichoderma (Khan et al. 2004; Moreno et al. 2009; Selim et al. 2014; MartinezMedina et al. 2017; Medeiros et al. 2017) but other biocontrol (Pochonia, Lecanicillium, Catenaria, avirulent or hypovirulent strains of Fusarium) (Larkin et al. 1996, Larkin and Fravel 1999; Monfort et al. 2005; Piskiewicz et al. 2009; Vu et al. 2006; Martinuz et al. 2015) and beneficial fungi such as mycorrhizae (Rhizophagus, Funneliformis, Glomus, Gigaspora) (Cordier et al. 1998; Pozo et al. 2002; De la Peña et al. 2006; Khaosaad et al. 2007; Elsen et al. 2008; Castellanos-Morales et al. 2011, 2012; Hao et al. 2012; Vos et al. 2012; Zhang and Franken 2014), endophytic basidiomycetes (Serendipita indica) (Cosme et al. 2016), and yeasts (Arxula) (Papasotiriou et al. 2013). The system has been widely implemented with biocontrol and plant growth-promoting rhizobacteria (Pseudomonas, Burkholderia, Sinorhizobium, Arthrobacter, Bacillus, Paenibacillus) and even nonpathogenic nematodes to explore plant systemic defenses against bacterial, fungal, oomycete, and nematode diseases. The defense induction by biological control agents belonging to bacterial and fungal species has been tested in semiwoody and woody plants such as cotton and olive, respectively, against the fungal pathogen Verticillium dahliae (Zhang et al.

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2012; Cabanas et al. 2017). Other studies including V. dahliae as target and using split roots have shown that biocontrol strains of bacteria, yeast, and hypovirulent filamentous fungi are able to induce systemic defense in vegetables like eggplant (Antonopoulos et al. 2008; Pantelides et al. 2009; Papasotiriou et al. 2013). Numerous works performed on the fibrous roots of monocotyledons have served to evaluate the ability of bacteria and mycorrhizal fungi against pathogens such as Gaeumannomyces graminis, Fusarium graminearum, and Pythium ultimum in barley and wheat plants (Aberra et al. 1998; Monfort et al. 2005; Khaosaad et al. 2007; Henkes et al. 2011; Jousset et al. 2011; Castellanos-Morales et al. 2012). However, split-root system has been widely used with tomato plants to test the biocontrol abilities of bacterial, fungal, nematode, and oomycete species against different kinds of attackers (Cordier et al. 1998; Pozo et al. 2002; Siddiqui and Shaukat 2002a, b, 2004; Zhu and Yao 2004; Dababat and Sikora (2007); Anwar and McKenry 2008; Moreno et al. 2009; Vos et al. 2012; Akram et al. 2013; Adam et al. 2014; Medeiros et al. 2017).

9.4

Conclusions and Outlook

We have demonstrated that the split-root methodology developed to study tomato responses to Trichoderma and RKN colonization is useful to explore the systemic signaling of plant defenses but also the heritable resistance induced by Trichoderma and the underlying mechanisms that support the beneficial activity of Trichoderma to plants in terms of growth and defense. Split-root methodology is a promising approach to advance our understanding of plant cross talk with the environment, the induction of systemic defenses against diseases and pests, the behavior of soilborne biocontrol agents and root colonizers, as well as comparative studies concerning microbial community structures and composition of rhizospheric and endophytic microbiomes induced by biotic and abiotic stimuli. Acknowledgments The research has been partially funded by the Spanish Ministry of Economy and Competitiveness for national project AGL2015-70671-C2-1-R and by the Regional Government of Castilla y León (Spain) for project SA009U16.

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Part III Experimental Approaches and Analytical Techniques in Rhizosphere Biology

Using a Tri-Isotope (13C, 15N, 33P) Labelling Method to Quantify Rhizodeposition

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Pierre Stevenel, E. Frossard, S. Abiven, I. M. Rao, F. Tamburini, and A. Oberson

Abstract

Belowground (BG) plant resource allocation, including roots and rhizodeposition, is a major source of soil organic matter. Knowledge on the amounts and turnover of BG carbon (C), nitrogen (N), and phosphorus (P) in soil is critical to the understanding of how these elements cycle in soil-plant system. However, the assumptions underlying the quantification and tracking of rhizodeposition using isotope labeling methods have hardly been tested. The main objectives of this chapter were to (i) review the different plant labeling techniques for each of the three elements; (ii) describe a novel method for the simultaneous investigation of C, N, and P rhizodeposition in sand; and (iii) test the methodological assumptions underlying quantification of rhizodeposition. Stable 13C and 15N isotopes were widely used to study rhizodeposition of plants either separately or in combination, while P radioisotopes (32P, 33P) were used to investigate root distribution. The combination of the 13CO2 single-pulse labeling with the simultaneous 15N and 33P cotton-wick stem feeding effectively labeled Canavalia brasiliensis roots and facilitated the estimation of rhizodeposited C, N, and P input from root systems. However, the isotope distribution was uneven within the root system for all three elements. Additionally, we observed a progressive translocation from shoot to roots for 15N and 33P over 15 days after P. Stevenel (*) · E. Frossard · F. Tamburini · A. Oberson Institute of Agricultural Sciences, Plant Nutrition Group, ETH Zurich, Zurich, Switzerland e-mail: [email protected] S. Abiven Department of Geography, Soil Science and Biogeography Unit, University of Zurich, Zurich, Switzerland I. M. Rao International Center for Tropical Agriculture (CIAT), Cali, Colombia Plant Polymer Research Unit, National Center for Agricultural Utilization Research, United States Department of Agriculture, Peoria, IL, USA © Springer Nature Singapore Pte Ltd. 2019 D. Reinhardt, A. K. Sharma (eds.), Methods in Rhizosphere Biology Research, Rhizosphere Biology, https://doi.org/10.1007/978-981-13-5767-1_10

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labeling, while the 13C tracer was diluted with newly assimilated non-enriched C compounds over time. Younger root sections also showed higher specific activities (33P/31P) than the older ones. The relatively high 33P radioactivity recovered in sand right away at the first sampling was attributed to an artifact generated by the stem feeding labeling method. Overall, our results suggest that the assumptions underlying the use of isotope methods for studying rhizodeposition are violated, which will affect the extent of quantification of rhizodeposition. The consequences of nonhomogeneous labeling of root segments of different age require further investigation. The use of a timeintegrated isotopic composition of the root is recommended to not only account for temporal variation of isotopes but also to improve the method of quantifying plant rhizodeposition.

10.1

Introduction

10.1.1 Significance of Plant Belowground C, N, and P Input in the Cycling of These Elements Belowground (BG) plant carbon (C), nitrogen (N), and phosphorus (P) inputs into the soil via roots and rhizodeposition play a key role in the cycling of these elements (Fig. 10.1). During their growth, plants release compounds through their root system (Neumann 2007), which can affect availability of nutrients and their cycling. This release is defined as rhizodeposition. Several definitions of rhizodeposition exist in the literature (Wichern et al. 2008). While some authors restrict rhizodeposition to

Fig. 10.1 Concept of belowground C, N, and P input

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release C from roots (Lynch and Whipps 1990; Meharg 1994; Nguyen 2003), others relate rhizodeposition also to other elements such as N (Janzen 1990; Mayer et al. 2003a). We will use the definition given by Uren (2001) who describes rhizodeposition as the release of all kinds of compounds lost from living plant roots, including ions and volatile compounds. Thus, contribution from belowground plant comprises both root biomass and rhizodeposition. Specifically for C, the additional respiration may be included in BG-C. Rhizodeposition can be divided into (i) lysates of sloughed-off cells and root debris and (ii) root exudates released from intact root cells including diffusates, secretions, and excretions (Neumann 2007). Knowledge on the amounts of BG-C, BG-N, and BG-P and their turnover in soil are critical to the understanding of cycles of these elements in soil-plant systems. Carbon rhizodeposition is a process of major importance for several reasons. Rhizodeposited C is (i) a net C loss and energy cost for plants, (ii) a direct C input to the soil, and (iii) an important microbial stimulation which increases the biological activity in the rhizosphere, especially regarding the enhancement of microbial nutrient turnover (Jones and Darrah 1996). A substantial proportion (20–60%) of C fixed during photosynthesis can be translocated belowground (Kuzyakov and Domanski 2000), where 1–25% of that C could end up as rhizodeposition. In a recent review, Pausch and Kuzyakov (2017) analyzed more than 280 data sets from literature on BG-C inputs under crops and grasslands. Even if results showed a higher C translocation to BG plant parts with grassland species (33% of total net fixed C) compared to crop species (21% of total net fixed C), the distribution of proportions of the C translocated belowground was similar in both cases. On average, 48% of the C translocated belowground was allocated to the root biomass, 36–38% was lost by roots as rhizosphere respiration, and 14–15% could be recovered as net rhizodeposition. This partitioning may change with environmental and management conditions affecting plant growth. Belowground plant inputs, comprising roots and rhizodeposition, also contain N taken up from the soil or, in case of N-fixing plants, N derived from the atmosphere (Janzen 1990; Mayer et al. 2003a). N2-fixing legumes have regularly been used in rhizodeposition studies, with the purpose to quantify the input into the soil of symbiotically fixed N. Thus, for legumes, rhizodeposition can provide a significant input of fixed N into the soil, which is often not accounted for in estimates of symbiotic N2 fixation. In a review, Wichern et al. (2008) reported that amounts of N rhizodeposits from experiments under controlled conditions ranged from 4% to 71% (median of 16%) of total assimilated plant N for legumes and from 4% to 56% (median of 14%) of total assimilated N for cereals. It appears that some studies overestimated the amount of N rhizodeposits because of the limitations of the methods used. Concerning BG-P, root biomass is a potential source of P for the following crop via decomposition and mineralization processes (Chen et al. 2002; Soon and Arshad 2002) since it can represent up to 50% of the total crop dry matter (Gregory 2006), substantially depending on plant species and growth stages. Phosphorus rhizodeposition is usually considered to be negligible (Foyjunnessa et al. 2014).

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However, a few studies reported losses of P from roots to the surrounding substrate (Emmert 1959; Pellet et al. 1996; Ritz and Newman 1985; Rovira and Bowen 1970). From 1 to 5% of 33P applied with foliar labeling was released into the rhizosphere of 22 days old wheat plants grown in rhizoboxes (McLaughlin et al. 1987).

10.1.2 Effect of Growth Conditions on BG-C, N, and P Input Amounts and quality of compounds released from roots vary with plant species, variety, and plant age as well as with abiotic and biotic factors influencing overall plant growth and root development (Dakora and Phillips 2002; Jones et al. 2004; Nguyen 2003). Plants are regularly exposed to a range of stresses at the root-soil interface, which influence root releases in response to soil parameters (Rovira 1969), nutrient deficiencies (Jones 1998), light intensity (Watt and Evans 1999), temperature (Bekkara et al. 1998), plant developmental stage (Aulakh et al. 2001), or CO2 elevation (De Graaff et al. 2007). The uptake of water and nutrients by roots is a major driving force for nutrient transfer and diffusion in the rhizosphere. Depending on the plant demand, depletion or accumulation of nutrients and solutes (including exudates and rhizodeposits) can occur in the rhizosphere via mass flow (Hinsinger et al. 2005). Depletion zones will induce plant mechanisms to acquire nutrient, which directly affect rhizodeposition. Belowground N was always around 40% of aboveground N in red clover growing over two seasons across a nutrient gradient in the field, but the proportions of rhizodeposited N on belowground N increased with time (Hammelehle et al. 2018). Exudation of low-molecular-weight organic acids is enhanced by many plants when P supply is severely limiting (Lambers et al. 2009; Louw-Gaume et al. 2017). Tropical forage legumes have developed strategies to acquire P from highly weathered soils with low available P contents by increasing P availability in their rhizosphere (Rao et al. 1999). Examples of such strategies include the secretion of organic anions or phosphatase enzymes, as reported for cowpea (Jemo et al. 2006). At the opposite, tropical grasses differ in their response to low P availability by developing an extensive root system and having a markedly higher P use efficiency (g dry matter produced per mg P uptake) than legumes (Rao et al. 1996). Such different plant mechanisms toward nutrient limitation will hence affect the quality of belowground C, N, and P inputs via the partitioning between roots and rhizodeposition but also via their nutrient ratios. Amounts and ratios of C, N, and P inputs to the soil are important drivers of soil microbial activity (Ehlers et al. 2010). The C:N:P ratios of plant residues affect the temporal dynamics of decomposition (Bünemann et al. 2004), microbial nutrient cycling (Oberson and Joner 2005), the release of plant available mineral N and P from these residues (Nziguheba et al. 2000; Vanlauwe et al. 1998), and also the microbial community composition (Ha et al. 2008). We therefore can expect that C: N:P ratios of belowground inputs will as well affect the cycling of these nutrients. For instance, low C:N and C:P ratios in root tissues result in fast decomposition and nutrient release (Gijsman et al. 1997). Jones et al. (2004) warned on the lack of the

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in-depth knowledge on the effect of the root exudate C:N ratio such as the balance of C supplied in sugars to N in amino acids, because the ratio will affect soil organic matter (SOM) cycling in the rhizosphere. Thus, amounts and ratios of nutrients contained in rhizodeposition need to be quantified, and their fate be studied simultaneously. Rhizodeposition may more likely determine the short-term nutrient turnover (rapid priming effect) than root decomposition. For instance, in the rhizosphere, root exudates can promote SOM turnover, even at very low amounts (μg g
1 soil) (De Nobili et al. 2001). Alternatively, at a given location, microorganisms will prefer to utilize the easily available rhizodeposition before the less available SOM, resulting in a decrease in SOM decomposition (Sparling et al. 1982). Including rhizodeposition could provide a more comprehensive picture on nutrient ratios of BG plant inputs in response to change in growth conditions. This calls for methods for simultaneously studying belowground input of C, N, and P. In the following chapter, we give an overview on methods currently used to study separately rhizodeposition of C, N, and P. We also show how methods can be combined for dual-isotope labeling investigations. We then discuss the methodological assumptions underlying quantification of rhizodeposition. Finally, we present a novel tri-isotope labeling approach to simultaneously study rhizodeposition of C, N, and P. While our results suggest that methods currently used to study BG input of elements do not meet the methods assumptions, we propose measures to obtain more accurate estimates of rhizodeposition.

10.2

Methods to Study Rhizodeposition

10.2.1 Method Overview While roots can be physically recovered from the soil, the quantification of rhizodeposition is commonly done with isotope enrichment techniques that enable to track plant-derived compounds released from roots into the soil. As outlined by Kuzyakov and Domanski (2000), rhizodeposition investigation is challenging mainly because of (i) low concentration of rhizodeposits compared to native soil substances, (ii) their fast decomposition, (iii) the restriction to a narrow zone of soil around the roots, and (iv) their chemical similarities to organic substances released by SOM decomposition and microbial turnover. Many reviews have been written on methods for quantitatively estimating C and N rhizodeposition from different agricultural crops and on investigating the fate of rhizodeposits in soil (Fustec et al. 2010; Kuzyakov and Domanski 2000; Kuzyakov and Schneckenberger 2004; Meharg 1994; Wichern et al. 2008). All these methods used for labeling plants may also influence the amount of predicted rhizodeposition (Mahieu et al. 2009; Meharg 1994; Merbach et al. 2000; Yasmin et al. 2006). Belowground C is usually labeled via the photosynthetic pathway using enriched CO2. Pulse or continuous labeling with 13C- or 14C-CO2 has been largely used to trace C compounds from the plant to the soil, such as root turnover and compound releases, usually using labeling chambers (Kong and Six 2010; Kuzyakov and

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Schneckenberger 2004). With pulse labeling, the dynamics of fresh assimilates can be traced, whereas continuous labeling gives more integrated results of C cycling in the soil-plant system, due to the more homogeneous labeling of the plants C pools and fluxes (Studer et al. 2014). Thus, pulse labeling provides information on C partitioning and C residence time in small C pools (Studer et al. 2014), distribution of C at a specific growth stage of plants, and kinetics of CO2 in soil-plant system (Kuzyakov and Schneckenberger 2004). Further, 13C repeated pulse labeling (i.e., successive periods of exposure to 13CO2 throughout plant growth) has been successfully used to determine the fate of BG-C in plant and soil pools (Bromand et al. 2001). In contrast to 13C labeling, most 15N enrichment techniques for determining the fate of N within the plant-soil system require plant manipulation (Fustec et al. 2010; Wichern et al. 2008). Approaches used include shoot fumigation (Janzen 1990), leaf feeding (McNeill et al. 1997; Palta et al. 1991), petiole feeding (Khan et al. 2002), stem feeding (Russell and Fillery 1996), and the split-root system technique (Sawatsky and Soper 1991). Only a few studies used 15N2 labeling techniques in order to estimate N fixation by legumes and the transfer of symbiotically fixed N to companion non-fixing plant (Lesuffleur et al. 2013; Ta et al. 1989; Warembourg et al. 1982). A commonly used labeling technique is the cotton-wick stem feeding where 15N-enriched urea is supplied to the plant stem via a wick soaking in a vial containing the labeled solution. For stem feeding, it is imperative that the plant used has a stem wide enough ( 2 mm) to avoid breaking the stem during the hole drilling prior to insertion of the wick. For plant species that are not suitable to stem feeding, the leaf feeding technique could be a good alternative for obtaining comparable results (Yasmin et al. 2006). More than five decades ago, a P radioisotope technique for determining the distribution and extent of living roots of annual crops was developed by Racz et al. (1964). Briefly, a P radioisotope (32P or 33P) was injected by means of a micro syringe into the stem of a plant, and its presence was determined in soil-root cores at different soil depths after a certain time. The activity measurements have been used to derive an active root profile in the soil when the results are calculated on the basis of relative distribution of counts within the soil sample volume (Racz et al. 1964). Thus the radiotracer methods are mainly appropriate to provide qualitative information on distribution of living roots such as gross root growth patterns and structure, as influenced by species, variety, soil type, and growth conditions (Abbott and Fraley 1991; Cholick et al. 1977; Halstead and Rennie 1965; Subbiah et al. 1968). Later, Foyjunnessa et al. (2014) tested the efficacy of 33P stem wick-feeding for labeling roots of canola (Brassica napus) and lupin (Lupinus angustifolius) in sandy soil and showed effective root labeling, facilitating the quantification of total P accumulation in plant root systems. Furthermore, they concluded in another experiment that P from canola belowground residues contributed up to 20% of the P uptake of seedling wheat as subsequent crop (Foyjunnessa et al. 2016). However, the contribution of P released from decomposing roots and from rhizodeposition was not distinguished in this study. To date, P radioisotopes have rarely been used for studying rhizodeposition. Emmert (1959) showed root loss of 32P to the P solutions

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of plants growing in a hydroponic system shortly after the 32P foliar application. In opposite, another study concluded negligible loss of 33P to soil via exudation from healthy growing roots of wheat plants even if 0.7% of total injected 33P was recovered in soil (McLaughlin et al. 1987). It has been suggested that most of the 33 P loss was due to isotopic exchange at the root cell membranes (Weigl 1968) and that loss of P by the roots to soil through exudation was negligible.

10.2.2 Method Assumptions Rhizodeposition is operationally defined by the plant-derived C, N, and P remaining in the soil after separation of roots (Fustec et al. 2010; Mayer et al. 2003a). In enrichment studies, the percentage of C or N in soil derived from rhizodeposition (% Cdfr or %Ndfr) has usually been quantified by relating the isotopic composition (IC) of soil to the IC of roots (Janzen and Bruinsma 1989): 13

%CdfR or%N dfR ¼ 13

C or 15 N atom%excess in soil  100 C or 15 N atom%excess in roots

ð10:1Þ

where 13C or 15N atom% excess are calculated from the isotope ratio in the labeled sample minus the isotope ratio in an unlabeled control sample. Usually the IC of total soil C and N concentrations are determined, but the approach as well applies to specific soil nutrient pools. This formula is also used for determining the rhizodeposition of P, with IC being the specific activity (33P/31P) of the respective pools. This root-soil isotope balance approach is based on three assumptions: (i) the IC of roots and rhizodeposition is identical (Janzen and Bruinsma 1989), (ii) the IC of roots is constant over time (Sawatsky and Soper 1991), and (iii) the IC is uniform within the root system (Jensen 1996). Several studies indicate that these assumptions may be violated under certain conditions. Pausch and Kuzyakov (2011) reported that the labeling with a single pulse of 14C-CO2 of Lolium perenne grown in soil did not yield homogeneously labeled roots but resulted in the presence of “hotspots” with high 14C activity. The highest 14C activity was detected at the root tips already at 6 h after the 14C assimilation, while the 14C tracer was concentrated along adventitious roots at 11 days after labeling. Indeed, as growing tissue, apical meristems of root tips are strong sinks for freshly assimilated C. This suggests that the apical meristem incorporates mainly unlabeled C when the 14C application is over, resulting in unlabeled tissue at the newly formed root tip. Regarding N rhizodeposition, Sierra et al. (2007) observed that root exudates of trees (Gliricidia sepium) 15N-labeled with leaf feeding were always depleted in 15N in relation to the roots, suggesting that the assumption on equal isotopic composition of root tissues and exudates was not met. Furthermore, Rasmussen et al. (2013) labeled white clover and perennial ryegrass with 15N-urea via leaf labeling and 14C either via a 14CO2 pulse labeling or with 14C-bicarbonate mixed with the 15N-urea solution through leaf labeling to evaluate the homogeneity of tracer distribution after two alternative labeling

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approaches. They observed a heterogeneous root incorporation of 14C (e.g., uneven root labeling) in both cases and a correlation between 15N and 14C enrichment, thus inducing high variations in 15N labeling within different root parts. Rasmussen (2011) therefore critically discussed the use of the isotope approach illustrating that root growth after a pulse labeling dilutes the enrichment of the root system resulting in lower enrichment in the root tissue and that reallocation of tracer from the roots to the shoot (e.g., regrowth after harvest) will deplete root C and N and change the root enrichment. By referring to Eq. 10.1, decrease with time of the enrichment of roots between labeling and sampling of the root system would result in overestimated rhizodeposition, because of the underestimation of the atom% excess in roots. Thus, one of the main factors influencing the rhizodeposition estimation is the distribution of the tracers within the root system. Additionally to the homogeneity problem, Gasser et al. (2015) observed the rapid appearance of 0.5% of the applied 15N through the root system within the first day after labeling by using leaf feeding labeling on clover with 15N-urea. Gasser et al. (2015) termed this rapid appearance of tracer in the soil as leakage, i.e., an artifact caused by the labeling technique, which leads to an overestimation of rhizodeposition. Recently, Hupe et al. (2016) compared the classical approach of Janzen and Bruinsma (1989) to an alternative shoot-root-soil mass balance approach, where recovered 15N in soil is related to the total 15N in plant and not only in roots, to quantify BGP-N in peas and assess the error associated with the classical calculation. Results showed that BGP-N calculated with a mass balance approach was significantly lower than with the classical calculation, suggesting substantial overestimation of N rhizodeposition in previous studies. Only a few studies investigated the P labeling homogeneity within the plant and release from the roots. While a few studies report that stem injected 32P moved rapidly into the root system and was uniformly distributed (Halstead and Rennie 1965; Vijayalakshmi and Dakshinamurti 1977), McLaughlin et al. (1987) concluded a nonuniform labeling in roots due to a faster 33P than 31P release from roots incubated after drying-rewetting. Racz et al. (1964) also suggested that 5-day equilibrium period is necessary to ensure that the injected 32P is distributed uniformly throughout the root system. However, this recommendation was based on the assumption that a constant activity in the whole root system would signify a homogeneous distribution of the tracer.

10.2.3 Net and Gross Rhizodeposition Gross rhizodeposition is defined as the total amount of organic and mineral compounds released by living roots into the soil (including those transformed by, e.g., microbial immobilization), whereas net rhizodeposition is the difference between gross rhizodeposition minus the part reabsorbed by plants and lost by gas emission such as CO2 and NH3. Indeed, Janzen (1990) observed that a proportion of labile root-derived organic N compounds can be quickly mineralized and reabsorbed by the plant. Plant reuptake can also occur when inorganic N is released directly

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from roots (Brophy and Heichel 1989). Thus, in an extreme case, a zero net rhizodeposition may be measured when all labeled rhizodeposits would have been reabsorbed, which may rarely happen because of rapid microbial immobilization of rhizodeposited N (Mayer et al. 2003a). In opposite, gross rhizodeposition may be determined when all 15N-labeled rhizodeposits remain in soil pools, and only unlabeled soil N is taken up by the plant (e.g., complete pool substitution), as discussed by Gasser et al. (2015). Thus, in most studies using enrichment techniques, net rhizodeposition (i.e., the amount of rhizodeposits that remained in the soil at harvest after separation of the roots) is commonly measured since plant reuptake is not considered in the approach of Janzen and Bruinsma (1989).

10.2.4 Experimental Conditions The study of rhizodeposition implies to work with a substrate suitable for recovery of the roots. To experimentally check the validity of the assumptions underlying isotope methods to quantify rhizodeposition, obtaining a soil or substrate solution may as well be needed (Gasser et al. 2015). When soils are used, e.g., to study the incorporation of rhizodeposited elements into soil microorganisms (Mayer et al. 2003b) or other nutrient pools (Mayer et al. 2004), then soil texture is an important criterion to facilitate physical root recovery. Indeed, the major error is due to the critical root-soil separation step where the extraction of all roots from soil, especially of fine roots