Handbook of Florists' Crops Diseases 9783319323749

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Fundamentals and Advances in Plant Problem Diagnostics Tim Schubert, Ayyamperumal Jeyaprakash, and Carrie Harmon

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . General Laboratory Diagnostic Techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Special Features Related to Ornamental Disease Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . An Argument for Continued Use of Fundamental Diagnostic Techniques . . . . . . . . . . . . . . . . 4.1 Attention to Details . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Discerning Indications of Abiotic Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Many Advanced Techniques Yield Only Binary Answers . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4 Interpretation Challenges with Genetic Sequencing Data . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5 The Complications of Host Predisposition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6 Balancing Diagnostic Costs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7 Diagnosticians Are the Major Ambassadors of Phytopathology . . . . . . . . . . . . . . . . . . . . The Challenges of Abiotic Disease Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A Universal Diagnostic Process . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1 Step One: Assemble a Comprehensive Syndrome Description . . . . . . . . . . . . . . . . . . . . . . 6.2 Step Two: Check for Spatial Patterns . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.3 Step Three: Establish the Disease Chronology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.4 Step Four: Assemble a Comprehensive History of the Horticultural Treatment of the Crop . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.5 Step Five: Consult Literature . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.6 Step Six: Begin Laboratory Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.7 Step Seven: Consolidate Information . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A Special Diagnostic Category: Indexing Plant Propagative Material for Cryptic Pathogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Diagnosis of Biotic Diseases: Direct Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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T. Schubert (*) • A. Jeyaprakash (*) Division of Plant Industry, Florida Department of Agriculture and Consumer Service, Gainesville, FL, USA e-mail: [email protected]; Timothy.Schubert@freshfromflorida.com; ayyamperumal. jeyaprakash@freshfromflorida.com C. Harmon (*) Department of Plant Pathology, University of Florida, Gainesville, FL, USA e-mail: clharmon@ufl.edu # Springer International Publishing Switzerland 2016 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florist's Crops Diseases, Handbook of Plant Disease Management, DOI 10.1007/978-3-319-32374-9_1-1

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8.1 Microscopic Methods for Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2 Microbiological Methods for Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.3 Serological Methods for Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.4 Metabolic Tests for Pathogen Identification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.5 Molecular Methods for Pathogen Detection and Identification . . . . . . . . . . . . . . . . . . . . . . 9 Indirect Methods of Plant Pathogen or Disease Detection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.1 Detection of Volatile Organic Compounds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.2 Spectral Imaging of Host Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10 Accurate and Timely Reporting of the Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract

Plant disease/problem diagnostics and pathogen detection are fundamental components of successful agriculture. Any corrective action for poor plant performance should arise out of thorough assessment and diagnosis of the problem(s), to be followed by appropriate management suggestions. Among the factors to consider in the choice of diagnostic techniques are speed, cost, accuracy, availability of technology, training, value of the crop, and likelihood of generating a successful management strategy. Many exciting new technologies are entering the field of diagnostics, and plant disease/problem diagnostics and pathogen detection are not exceptions. Among the favorites are molecular-based tests that rely on the specificity of gene sequences, serological tests in various formats, direct and indirect biosensor technologies, and spectral imaging. We have entered an era in which the entire biome both inside and outside a plant can be identified, diseased plants can be analyzed non-destructively, and the overall health of an entire crop is quickly discernible. Now we must learn how to interpret all that data and put it to practical use. In spite of the enormous potential of all these new developments, successful disease diagnosis will still depend fundamentally on basic skills that have been honed and perfected in practice for more than a century. Without the foundational information from thorough and focused observation on-site and access to all horticultural information and inputs for the crop in question, no amount of advanced technology can reliably correct for deficiencies in the primary steps in diagnosis. Keywords

Plant disease diagnosis • Plant problem diagnosis • Triage • Microscopy • ELISA • PCR

1

Introduction

When a crop fails to perform up to expectations, it is logical to turn to a diagnostic process to determine what is preventing the attainment of full potential. This chapter will explain the full spectrum of options for determining what might be limiting the growth of a particular plant while considering various factors that influence which

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tools are most suitable for the particular diagnosis in question. In an era of rapidly advancing modern technologies promising extraordinary accuracy and precision heretofore practically unimaginable, the wise deployment of the entire spectrum of available tools maximizes the accuracy and usefulness of the diagnosis for the client and the grower community with a minimum of expense in a reasonable amount of time. There are three caveats at the beginning: Firstly, for this discussion, we use two different terms or phrases to describe the process of figuring out what is wrong with a plant – plant problem diagnosis and plant disease diagnosis. It is not necessary for the reader to accept any strict definitions here, as the two can have nearly the same meaning, and that is how we intend to use them. We draw attention to both terms to make certain that the reader understands that we are including some techniques for the diagnosis of plant problems that have no biotic (living) causal agent, i.e., they are due to some abiotic (nonliving) cause. Some consider plant disorders caused by abiotic influences something other than disease, with the term “disease” restricted to plant disorders caused by living causal agents (giving some latitude for the viruses and viroids and perhaps prions which are incapable of life functions outside a host cell). Others classify these disorders as noninfectious diseases or physiological diseases. In this abiotic category are factors such as excesses or deficiencies of nutrients, water, or light; poor-quality water, soil/media, or air in the growing environment; temperatures outside optimal ranges for plant growth; mechanical damage; etc. Though some may prefer to make a distinction, we will be using plant problem diagnosis and plant disease diagnosis as essentially the same thing. Secondly, a further clarification of terms is helpful in our quest for plant health. Plant disease or plant problem diagnosis represents the final step or culmination of a process that is preceded first by plant disease surveillance, followed by plant disease or plant pathogen detection. The semantics here are important. A grower must engage in continual surveillance to monitor overall plant health. This answers the question, “Is the crop normal? Is it on schedule? Is there a problem?” Plant disease or plant pathogen detection is the process of following up on the findings of surveillance. It addresses the question, “Is there a problem?” by defining the nature of the problem found, and discovers what organisms or causal agents are possibly associated with the syndrome detected. Finally, plant disease or problem diagnosis is the term reserved for the process of assigning the blame for the problem detected in the surveillance. The techniques and technologies discussed in this chapter can help in one or all of these three steps (Stack and Fletcher 2007; Miller et al. 2009). Thirdly, many clients of plant disease diagnostic services automatically assume that the diagnosis takes place entirely or mostly in the lab using esoteric methods, specialized knowledge, and complicated machinery. Actually, overlooking the field aspects of the diagnosis can derail the entire process and yield faulty conclusions. Though we will start our discussion on lab techniques, at several points through the chapter, we will remind the reader of what must be accomplished in the field with the goal of reinforcing the idea that good diagnostics requires more than modern equipment and skilled operators. The indispensable detective work in the field at the time the sample is taken sets the stage for an accurate and holistic diagnosis that

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leads to optimal management decisions. Furthermore, very often the sample-taker does not participate in the lab work and vice versa, so good communication networks between all the parties (the client, the sample-taker, and the lab scientist) are vital.

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General Laboratory Diagnostic Techniques

In general, the techniques normally restricted to the laboratory and used for plant disease or plant problem diagnosis fall into these four major categories: • Microscopy – examining the afflicted plant’s tissues at various magnifications to visualize symptoms and evidence of any pathogen(s). This would include a hand lens, dissecting (reflected light) and compound (transmitted light) microscopy, and scanning and transmission electron microscopies. • Microbiology – using various culture methods to separate the pathogen from diseased host tissue to study it in isolation for identifying features. If a pathogen can be isolated from its host and grown in culture or, in the case of viruses and viroids, purified by various methods, proof of pathogenicity by reinoculation is possible using Koch’s postulates. Failure to recover or witness any organisms associated with the syndrome would push the diagnosis in the direction of an abiotic causal agent. • Serology – using antisera developed against specific antigens from suspect pathogens to identify the causal agent in whatever matrix it may be found. • Molecular biology – using nucleic acid visualization and sequencing to find identifying sequences in DNA or RNA extracts from isolated suspect pathogens or taken directly from samples of diseased tissues suspected of containing the pathogen. This technology has the sensitivity to detect latent disease and inform management decisions proactively (Michailides et al. 2005). These categories are what many clients in need of diagnostic services believe to be the major or even sole methods employed. Each is in a typical sequence of steps and in relative order of cost to perform, and each requires special equipment and training to be successful in their use and interpretation. As technology advances, some of the work formerly confined to the lab is being “field hardened” to permit quick deployment on-site (Boonham 2014). Examples are more sophisticated field microscopes, dipstick and lateral flow devices for serological detection of microbes, and field PCR units. These methods can be used separately or in combination to reach the desired level of specificity in the diagnosis. However, it is extremely important to remember that for the best diagnosis, much of the successful process takes place in the field, prior to sample processing in the lab. While still in the field at the time of sample taking, a diligent effort must be made to acquire background information about the problem. Further questions may arise during lab processing that will make it necessary to revisit the site, resample, discuss issues again with growers, etc. These frequently underappreciated, overlooked, or poorly performed primary aspects of the diagnostic process are covered in greater detail in Sect. 6.

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5

Special Features Related to Ornamental Disease Diagnosis

At the outset, it is important to note that plant problems in ornamental crops as opposed to food, fuel, or fiber crops present some unique features that need special consideration in the diagnostic context. First, the value of ornamental plants is aesthetic – based upon appearance. Perfection (or near perfection) in appearance can make a huge difference in profitability of the enterprise, so much so that addressing any factor(s) that prevents attainment of that aesthetic standard could require the utmost in diagnostic expertise and technology. Plus, specifically ornamental floral crops are by their very nature more fragile than even other ornamental plants such as foliage or woody ornamentals. Conversely, cosmetic imperfections that blemish the non-harvested portions of food, fuel, or fiber crops (or even the harvested portions to a degree) can be tolerated with little or no impact on yield or product value. (In making note of this fact, we must at the same time admit much room for improvement in consumer tolerance of minor blemishes in food products in order to feed the growing masses and still provide a fair economic return to the grower.) Calculating the amount of pest damage to tolerate on any crop with no loss in value is fundamental to good stewardship and maximizing profits. So, in the final analysis, that allowance for imperfections while remaining profitable tends to be much smaller for the ornamental crop, especially floral products. Furthermore, commerce involving the propagative material of ornamental plants often comes with a much greater plant pest regulatory risk than for food, fiber, and fuel crops because the ornamental commodity is frequently destined not for some devitalizing use such as consumption as food, manufacture into an inanimate object, or burned as fuel. Instead, it is planted in a new location for a continued existence, complete with all the pests and pathogens that accompanied it on its original journey. The default “apparently free of pests and diseases” standard for most routine interstate and international phytosanitary certification leaves considerable latitude for cryptic organisms to accompany and disperse from propagative material. This added risk potential makes high-level diagnostics especially appropriate in the ornamental plant arena. And lastly, high-value ornamental crops as a rule receive much more attention in the way of horticultural inputs, each one designed to maximize quality production at minimum cost in the shortest amount of time. Crops “pushed hard” in this manner are more apt to suffer abiotic disorders from such things as excesses in fertilizer or growth regulators, phytotoxic pesticide applications and/or mechanical injury from repotting and handling for merchandizing.

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An Argument for Continued Use of Fundamental Diagnostic Techniques

Before addressing what might be considered by many to be the main topic in this chapter (that of advancements in biotic disease diagnostic techniques), we consider it entirely fitting to present a strong case for continued employment of more fundamental standard diagnostic methods. For the most part, few diagnostic tests are

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necessarily outmoded or automatically unsuitable with the passage of time. Especially neglected in this category are macroscopic and microscopic diagnostic approaches. Several reasons support this position, and these are addressed singly and in order.

4.1

Attention to Details

First, one’s initial approach to a diagnostic puzzle should commence with fundamental and historically proven tactics which depend upon keen attention to all aspects of the disease syndrome. Unfortunately, in every aspect of modern life, forces work against a person’s ability to simply give attention, to focus intently on a subject, and to notice pertinent details (Crawford 2015). It is at this initial stage where disciplined and polished observational skills and inquiries pay great dividends and start the diagnostic quest in the right direction. Furthermore, in most cases, the sample collection process is performed by someone other than the diagnostician, so good collaboration and communication with the sample collector is critical. It is no exaggeration to assert that the sample collector is an absolutely essential participant in the diagnostic process. Every available means of modern communication (both real time and recorded) should be made available and used to help the lab personnel grasp the disease presentation in the field and guide the sample collection process to full advantage. Done properly, much information will be gathered in these early stages. Much of it will prove unhelpful and tangential to the final diagnosis, but there is no substitute for a thorough job at this stage. The essential clues will be distilled from that voluminous assemblage of seemingly random information. A sloppy job here will frequently alter the course of the entire diagnosis, and advanced techniques have little or no inherent value to correct the situation.

4.2

Discerning Indications of Abiotic Disease

Second, it is important to remember that abiotic causes are responsible for about half the samples submitted to a typical plant disease diagnostic clinic. The fact that ornamental plants are subjected to a high degree of deliberate manipulation by caretakers translates into a quite reasonable expectation that abiotic diseases might be even more common here. Assignment of an abiotic cause is, of course, based on our current fragmentary understanding of what can cause various symptoms in plants, but it would be unwise to assume that our understanding does not remain fragmentary even with advanced diagnostic tools at our disposal. Abiotic diseases seldom leave signs of the causal agent in the classic sense (a recognizable part of the causal agent left behind at the scene), as is more often the case with a pathogen/biotic disease. Only symptoms remain, and symptoms are notoriously general with multiple paths of causation. Genomic tools are not as valuable under such abiotic disease circumstances, though one day, transcriptome (RNA molecules present) or

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metabolomics (chemical fingerprints) characterization technologies might prove helpful in understanding how a syndrome developed, and that could yield clues as to what abiotic condition may be responsible (Adams 2013).

4.3

Many Advanced Techniques Yield Only Binary Answers

Third, many (not all) of the more advanced diagnostic techniques are limited to providing only yes/no answers inasmuch as they can only detect a single organism. These are more accurately deemed detection tools that contribute to a diagnosis and do not constitute a diagnosis in and of themselves. This is especially true of PCR techniques and also serological tests to a lesser degree. They are more suited and useful for diagnostic confirmation than for diagnostic exploration.

4.4

Interpretation Challenges with Genetic Sequencing Data

Fourth, more advanced nucleic acid sequencing techniques and metagenomics have enormous potential to reveal genetic clues about all the organisms present in a sample of diseased plant tissue. One must be careful in the interpretation of the data procured with sequencing methods however. Our understanding of apparently innocuous or beneficial endophytic and epiphytic organisms associated with plants is too rudimentary to jump to conclusions about causality for a plant disease syndrome. Furthermore, associating gene sequences of an organism and its ability to cause disease is still a loose and tenuous relationship for many pathogen (e.g., Fusarium oxysporum). Many surprises await us, and understanding the significance of the biome and microbiome associated with a diseased plant will probably be a long learning curve. The more traditional diagnostic skills of isolating a pathogen from diseased tissues offer a cleaner substrate to take to the lab for nucleic acid sequencing, but limitations in what can be successfully isolated from diseased tissues have been known at some level for a long time. The automatic bias against fastidious microbes as pathogens is something the diagnostician must consciously avoid. The more we learn about the plant biome/microbiome, the more we understand about the limitations of culturing or otherwise separating microbes from diseased plant tissue. Even though these limitations are greater than we may have imagined, it is still a good idea to attempt to isolate the suspected causal agent from the host tissues because the clarity of genomic analyses of the clean isolate is far greater than an extract taken directly from diseased plant tissues.

4.5

The Complications of Host Predisposition

Fifth, as diagnostic skills advance, it is becoming more apparent that, under certain (perhaps many) circumstances, predisposing abiotic events precede and may even be

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essential for biological invasions leading to plant disease development. Since plant tissues can behave something like ecosystems in miniature, it is not unreasonable to assume that abiotic predisposing events and colonization events could very well take place in a somewhat orderly succession. Although it is helpful to know the biotic agent responsible for the eventual symptoms, the most effective plant disease management techniques would recommend the avoidance or minimization of the predisposing conditions to lessen or prevent the biotic phase(s) of the disease altogether. Starting with metagenomics in such a situation would render a very complicated and incomplete diagnosis and probably generate an ineffective management scheme. And if preliminary genomic findings have any early message for growers and diagnosticians, we should set aside notions of genetic stability of pathogens and, as a consequence, discount any parallel belief that a sequencebased diagnostic method will reach perfect performance and permanence. Clearly, pathogens are much more dynamic and adaptable than once thought. Fortunately for both client and diagnostician, a 100% accurate diagnosis is not always necessary to yield a helpful (if incomplete) determination(s) and provide recommendations that improve plant health.

4.6

Balancing Diagnostic Costs

Sixth, it remains imperative that any clinical methods be commensurate with the goal of the diagnosis, triage in one sense of the word. Every sample does not call for advanced techniques. With no intent to diminish the importance of any diagnosis, clinics need to maximize the resources at their disposal to provide as much service to society as possible without limiting their capacity to respond to extraordinary circumstances with every tool possible when the need arises. Most plant problem clinics operate with some level of public support, so there is an obligation to serve any tax-paying client to the fullest extent possible and warranted. Charging a fee for diagnostic services to supplement operating budgets and recoup costs may make larger staff and more costly procedures possible, but those same fees can also interrupt vital links to the very community that can serve as front-line detectors and “citizen scientists” for early disease detection and effective intervention. These fragile links need nurturing, and diagnostic fees as such tend to be counterproductive. The balance between affordability, applicability, accuracy, and sensitivity must be decided jointly by the clinician and client while keeping all these competing factors in view. One trend that helps in this regard is that of illustrated electronic communications to initiate a diagnosis. Assuming good photographic skills on the part of the client or disease surveyor/scout, a good diagnostician should be able to recognize the routine problems and dispatch them with minimum expense and time to save resources for the major events that will require “all hands on deck” and full spectrum diagnostics. In short, when it comes to advanced techniques, discretion in deployment may be as important as capabilities.

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Diagnosticians Are the Major Ambassadors of Phytopathology

Seventh, related to the theme of how the costs of diagnostic services are met, we must further acknowledge that disease diagnostic services and the essential cooperative nature of this enterprise with the client are almost certainly the sole link between the phytopathology profession and most of the general public. That being the case, the diagnostic clinic is well advised to put its best foot forward for the reputation of the profession. Additionally, even though all life on planet Earth is based directly on plant health, the scarcity of providers of diagnostic plant pathology services contributes to the general obscurity of phytopathology in the public service arena. For comparison purposes, according to the US Bureau of Labor Statistics, the medical profession has in the neighborhood of 700,000 diagnostic physicians and surgeons and the veterinary profession about 70,000. Diagnostic phytopathologists, on the other hand, are very generously estimated to number only about 1000 nationwide based on the number of members in the American Phytopathological Society that participate in the Diagnostics Committee plus the estimated number of employees at state- and federally sponsored diagnostic labs and clinics around the country. Diagnosticians undoubtedly serve as the foremost ambassadors for the entire profession. Deciding on the method(s) to be used in a diagnostic project must take into account much more than cost and applicability but must also consider the representation of the profession in the public forum and the potential contributions to be made to the common good.

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The Challenges of Abiotic Disease Diagnosis

In many respects, the diagnostic approach for any disease, biotic or abiotic, will follow the same pathways. Exceptions are granted when the diagnostician knows a crop extremely well and is fully aware of all the horticultural inputs and their effects. Familiarity with common biotic diseases for the crop also helps narrow the diagnostic focus. Usually, however, a thorough abiotic disease diagnosis is a meticulous process of eliminating possible biotic causal agents, then resorting to attempts to match abiotic causal agents to the symptoms observed.

6

A Universal Diagnostic Process

Stepwise, here is a rational process for any disease diagnosis. We emphasize that much of a successful and accurate diagnostic process occurs before moving into the lab. It gathers enough information to evaluate both categories, biotic and abiotic:

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Step One: Assemble a Comprehensive Syndrome Description

Begin with a careful examination of the syndrome to record all symptoms plus any signs of a pathogen with the goal of fully describing the problem. Is the presentation actually abnormal? How should the crop appear at this stage in the production cycle? At this early juncture, it is essential that detailed record keeping be initiated and practiced throughout the diagnostic process. These documents are vital for legal disputes and for trace back – trace forward investigation.

6.2

Step Two: Check for Spatial Patterns

Look for any patterns in the symptom expression on both the individual plants and within the plant population as a whole. Do not confine the exam to just the single crop, but also evaluate neighboring flora, even weeds if they are present. Examine whole plants as much as possible, internally and externally both above and below ground, and be prepared to sacrifice representatives to supply samples for further lab work.

6.3

Step Three: Establish the Disease Chronology

Determine the chronology of symptom appearance and any changes in the syndrome over time. When did the syndrome start? Was the onset sudden? Do symptoms abate, stay the same, or get worse?

6.4

Step Four: Assemble a Comprehensive History of the Horticultural Treatment of the Crop

Find out as much as possible about how the crop has been grown. Is the crop new to the grower, a new variety, growing in a different structure, or has something changed in the horticultural regimen (physical and chemical qualities of the growing media, water quality and amount, fertility, light levels and quality, temperature, growth regulator use)? Pay particular attention to the mode of delivery of any horticultural treatments. Phytotoxic agents can reach a crop in any number of ways, including by direct and deliberate application, drift, contamination of pesticide or fertilizer product or application equipment, rate miscalculations, poor application techniques, contamination of soil or surroundings, mulch or soil amendments, in irrigation water or runoff, etc.

6.5

Step Five: Consult Literature

Check references on the plant in question to learn of any unusual sensitivities, preferred growing conditions, idiosyncrasies in growth cycles and plant appearance,

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and reported diseases and disorders. APS compendia and reference manuals such as this one are good sources of information (Chase et al. 1995; Daughtrey et al. 1995; Gill et al. 2006; Gleason et al. 2009; Horst and Cloyd 2007; Horst and Nelson 1997; Horst 2013).

6.6

Step Six: Begin Laboratory Analysis

Based on the information gathered, determine what laboratory testing might be warranted to attempt confirmation. (It is here that the more technical laboratory work introduced earlier in Sect. 1 and addressed in more detail in Sect. 8 usually begins). Determine whether any biotic agents discovered using laboratory techniques and coupled with symptoms and disease chronologies are primary or secondary. Soil and tissue analysis are advised for confirmation of symptoms associated with known syndromes of nutrient deficiencies/excesses or soil pH problems. A bioassay using indicator plant species can confirm problems with suspected harmful soil residues. Testing of pesticide products for foreign components or testing of plant tissues for residues of pesticides is much more involved and expensive, usually requiring some idea of the nature of the residue sought.

6.7

Step Seven: Consolidate Information

Review all the gathered information to synthesize a likely diagnosis. The process for an abiotic disease diagnosis can eventually assemble large amounts of information because it is fundamentally a process of elimination, and much of the facts gathered may end up having little or no bearing on the final conclusions. Nevertheless, the quality of the diagnosis depends on thoroughness of the overall investigation. As stated earlier, using the most advanced diagnostic tools available cannot make up for deficiencies in steps 6.1 through 6.5.

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A Special Diagnostic Category: Indexing Plant Propagative Material for Cryptic Pathogens

Indexing for plant pathogens in propagative material is a different sort of diagnostic challenge altogether. The most sensitive detection methods available deserve top consideration, especially if the propagative material is to be moved to a new location within a country or to another country and must meet the attendant phytosanitary requirements. The potential gains in overall plant health (and thus profitability) from indexing of asexually propagated perennial plants in particular can be substantial. The rewards for indexing annual plants are different but similarly valuable because the crop customarily starts from seed each year anyway. With annuals, indexed seed is the goal. Indexed seed and stock plants are extremely valuable, though they can be quite expensive to acquire and maintain. Furthermore, these costs come up front,

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before any actualization of profits. Rewards may or may not prove to be worth the investment. Costs of the alternative (imperfect regulatory protection in place of starting with clean stock) are seldom fully considered in any cost-benefit ratio calculations; indeed such values are difficult to commodify without advanced economic skills which are outside the circle of most growers’ acquaintances. Assigning costs to any of the parties involved, especially at the outset, is also controversial, but eventually the potential for a higher profit margin and premium price for the superior product could supply ample financial returns to cover the setup and day-to-day operating expenses. Acquiring the equipment for the most advanced indexing techniques is another matter. Prices for advanced sequencing equipment and subsequent data processing are becoming more affordable with the passage of time. Grants and sharing resources collaboratively with other professions needing similar services (e.g., medical/veterinary health and ecosystem services) may be a possible solution. Still, in a free market, there will always be a niche for the cheapest items of any commodity, so we should not fool ourselves into thinking that every grower will eagerly adopt propagation from high-quality indexed stock. Regulatory oversight and public funding to protect consumers and natural resources from the consequences of substandard plant material are still necessary and advisable.

8

Diagnosis of Biotic Diseases: Direct Methods

Historically, plant disease diagnosis has targeted direct detection of the organism (s) associated with a syndrome to arrive at a diagnosis. In the case of an abiotic disease, there may be no physical evidence of a causal agent to detect and obvious shortcoming. Still, direct methods can be used in a process of elimination toward an abiotic diagnosis, and some abiotic agents do indeed leave physical evidence amenable to direct methods. Lately, progress has been made in indirect methods of disease diagnosis. These are covered in Sect. 9.

8.1

Microscopic Methods for Diagnosis

The careful observation of diseased plants and tissues using various means of magnification is the oldest, potentially the quickest, lowest cost, and most accessible diagnostic method available. Unfortunately, in many respects, the more advanced microbiological, serological, and molecular methods have distracted diagnosticians from the inherent value of simple and thorough light microscopic techniques. Skill in microscopic methods definitely requires good equipment and the time and material for plenty of practice. An apprenticeship with an adept microscopist is a great advantage. The more one practices microscopy and the discriminating observation and attention skills required to spot signs of a pathogen, the more exciting discoveries appear. Some good advice when examining diseased plant tissues for clues about the pathogen: have patience and expect to discover useful information using the microscope, do not give up easily, and do not restrict your examination to just a

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few pieces of symptomatic tissue. Carefully examine all stages of the symptoms insofar as you can reconstruct a time frame for the syndrome. If you see nothing noteworthy upon initial examination, try a moist chamber incubation for a few days to encourage fungal sporulation, and then reexamine. Inducing fungal sporulation to permit microscopic examination might require many weeks and manipulation of incubation conditions in some cases. In the emerging era of identification at the molecular level, there remains a significant place for morphometric identifications based on reproductive structures (Hyde et al. 2010) using fungal identification literature (Barnett and Hunter 1998; Leslie et al. 2006; Siefert et al. 2011). Highresolution digital photographic equipment for both macroscopic and microscopic images allows the sharing and cataloguing of images with and by experts over the Internet, thereby enhancing the technique even further. Good light microscopic equipment for both reflected and transmitted light microscopy equipped with digital cameras can cost from $3,000 to $4,000 at the low end to as much as $15,000 at the high end. Electron microscopy provides even more imaging capacity which can prove especially useful for diagnosis of viral and viroid diseases. Virus and viroid particles can be fairly quickly detected in expressed sap dried down and negatively stained on a coated grid. Electron microscopes small enough for a lab bench or tabletop capable of both scanning and transmission electron microscopy can be obtained for around $175,000 to $200,000. However, the specialized training and consumables make this equipment a likely purchase only when in support of a larger program or one with an emphasis in virus diagnosis or morphologically supported taxonomy and systematics. Microscopic detection methods suffer from the sensitivity inherent in the process. It is impossible to examine large volumes of host tissues or the environment using a light microscope to find suspected targets. The sampling problem is compounded when using the electron microscope. Extraordinary observation skills, training, and experience can lessen the inefficiency to a degree.

8.2

Microbiological Methods for Diagnosis

Several excellent treatises and lab manuals for the microbiological techniques traditionally used for plant disease diagnosis are available (Burns 2009; Dhingra and Sinclair 1995; Dugan 2006; Schaad et al. 2001; Shurtleff and Averre 1997; Streets 1979;Tuite 1969; Waller et al. 2002) plus there are many methods described online at http://wiki.bugwood.org/Diagnosticians_cookbook. Bacterial differentiation to the genus level is possible with the use of a few semiselective culture media coupled with tests for oxidase, Gram reaction, anaerobic growth, and hypersensitive response. These tests are not particularly difficult, but selection of isolates to carry forward from streaked plant tissue is based on experience. Careful attention to the chemistry and recipes of the culture media will encourage success. One is reminded to use fresh solutions and young cultures for best results. A few useful additions to the diagnostician’s bench include

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semiselective media for fungi that can help to separate out oomycetes from true fungi, can encourage sporulation, and can be amended to identify fungicide resistance.

8.3

Serological Methods for Diagnosis

Serological methods rely on the specificity inherent between an antigen and antibody developed as they are naturally formed by the immune system after injecting the antigen into the bloodstream of an animal, usually a mammal. This specific recognition phenomenon can be exploited to great advantage to seek out pathogen-specific antigens in any matrix (expressed plant sap, ground tissue extracts, etc.) and coupled with stains for light and/or electron microscopy, spectrophotometry, etc. Antisera produced in this traditional fashion are polyclonal and a mixture of antibodies generated to recognize different parts of a surface protein molecule or antigen. Most antibodies are now produced using hybridoma technology in tissue cultured cells that can be kept indefinitely to produce more antisera as needed. Antisera produced via hybridoma technology are monoclonal and recognize as a specific part of the antigen. Immunodiagnostics are quite reliable depending of the selectivity and sensitivity of the antiserum. Several enzyme-linked immunosorbent assays (ELISAs) have been developed to detect pathogens, especially bacteria and viruses. However, this does require that a species-specific antibody is already available to perform ELISA. The collective commercially available catalog of ELISA assays that have been developed so far to diagnose selected viral and bacterial pathogens is limited, but the availability of commercially available positive and negative controls encourages standardization of testing and verification of results. Interpretation of results must be done with care. Sutula et al. (1986) discuss the potential pitfalls and how to avoid erroneous interpretation. Once set up, an ELISA assay can be performed rapidly, but the costs of superior antigen identification and antibody development plus signal attachment can be a constraint. Once all reagents are developed and validated, a 96-well plate ELISA assay format can be utilized to reduce cost when the test volume is high. A microplate reader will be required to perform ELISA. Some serological assays have been incorporated into a very handy single sample dipstick or a lateral flow format that can be employed even in the field when and where the samples are collected. Both types of antisera, polyclonal and monoclonal, are often used in double-antibody sandwich ELISA (DAS-ELISA) and the aforementioned dipstick or lateral flow device. Monoclonal antibodies are more expensive to produce but since they are highly specific to the pathogen antigen, can detect single strains of a virus or a bacterial species or potentially subspecies. Results between tests are highly reproducible. Polyclonal antibodies are produced as a mixture, which has the advantage of being less expensive to produce as well as detecting multiple strains of a virus or multiple species of a bacterial genus. However, they are more likely to cross-react with other proteins. The lower specificity of polyclonal antisera

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may have an additional advantage in that they may provide clues to help identify unknown or as-yet uncharacterized viruses. DAS-ELISA kits including all reagents and controls are available from many vendors globally, costing between $1–12 per test well in kit cost. This plate-based assay is scalable, making diagnosis of large numbers of samples relatively easy and decreasing cost per sample. Quick tests such as dipsticks and lateral flow devices are readily available but for a smaller number of pathogens than plate-based ELISA and cost $1–5 per test. The limits of detection using ELISA are 105 to 106 CFU/ml. Serological specificity has been adapted a step further by attaching the antibodies to a fluorescent marker that emits a visible signal when viewed under a specially equipped microscope upon binding with its antigen while illuminated with a particular wavelength of light, usually in the UV range. This immunofluorescent microscopy depends on the availability and specificity of the antiserum. Flow cytometry (or FCM), used sparingly in plant pathology, can also take advantage of immunofluorescence technologies to detect pathogen antigens in large volumes of individualized plant cells. Flow cytometry offers enormous potential for indexing plant tissues for cryptic pathogens or a genetic sequence of interest, targeting any antigen or genetic sequence for which a specific antibody can be devised and a fluorescent probe attached (D’Hondt et al. 2011). It is even possible to sort the scanned cells based on the signal detected, thus permitting further work specifically with the selected tissues. Pathogen detection and viability in environmental samples is another application of flow cytometry (Chitarra and van den Bulk 2003). The need to develop antibodies and affix a fluorochrome is unnecessary to make use of flow cytometry if the target cells have an innate signal fluorescence when illuminated by a particular wavelength of light from the laser light source in the instrument. A basic FCM instrument without sorting capabilities costs from approximately $40,000 to $100,000. The sensitivity of FCM is in the range of 104 CFU/ml.

8.4

Metabolic Tests for Pathogen Identification

Microplate-based metabolic assays such as Biolog ® have successfully guided bacterial diagnosis, and also limited fungal diagnosis of plant pathogens. This system records the ability of a given active culture to metabolize specific carbon sources; the resulting activity matrix is compared to a library of known responses for over 2000 bacteria and yeasts. A separate setup can identify a smaller number of filamentous fungi. Originally developed for assessment of bacterial mammalian pathogens, the current GenIII library includes many plant pathogens as well, and also tests for Gram reaction. However, the use of this technology to identify specific organisms may be limited to genus, with a suggestion as to the species. Additionally, this system is limited for obvious reasons to culturable organisms. But it requires little specialized expertise and can be piggy-backed onto a plate reader that can also be used for ELISA, creating some efficiency in equipment purchase and upkeep for both systems. Biolog ® can cost about $20 per sample, assuming the use of the lesscostly nonautomated equipment.

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Identification of bacteria and fungi via fatty acid methyl ester (FAME) analysis is exemplified by the SherlockTM Microbial ID system (MIDI). It is based on gas chromatography and, in disease diagnosis, has been most heavily used for identification of culturable aerobic plant pathogens. This well-plate-based assay can be fully automated to resolve the identification of a microbe and compare its profile to a large library of known organisms. As with Biolog, this system generally resolves bacteria to genus reliably and suggests species and subspecies. The MIDI system develops a dendrogram of relatedness with references in the library, as chosen by the user, and generates a similarity index to the closest match. The MIDI tests generally cost $30–50 per sample, with savings possible for increasing numbers of samples run together. Diagnosticians tend to couple data from Biolog ® or FAME tests with other sequence-based, cultural, or plant-based tests to confirm the identified organism is capable of causing the disease symptoms of the affected plant. One thing these two tests can provide that molecular detection such as PCR and sequencing cannot is the verification that the detected organism is viable. If any or all these traditional approaches fail to provide a complete and/or clear diagnosis, molecular diagnosis at DNA or RNA level (Sect. 8.5) is an appropriate next step.

8.5

Molecular Methods for Pathogen Detection and Identification

Plant pathogen identification has always been a challenging task requiring increasingly more sophisticated tools and techniques as our understanding of host-parasite relationships and pathogenicity advances. As stressed earlier, even with molecular methods, initial assessment still requires techniques of visual inspection to recognize the symptoms caused by the infection such as chlorosis, necrosis, stunting, distortion, dieback, etc. Properly targeted sampling and careful shipment of the selected plant parts or nucleic acid extracts to a diagnostic laboratory is required for a methodical analysis to identify the pathogens at genus, species, or strain level. Molecular analysis requires various pieces of equipment (water bath, refrigerator, freezer, microcentrifuge, tissue homogenizer, nucleic acid quantification equipment, PCR and real-time PCR machines, gel-documentation system, Sanger and next-generation sequencing platforms, sequence analysis software and computers), funding for the lab supplies (extraction kits, reagents, pipettes tips and tubes, etc.), and a skilled technician to do the work. Some labs charge a price for every sample submitted for analysis to cover the operational cost. Other labs cover costs using grants from grower associations and local and federal governments, thus providing diagnostic help without any additional charge to the sample submitter. Cost estimates are given under the individual subject headings.

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8.5.1 DNA and RNA Extractions The starting material for DNA and RNA extraction varies depending on the pathogen. Harvested bacterial/fungal cultures grown on plates or finely ground up infected plant leaves, stem, or root tissues (100–200 mg) are extracted using spin-column kits or chemical purification such as CTAB methods kit to obtain genomic DNA. Elution using ultrapure, PCR-grade water (100–200 μl) is always preferred because the elution buffers provided in many extraction kits contain enzyme inhibitors which could interfere with downstream applications. The majority of the viral diseases are caused by RNA viruses, and their detection will require extracting total RNA from infected plant tissues (100–200 mg) using a kit such as a QIAGEN RNeasy Plant Mini Kit. A microcentrifuge is required to perform this task. The nucleic acids can be stored at
20 C until ready for analysis. The cost for the QIAGEN column is reasonable ($2–3). The quality of the DNA and RNA extracted must be quite good for successful downstream applications. 8.5.2 Molecular Identification The precise strategy used for identification of plant pathogens can vary depending on the organism and the information available about the pathogen. The NIH GenBank ® database (www.ncbi.nlm.nih.gov/genbank) has nucleic acid sequences for numerous well-known species, and the database is growing constantly as scientists from all over the world deposit sequences. Several million sequence deposits already exist in the GenBank, and that includes sequences from a large number of important plant pathogens. There is a good chance that the sequence information about a suspected plant pathogen could be already present in the GenBank. Sequence information for rare plant pathogens may not exist in the GenBank, so this would require using universal primers (ribosomal RNA or mitochondrial COX1) or degenerate primers (β-tubulin, actin, histone, etc.) to amplify a marker sequence by PCR and perform phylogenetic analysis to match to a closely related species for identification. Therefore, a strategy for molecular identification has to be selected carefully. 8.5.3

Fungal Species Identification

Species-Specific PCR Positive identification of plant-associated fungi is tremendously aided by modern genetic analysis methods. One must remember, however, that simply identifying a plant-associated fungus based on a genetic sequence is not strictly the equivalent of a disease diagnosis, though it can be a step in the process. Pathogenicity testing will be required to confirm the virulence of any isolates from the diseased tissues. Proving pathogenicity can be a simple or very complicated process which may need to take into account groups of organisms which work in concert in a chronological set or physiological sequence. Furthermore, environmental predisposition of the host to achieve a susceptible state can be complicated to determine and recreate. If a fungal plant pathogen is suspected to be a particular species based on morphology and culture characteristics, but a confirmation is required, then the

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sequences can be obtained from the GenBank for that species, and species-specific primers can be designed using CLUSTAL W sequence alignment generated by comparing to all closely related species and then selecting a unique sequence sites to design a species-specific primer. A straightforward PCR assay can be performed, and the size of the DNA band amplified should match the sequence information already existing in the GenBank, thus providing confirmation. Further confirmation can be derived by digesting the PCR products with a restriction enzyme and developing a banding pattern to detect unique restriction fragment length polymorphisms (PCR-RFLP), which should match the expected sizes. This approach has an added advantage, as it can be used for plant pathogens that are difficult to culture because the DNA extracted from infected plant could be used for PCR. The primers and a PCR test per sample could cost around $20. A PCR machine and an electrophoresis gel system would be required. A basic gradient block PCR thermocycler can be purchased for as little as $5000; gel electrophoresis equipment can be $12,000 or more, depending on the camera and size of the setup. Sequencing Several nuclear or mitochondrial marker sequences can be used to identify plant pathogenic and other plant-associated fungi. This strategy requires using universal primers or degenerate primers to amplify a specific nuclear or a mitochondrial marker, sequencing, and then comparing the sequences to those already existing in the GenBank using a BLAST search tool. The markers generally used for fungal taxonomy are as follows. Ribosomal RNA The ribosomal RNA region (18S-ITS1-5.8S-ITS2-28S) has been well investigated from numerous fungal pathogens. 18S rRNA, 28S rRNA, and ITS1-ITS2 spacers are the markers that are extensively investigated from fungal pathogens. The ITS1 and ITS2 spacers are known to accumulate mutations, and it is often possible to find unique species-specific sequences in this region to generate phylogenetic trees for identification or to design species-specific primers. The ITS region is multicopy, but surprisingly all the copies that are present in a fungal species turned out to be identical. This was exploited by fungal taxonomists to build a rich collection of sequence information in the GenBank database. Universal primers are available that amplify well from both known and unknown fungal cultures grown on media (Table 1) (White et al. 1990). Culturing on a plate is necessary before performing this procedure to eliminate all the other background fungi present in the diseased tissue. Other Markers Mitochondrial cytochrome oxidase 1 (COX1), the nuclear elongation factor-1 α (EF1α), β-tubulin (Tub), histone (His), etc. could be used as well, but the sequence information library is not as rich for fungal species as it is for the ribosomal sequences. Multilocus sequence analysis (MLSA) involving several different markers greatly improves fungal species identification. The PCR products can be

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Table 1 Generic primers for amplifying fungal and bacterial pathogen DNA Organism Fungi

Marker 18S ITS

28S

Bacteria

16S

Primer NS1 NS8 ITS1 ITS4 ITS5 LROR LR5 LR7 27f 1495r

Sequence 50 –30 GTA GTC ATA TGC TTG TCT C TCC GCA GGT CAC CTA CGG A TCC GTA GGT GAA CCT GCG G TCC TCC GCT TAT TGA TAT GC GGA AGT AAA AGT CGT AAC AAG G ACC CGC TGA ACT TAA GC TCC TGA GGG AAA CTT CG TAC TAC CAC CAA GAT CT GAG AGT TTG ATC CTG GCT CAG CTA CGG CTA CCT TGT TAC GA

shipped to a commercial sequencing company or university core laboratory to obtain the sequence information. The cost for sequencing PCR products or cloned inserts is steadily decreasing, around $10 per sequencing run. Multiple sequence runs as well as sequencing both forward and reverse strands would greatly improve sequence accuracy and provide a clear identification. Occasionally, diagnostic labs do have their own Sanger sequencing machine (Applied Biosystems 3130XL Genetic Analyzer), with equipment costs starting around $150,000. Real-Time PCR The conventional standard PCR procedure can be quite labor-intensive, is prone to aerosol contamination, and is good for screening only a few samples. If the sample size is large, then a real-time PCR (also called quantitative PCR or qPCR) assay, such as one based on TaqMan technology, is ideal. This does require two primers (forward and reverse) and a species-specific fluorescent probe (6FAM or TET). This technology exploits the Taq DNA polymerase exonuclease activity, which shreds the probe bound to the species-specific site releasing the fluorescent marker and producing a positive result (Ct value 16–32). The probe fails to bind other non-target species DNA due to mismatches, producing a negative result (Ct value 32–40). The Ct values 30 % (Garcés de Granada et al. 1994). Symptoms/signs. During rooting cuttings, the pathogen can cause partial or complete decay of the plantings resulting in an irregular and uneven rooting, affecting the quality of the cuttings (Fig. 5). Also, it has been found that the fungus

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Fig. 5 Pythium rot of cuttings. Soft rot is spread by contact of affected leaves (SM Wolcan)

in adult plants behaves as a minor pathogen, causing a reduction in plant growth and root activity with a decrease in yield and quality of flowers. Disease symptoms are more evident in soils with a history of continuous Gypsophila cropping (Galindo and Arbeláez 1995; Gamliel et al. 1993). Biology and epidemiology. Refer to the information presented in ▶ Diseases of Carnation, Sect. 2.10. A high cutting density in the rooting benches, overwatering, and high humidity in the propagative greenhouses favor the production and spread of basal rot in the cuttings or rooted cuttings (Fig. 5). Management • Cultural practices – Avoid high relative humidity and overwatering in the rooting trays as well as in production plots. Use well-drained soils. Control weeds inside and outside of greenhouses to keep better air circulation among plants and avoid possible alternative hosts of Pythium species. Also, avoid transplanting rooted cuttings with dark roots. • Biological control – The application of Trichoderma harzianum isolate T17 increased the number of flower by 26.0 % and 15.6 % compared with an untreated control and commercial control, respectively (Galindo and Arbeláez 1995; Sivan et al. 1984). Some species of Pythium act as mycoparasites. Pythium oligandrum, Pythium acanthicum, and Pythium periplocum are well known as parasites of P. irregulare and P. ultimum (Van der Plaats-Niterink 1981). • Fungicides – Mefenoxam is helpful when used in preventative programs (Koike et al. 2009). Gould (2012) recommends the use of etridiazole, mefenoxam, and metalaxyl applied as a soil drench during propagation and also at transplanting. Metalaxyl produced a 11.4 % increase in flower bunches compared with the untreated control (Galindo and Arbeláez 1995). • Soil disinfestation – Solarization: Gamliel et al. (1993) used clear, 40–50-μmthick polyethylene mulch to solarize both infested soils and substrates infested

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Fig. 6 Symptoms of dry and shredded rot in a rhizome of G. paniculata caused by Rhizoctonia solani (SM Wolcan)

with Pythium spp. and Rhizoctonia solani for 40–55 d (max temperature 45–48  C) and obtained increases in total yield of flowers (flower weight and number) and extended the cropping time. Fumigation: In the same experiment, metham sodium applied at a rate of 120 ml/m2 via sprinkler irrigation produced the same results.

2.7

Rhizoctonia Rot and Rhizoctonia Stem Base Rot [Rhizoctonia solani Kühn Teleomorph: Thanatephorus cucumeris (Frank) Donk]

Geographic occurrence and impact. The disease has been reported in Argentina, Brazil, Israel, Korea, and Poland (Bueno et al. 2013; Elad et al. 2014; Kang et al. 2009; Werner and Antkowiak 2002; Wolcan et al. 2007). Symptoms/signs. Refer to the information on Rhizoctonia in Sect. 2.12 of ▶ Diseases of Carnation. In the field, symptoms of Gypsophila plants affected by R. solani may be mistaken for those produced by Phytophthora crown rot. In the case of Rhizoctonia, the stems are not easily pulled out. A dry and shredded appearance is observed when the rotted stem or the rhizome are split longitudinally (Fig. 6). Biology and epidemiology. Refer to Sect. 2.12 of ▶ Diseases of Carnation. Rhizoctonia solani populations are divided into anastomosis groups, and according to studies of Bueno et al. (2013) in Brazil, the pathogen from Gypsophila corresponds to the AG-4 HG III group.

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Management • Cultural practices – Refer to Sect. 2.12 of ▶ Diseases of Carnation. • Fungicides – Gould (2012) recommends the use of iprodione, iprodione + thiophanate-methyl, and thiophanate-methyl + etridiazole applied by soil drench at the seedling stage and transplanting. Treat soil or the planting medium with granular PCNB before planting. Spray the base of plants before or after planting with iprodione or PCNB. Keep PCNB off foliage (Koike et al. 2009). • Biological control – Bacillus subtilis var. amyloliquefaciens strain FZB24 is recommended by Gould (2012) to be applied as a soil and transplant drench. • Soil disinfestation – Solarization: Gamliel et al. (1993) reduced disease caused by Rhizoctonia and Pythium and increased yield in Gypsophila using soil solarization (see Management of Pythium above). Fumigation: In the same experiment, metham sodium applied at a rate of 120 ml/m2 via sprinkler irrigation produced the same results. Additional fungal diseases – The following fungal pathogens of Gypsophila spp. have also been reported (see management of the same pathogen genera in the “▶ Diseases of Carnation”): – Alternaria dianthi: in Mexico (USDA, ARS, US National Fungus Collections (BPI) 1946). (http://mycoportal.org/portal/collections/individual/index.php?occid=1350477) – Alternaria dianthicola Neergaard: in Poland (Werner and Antkowiak 2002). – Alternaria mold on flowers (Alternaria axiaeriisporifera E.G. Simmons & C.F. Hill). This species was recorded in New Zealand (Simmons 2007; Woudenberg et al. 2013). – Downy mildew (Peronospora gypsophilae Jacz., Peronospora pulveracea Fuckel). P. gypsophilae was recorded in Israel (Ben-Ze’ev et al. 2006) and P. pulveracea in Ecuador (Puma Quinchuango 2010). – Phoma leaf spot – Phoma gypsophilae Hollós was reported in Armenia (Simonyan 1981) and Phoma pinoggii Servazzi in Italy (Servazzi 1942). – Phyllosticta leaf spot – Phyllosticta sp. was reported in the USA (Alfieri et al. 1984). – Rust – Puccinia arenariae (Schumach.) G. Winter was reported on G. paniculata in Africa and on G. elegans in Chile, Germany, Portugal, and the UK (Farr and Rossman 2016). – White mold – Sclerotinia sclerotiorum (Lib.) de Bary, Sclerotinia serica M.A. Keay. S. sclerotiorum was reported on G. paniculata in Israel and the USA (Elad et al. 2014; Koike et al. 2009), and S. serica was reported in the UK on G. elegans (Keay 1937).

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3

Bacterial and Phytoplasma Diseases

3.1

Crown Gall [Agrobacterium tumefaciens (Smith and Townsend 1907) Conn (Formerly Bacterium tumefaciens, Pseudomonas tumefaciens, Bacillus tumefaciens, and Phytomonas tumefaciens)]

Geographic occurrence and impact. Crown gall is one of the most widely distributed bacterial diseases and has been reported from all five continents (Hayward and Waterston 1965; CMI 1976). Symptoms/signs. Crown gall tumors are observed most often at the base of the stems and below ground on the main roots. Once initiated, tumors continue to growth into large, friable masses that dissociate into small pieces as the tumor ages and dries (Kado 2010). Biology and epidemiology. Agrobacterium tumefaciens is a soilborne bacterium that infects dicotyledonous plants from over 90 different plant families. Crown gall tumors are the culmination of A. tumefaciens plasmid-mediated transformation of cells surrounding the infection court. Infection courts where bacteria proliferate are established mainly through injuries caused by mechanical trauma, grafting, freezing temperatures, and root-invading insects. High-moisture levels favor infections leading to transformation which take place in less than 3 h. Transformed cells proliferate into visibly abnormal tumors. As the tumors grow, A. tumefaciens is shed into the surrounding soil, particularly when rain or sprinkler irrigation occurs (Kado 2010). A. tumefaciens naturally resides on the rhizoplane of woody and herbaceous weeds. Its presence in soils originates from galls that were broken or sloughed off from infected plants during cultivation practices. Irrigation aids in further dissemination of the A. tumefaciens bacterial cells. Agrobacterium tumefaciens is also spread by infected and infested planting material originating from nursery stock from uncertified sources. Diagnosis is based on isolation in selective and semi-selective media (Schaad et al. 2001), bioassays, and PCR-specific detection (Haas et al. 1995). Management – refer to ▶ Diseases of Carnation, Sect. 3.4.

3.2

Fasciation [Rhodococcus fascians (Tilford 1936) Goodfellow 1984], also Called Leafy Gall

Geographic occurrence and impact. R. fascians is widely distributed and has been reported in Australia, Belgium, Canada, former Czechoslovakia, Denmark, Egypt, France, Germany, Hungary, Iran, Mexico, the Netherlands, New Zealand, Sweden, Russia, the UK, and the USA (Bradbury 1986; Putnam and Miller 2007). Symptoms/signs. Excessive and abnormal proliferation of the leaves, flowers, and shoots, including gall formation on leaves and at wounds on the stems as a result of the hyperplastic growth of the affected tissues (Kado 2010). Biology and epidemiology. Refer to ▶ Diseases of Carnation, Sect. 3.5. Management. Refer to ▶ Diseases of Carnation, Sect. 3.5.

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Gypsophila Gall, also Named Crown and Root Gall Disease of Gypsophila [Pantoea agglomerans pv. gypsophilae (Brown 1932), Pantoea agglomerans pv. gypsophilae (Formerly, Erwinia herbicola pv. gypsophilae, Agrobacterium gypsophilae, Phytomonas gypsophilae, Pseudomonas gypsophilae, and Bacterium gypsophilae)]

Geographic occurrence and impact. The host range of Pantoea agglomerans pv. gypsophilae is restricted to Gypsophila spp., and it has been reported in Israel, Italy, the Netherlands, Japan, Scotland, Taiwan, and the USA (Bradbury 1986; Kado 2010); whereas, P. agglomerans pv. betae is pathogenic on Beta vulgaris and Gypsophila (Manulis and Barash 2003). Symptoms. Galls are formed, particularly, on the base of the stem. Symptoms resemble those of crown gall disease (Fig. 7). The galls may show water-soaked areas. The galls weaken the plants and cause defoliation and ultimately plant death (Manulis et al. 1998). Biology and Epidemiology. The cut ends of stem cuttings are the sites of entry and colonization by P. a. pv. gypsophilae, f and gall formation. Galled Gypsophila cuttings are the primary sources of inoculum; bacteria are disseminated via cutting tools and splashing water (Kado 2010). Apparently the bacterium moves from the galls to the shoots via the xylem vessels (Manulis and Barash 2003). The bacterium can be reliably detected by Nested PCR or Bio-PCR in symptomless Gypsophila cuttings as reported by Manulis and co-workers (Manulis et al. 1998).

Fig. 7 Symptoms of Gypsophila gall on: cutting (left) and plants (center and right) (Courtesy of S Manulis-Sasson, Department of Plant Pathology and Weed Research, ARO The Volcani Center, Bet Dagan, Israel)

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Management • Cultural practices – The only control measure is by the production of pathogenfree planting material through culture indexing, followed by strict sanitation practices. The practice of placing freshly cut, bundled shoots in water effectively disseminates the pathogen to all cuttings (Kado 2010). Neither resistant clones nor effective chemical treatments are available (Manulis et al. 1998). • Sanitation – Avoid recycling irrigation water because it can serve as a reservoir of the pathogen.

3.4

Phytoplasma Diseases

Geographic occurrence and impact. Aster yellows disease of Gypsophila has been reported in the USA (USDA 1960) and Canada (Northover 2007). Another phytoplasma disease affecting Gypsophila has been identified in Israel (Gera et al. 2007) and Poland (Kaminska et al. 1996). Phytoplasmas have a very broad and extensive host range covering at least 60 families (Kado 2010) indicating a lack of specificity. Symptoms/signs. Aster yellows disease symptoms begin with a general yellowing of the foliage followed by growth distortions, such as the loss of apical dominance that results in shoot and floral proliferation into a witches’ broom appearance. The yellowing leads to a general decline of the plant, culminating in death. Phyllody and alteration of tissue pigments (purple, yellow, red, and bronze) have been observed in some hosts (Kado 2010). Symptoms of stunting and shoot proliferation (witches’ broom or “asparagus fern”) and poor flower set have been observed in Gypsophila sp. in Israel (Gera et al. 2007) and Poland (Kaminska et al. 1996). Biology and epidemiology. Phytoplasmas are members of the bacterial class Mollicutes which are Gram-negative pleomorphic cells without walls. Currently, they are not cultivable in vitro and are considered obligate parasites that reside in the plant phloem. Phytoplasmas are able to replicate in the plant host and in the insects that vector them. In the USA and Canada, C. phytoplasma spp. are vectored by the aster leafhopper or six-spotted leafhopper (Macrosteles quadrilineatus = M. fascifrons). In Israel, three leafhopper vectors of phytoplasmas were identified: Orosius orientalis, Circulifer haematoceps, and C. tenellus (Gera et al. 2007). The universal primers P1/P7 for PCR reaction have been developed for phytoplasma’s detection (Deng and Hiruki 1991). Management. Management of phytoplasma diseases is based on monitoring and reduction of insect vector populations through chemicals or biological means and vector exclusion through the use of fine mesh screening. Once a plant is infected, there is no curative treatment. Crop rotation is ineffective for phytoplasma management due to their wide host range. Market considerations must be made for phytoplasma disease management, and management strategies employed in certain crops will not necessarily be feasible in others.

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4

Viral Diseases

4.1

Nepoviruses

Tomato ringspot virus (ToRSV) Geographic occurrence and impact. The disease was reported in Lithuania (Samuitiené and Navalinskienné 2001). Symptoms/signs. Infection reduces the plant growth, yield, and aesthetic quality. The main symptoms are plant stunting, malformation of leaves and flowers, chlorotic and necrotic spots and streaks, ring spots on leaves, vein necrosis, and shortening inducing leaf crinkling. When the infection is severe, plants may die early (Samuitiené and Navalinskienné 2001). Biology and epidemiology. Transmission of ToRSV is by nematodes (Forer and Stouffer 1982) and vegetative propagation from infected plants. Also, pollen and seeds may disseminate the virus (Kahn 1956). Disease is widespread in field perennial flowers that may serve as reservoirs for viruses and a source of infection for other crops (Rosenberger et al. 1983). Management. Propagation of selected healthy planting material, monitoring of plants for the symptoms, and elimination of affected plants. Control nematode populations by soil disinfestation through fumigation, soil solarization or steam treatment, or the use of chemical nematicides. Crop rotations can sometimes reduce nematode populations. The use of virus-free stock plants obtained by heat treatment and meristem culture ensures the start of a healthy plantation (Samuitiené and Navalinskienné 2001).

4.2

Cucumoviruses

Cucumber mosaic virus (CMV) Geographic occurrence and impact. The disease was reported in Korea (Park and Choi 2009). Symptoms/signs. Produces mosaic symptoms on Gypsophila paniculata (Park and Choi 2009). Biology and epidemiology. The virus is transmitted by more than 60 species of aphids, notably Aphis gossypii and Myzus persicae (Francki et al. 1979). Seed transmission has been reported in more than 20 plant species, with varying efficiencies from 150 plant genera and to survive as mycelium in plant debris (Booth and Waterston 1964). McGovern et al. (2003) indicated that F. avenaceum could survive on Styrofoam transplant trays and as an endophyte in the root systems of a number of ornamentals including wax begonia (Begonia x semperflorens-cultorum), carnation (Dianthus caryophyllus), exacum (Exacum affine), Gebera daisy (Gerbera jamesonii), Madagascar periwinkle (Catharanthus roseus), French marigold (Tagetes patula), pansy (Viola tricolor subsp. hortensis), petunia (Petunia x hybrida), rudbeckia (Rudbeckia sp.), salvia (Salvia sp.), and verbena (Verbena sp.). In addition, isolates of the fungus from several other hosts were pathogenic to

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Fig. 6 Fusarium crown and stem rot symptoms/signs: progression of symptoms (left to right) (left); mass of orange conidia (arrow) in stem lesion (right) (R. J. McGovern)

lisianthus, indicating that F. avenaceum may be pathogenic on lisianthus regardless of its phylogenetic origin (Nalim et al. 2009). Seed infection by F. avenaceum has been reported in a number of hosts (Booth and Waterston 1964), but the fungus was not isolated from lisianthus seed (McGovern et al. 2003; Pecchia et al. 2000). Short-distance aerial movement of macroconidia of F. avenaceum in a lisianthus transplant production house in California and two lisianthus cut-flower facilities in Florida was documented (McGovern et al. 2003; Seijo et al. 2000). The fungus is readily spread over long distances via infected transplants, and serious outbreaks of Fusarium crown and stem rot in cut-flower production facilities in the USA were attributed to infected transplants resulting from propagation in reused transplant trays without prior disinfestation (McGovern et al. 2003). The fungus can also be spread by fungus gnats (Bradysia spp.), shore flies (Scatella spp.), and moth flies (Psychoda spp.) (McGovern and Harbaugh 1997; El-Hamalawi and Stanghellini 2005). Common feeding sites for fungus gnats in lisianthus are roots at the plant base which are covered by decomposing leaves closely appressed to the growing medium; these sites provide moist infection courts for F. avenaceum. The fungus may also be spread from plant to plant on cutting tools (McGovern et al. 2003).

2.6.4 Management • Cultural practices – Use pathogen-free transplants. Avoid overwatering to prevent the buildup of fungus gnat and shore fly populations and control these vectors by biological or chemical means. • Sanitation – Avoid reusing transplant trays without thorough chemical or steam disinfestation; an alternative is the use of disposable plastic inserts that fit within

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Styrofoam trays. Disinfest cutting tools between plants. Locate cull piles away and downwind from production facilities. Eliminate weeds. Disinfest soil between crops using chemical (fumigation) or physical measures (steam or soil solarization). • Fungicides – McGovern et al. (2002a) reported that preventive application of azoxystrobin reduced disease incidence and plant mortality to ~20% and 0%, respectively, in a growth chamber trial; fluazinam, fludioxonil, myclobutanil, and thiophanate methyl were generally less effective. In a greenhouse experiment in the Netherlands, van der Wurff and Hamelink (2007) found that fludioxonil + cyprodinil and trifloxystrobin were effective in reducing Fusarium crown and stem rot. • Resistance – Harbaugh and McGovern (2000) evaluated the susceptibility of 46 lisianthus cvs. under high disease pressure in growth chamber experiments and found that the lowest disease frequencies 55 days after inoculation with F. avenaceum occurred in “Ventura Deep Blue” and “Hallelujah Purple” (25%), “Bridal Pink” (23%), and “Heidi Pure White” (53%), representing the blue/ purple, pink, and white flower color groups, respectively.

2.7

Fusarium Wilt (Fusarium oxysporum (Schlechtend):Fr. f. sp. eustomae)

2.7.1 Geographic Occurrence and Impact Fusarium wilt of lisianthus has occurred in Ecuador, Israel, Italy, Japan, Korea, Poland, the Netherlands, the UK, and the USA (de Werd 2003; Elad et al. 2014; Hahm 1998; McGovern unpublished data; O’Neil and Green 2010; Orlikowski 2001; Raabe 1991; Rapetti et al. 2002; Tomita et al. 2004). Losses of 50–70% observed at a lisianthus cut-flower facility in the Netherlands were attributed to F. oxysporum f. sp. eustomae possibly in combination with Pythium irregulare (de Werd 2003). Losses of 10–50% and 100% were observed at cut-flower sites in Ecuador and the USA (California), respectively (McGovern, unpublished data). Fusarium wilt was reported to be widespread throughout the ROK especially in alpine production locations where it occurred at incidences of 5–30% (Hahm et al. 1998). Up to 40% crop loss occurred in Poland (Orlikowski 2001).

2.7.2 Symptoms/Signs The F. oxysporum f. sp. eustomae infects and causes a dark discoloration of roots and the stem base. The pathogen moves upward through the xylem causing yellowing of leaves and wilting; vascular discoloration is visible when infected stems are cut lengthwise. As the disease progresses white masses of spores (sporodochia) are produced on stems, the leaves become tan, and plants wilt entirely and die (Fig. 7). The pathogen may also cause pre- and postemergence damping-off.

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Fig. 7 Fusarium wilt symptoms/signs: root and basal stem rot and white masses of conidia in sporodochia (arrow) on the stem of a wilted plant (R. J. McGovern)

2.7.3 Biology and Epidemiology Fusarium oxysporum f. sp. eustomae produces three types of asexual spores: microconidia and macroconidia which enable aerial dissemination and thick-walled, stress-resistant chlamydospores which enable long-term survival. Survival by the fungus in association with plant debris similar to other formae speciales of F. oxysporum is very likely. The fungus has also been shown to survive as a symptomless endophyte in the roots of Aubrieta sp., rapeseed (Brassica napus), and stock (Matthiola incana) (O’Neil 2006a). The sexual stage of F. oxysporum has not been reported. The disease is favored by warm temperatures (20–28  C/ 68–82  F) (Bertoldo et al. 2015; McGovern et al. 2002b). Genetic and virulence variability of the pathogen was reported in Italy (Bertoldo et al. 2015), and a PCR assay was developed which enabled its detection in the roots and stems of infected but symptomless plants (Li et al. 2010). 2.7.4 Management • Biological control – A commercial formulation which combines Bacillus sp., Pseudomonas sp., Streptomyces sp., and Trichoderma sp. Consistently reduced plant mortality from F. oxysporum f. sp. eustomae in growth chamber

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•

• •

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experiments (McGovern et al. 2002c; McGovern unpublished data). The biocontrol Trichoderma lignorum significantly reduced the incidence of damping-off caused by Pythium sp. and F. oxysporum and increased grower profitability in research in commercial facilities in Ecuador (Sacoto Bravo 2009). Fungicides – In controlled environment experiments, fludioxonil and triflumizole were very effective in reducing both the incidence of and mortality due to F. oxysporum f. sp. eustomae; azoxystrobin, myclobutanil, thiophanate methyl, and trifloxystrobin were less effective, and thiophanate + chlorothalonil was ineffective (McGovern et al. 2002b, c). Chase (2011) reported that the severity of the disease was significantly reduced by the fungicide metconazole and the inducer of systemic acquired resistance, acibenzolar-S-methyl. Soil disinfestation – Preplant soil disinfestation is an important tool in the management of Fusarium wilt. Failure to fumigate the soil between successive lisianthus crops and inadequate fumigant distribution due to high soil compaction led to serious outbreaks of the disease in the USA (California) and Colombia, respectively (McGovern, unpublished data). O’Neil and Green (2010) reported that sheet steaming (>80  C/176  F for 10 h), a steam plough (>80  C/176  F for 1 h), chloropicrin applied via drip line irrigation, dazomet incorporated into the soil, and formaldehyde and metam sodium drenched onto the soil significantly reduced but did not eliminate the incidence of Fusarium sp. in buried, naturally infected roots and stems of lisianthus in commercial greenhouse experiments in the UK; calcium cyanamide incorporation was ineffective. Preplant steaming and application of metam sodium and dazomet provided acceptable control of Fusarium sp. for cut-flower production in Argentina (Salles et al. 2001). Resistance – Greenhouse trials conducted in Italy indicated that the lisianthus cvs. Mariachi Green and Echo Dream Yellow were partially resistant to the disease (Gilardi et al. 2006). Integrative strategies – O’Neil and Green (2007) found that the efficacy of sheet steaming in reducing the viability of F. oxysporum was not improved by combination with a number of biocontrols (Gliocladium catenulatum, nonpathogenic F. oxysporum, Trichoderma harzianum) or fungicides (calcium cyanamide, carbendazim).

2.8

Phomopsis Stem and Leaf Blight (Phomopsis sp.)

2.8.1 Geographic Occurrence and Impact The disease occurred in the USA (Florida) where losses ranging from 4% to 80% were observed in two potted lisianthus production facilities (McGovern et al. 2000a). 2.8.2 Symptoms/Signs Stem necrosis is rapidly followed by leaf blight and the production of numerous dark pycnidia in diseased tissue. As the stem blight progresses, infected plants collapse and die (Fig. 8).

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Fig. 8 Phomopsis stem and leaf blight symptoms/signs. Note the production of numerous black pycnidia (arrow) in infected tissue (R. J. McGovern)

2.8.3 Biology and Epidemiology The fungus is spread by spores and infected plants and survives as pycnidia and mycelium in infected tissue. 2.8.4 Management Refer to the introductory chapters on integrated disease management.

2.9

Powdery Mildew [Leveillula taurica (Lév.) G. Arnaud (Anamorph: Oidiopsis taurica (Lev.) E. S. Salmon)]; Oidium sp.

2.9.1 Geographic Occurrence and Impact Powdery mildew of lisianthus has been reported to be caused by L. taurica (Oidiopsis spp.) in Brazil, Israel, Poland, Spain, Venezuela, and the USA and by Oidium sp. in Argentina and Japan (Cabrera et al. 2009; Cedeño et al. 2009; Elad et al. 2007; Koike et al. 1995; Melgares de Aquilar Cormenzana 1996; Okamoto et al. 2002; Orlikowski 2001; Reis et al. 2007). Genetic analysis should elucidate the taxonomic relationships among the various powdery mildew pathogens of lisianthus. 2.9.2 Symptoms/Signs Symptoms and signs include yellow blotches on both the upper and lower leaf surfaces and stems which become covered with profuse white fungal growth (mycelium, conidiophores, and conidia). Floral infection and leaf distortion and premature abscission may also occur (Fig. 9).

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Fig. 9 Powdery mildew symptoms/signs: yellow blotches on leaves covered with profuse, white fungal growth (mycelium, conidiophores, and conidia) (R. J. McGovern)

2.9.3 Biology and Epidemiology Only the anamorphic stage of powdery mildew fungi has been observed on lisianthus; the role of the teleomorphic stage is currently unknown. The fungus infects the epidermis but, unlike most powdery mildew pathogens, it also colonizes mesophyllic tissue by means of stomata through which conidiophores subsequently emerge. Elad et al. (2007) found that conidial germination of O. taurica was optimal at 20  C/68  F and a relative humidity of 75–85% and that severe infection occurred at 15–20  C/59–68  F in pepper research in Israel; prolonged temperatures above 25  C/77  F decreased disease severity. Conidia of the fungus are spread by air currents. It has been suggested that L. taurica is a composite species consisting of a number of host-specific races (Braun 1987, 1995). However, Correll et al. (1987) found that L. taurica isolates from a number of different genera and families could cause powdery mildew in tomato (Solanum lycopericum). Okamoto et al. (2002) demonstrated that an isolate of Oidium sp. from 4 o’clock flower (Mirabilis jalapa) could cause powdery mildew symptoms on both lisianthus and broad bean (Vicia faba). Reis et al. (2007) reported that in addition to lisianthus, nasturtium (Tropaeolum majus), calla lily (Zantedeschia aethiopica), impatiens (Impatiens balsamina), and balloon plant (Asclepias physocarpa) were hosts of Oidiopsis. 2.9.4 Management The endoparasitic nature of Leveillula taurica makes it more difficult to control by contact fungicides than ectoparasitic powdery mildew pathogens which infect host tissue more superficially. • Cultural practices – In greenhouse research on pepper (Capsicum annuum) in Israel, Elad et al. (2007) found that powdery mildew caused by Leveillula taurica was decreased by increasing nighttime temperatures by heating and daytime temperatures by manipulation of the greenhouse vents. • Fungicides – Numerous fungicides and a number of biocontrols are available for management of powdery mildew (refer to chapter 6 “Fungicides and Biopesticides for Florists’ Crops Disease Management”). Fungicides should be rotated by mode of action and applied preventively.

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Fig. 10 Pythium root rot symptoms: discolored roots (left), rapid wilting (right) (R. J. McGovern)

2.10

Pythium Root Rot (Pythium spp.)

2.10.1 Geographic Occurrence and Impact Pythium root rot in lisianthus may be caused by a number of Pythium species including P. irregulare, P. myriotylum, P. spinosum, as well as unidentified species of the pathogen and has been reported in Ecuador, Israel, Japan, Norway, Spain, and the USA (Aegerter et al. 2002; Elad et al. 2014; McGovern unpublished data; Melgares de Aquilar Cormenzana 1996; Sacoto Bravo 2009; Tomita et al. 2004; Toppe 2005). Losses exceeding 75% occurred in greenhouse in the USA (California) (Aegerter et al. 2002).

2.10.2 Symptoms/Signs The root systems become darkly discolored and rotted, and infected plants rapidly wilt. The outer rotted and discolored layer of Pythium-infected roots is often easily pulled off leaving behind the central, lighter-colored stringy root fiber (stele). The pathogen may also cause pre- and postemergence damping-off (Fig. 10).

2.10.3 Biology and Epidemiology The genus Pythium is closely related to Phytophthora and also is fungus-like but not a true fungus. The pathogen is disseminated as zoospores through irrigation, surface water, and rain, and long-term survival is enabled by chlamydospores and sexually produced oospores. Pythium root rot is common in waterlogged soils with poor aeration; cycles of under- and overwatering which lead to root damage are very conducive to outbreaks of Pythium. Fungus gnats (Bradysia spp.) create infection

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sites through their root-feeding and may disseminate the pathogen. Pythium, depending on the species, may have a very broad host range.

2.10.4 Management • Biological control – The biocontrol Trichoderma lignorum significantly reduced the incidence of damping-off caused by Pythium sp. and F. oxysporum and increased grower profitability in research in commercial facilities in Ecuador (Sacoto Bravo 2009). • Fungicides – A number of isolates of P. irregulare recovered from lisianthus in the USA (California) were found to be resistant to the commonly used fungicide mefenoxam (Aegerter et al. 2002).

2.11

Rhizoctonia Crown and Stem Rot Blight [Rhizoctonia solani Kühn (Teleomorph: Thanatephorus cucumeris (A.B. Frank) Donk)]

2.11.1 Geographic Occurrence and Impact Rhizoctonia root, crown, and stem rot is a major plant disease and has been documented in lisianthus in Israel, Japan, Spain, and the USA (Alfieri et al. 1994; Meir et al. 2010; Melgares de Aquilar Cormenzana 1996; Yoshimatsu 1993). Tomita et al. (2004) observed 2.8–31.6% losses due to damping-off in the field from R. solani in Japan. 2.11.2 Symptoms/Signs Rhizoctonia solani can infect the roots, stems, and foliage of lisianthus. The fungus most typically produces a dark discoloration and rot in stems at the soil line, leading to wilting and rapid collapse of the entire plant. Under humid conditions, mycelial growth of R. solani may rapidly envelop and blight leaves and shoots; this form of the disease is known as Rhizoctonia aerial blight. Discrete leaf lesions may have a concentric appearance. The pathogen may also cause pre- and postemergence damping-off (Fig. 11). 2.11.3 Biology and Epidemiology Rhizoctonia solani has a very large host range which encompasses most economically important cultivated plants including ornamentals and weeds; but some host specialization has been demonstrated (Mordue 1974). The fungus is very active at warm temperatures and effectively colonizes and survives as hyphae and sclerotia in plant debris, soil, and other growing media. Although R. solani does not produce spores, it can be spread by infected propagative material including seeds, water splash, and airborne particulate matter. The relationship of the sexual stage of R. solani to plant disease has not been well studied.

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Fig. 11 Rhizoctonia crown and stem rot and foliar blight symptoms (R. J. McGovern)

2.11.4 Management • Biological control – Meir et al. (2010) observed a consistent though nonsignificant increase in lisianthus survival in soil infested by both F. solani and R. solani by prior inoculation with the arbuscular mycorrhizal fungus Rhizophagus intraradices (formerly Glomus intraradices). • Soil disinfestation – Chloropicrin applied by chemigation (through drip irrigation tubing) resulted in significant but minor reduction of R. solani soil densities in greenhouse research in the UK (O’Neil 2006b).

2.12

Thielaviopsis Black Root Rot [Thielaviopsis basicola (Berk. and Broome) Ferraris (1912)]

2.12.1 Geographic Occurrence and Impact The disease has occurred in Canada, Switzerland, and the USA (Florida) (Joshi 2000; McGovern unpublished data, Michel 2015). A loss of ca. 70% occurred with one planting of a single cultivar in Florida. 2.12.2 Symptoms/Signs A black discoloration occurs in the small, feeder roots, or these roots are entirely absent. Infected plants are stunted, turn yellow, wilt, and die (Fig. 12). 2.12.3 Biology and Epidemiology Thielaviopsis basicola produces black, thick-walled, resistive chlamydospores (also referred to as macroconidia), and microconidia, which are involved in its survival and dissemination. The fungus is worldwide in distribution especially in cool, wet climates and infects a broad range of cultivated plants in many unrelated families.

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Fig. 12 Thielaviopsis black root rot symptoms: note the black discoloration of small, feeder roots (R. J. McGovern)

Black root rot is most severe in cool, wet soils (17–23  C/62–73  F) and at a soil pH above 5.5 (Shew and Lucas 1991).

2.12.4 Management • Cultural practices – The disease can be reduced by lowering the soil pH to below 5.5 if feasible, and crop rotation. 2.12.5 Additional Fungal Diseases The following fungal diseases of lisianthus have also been reported: – Alternaria blight (Alternaria sp.) (Israel, Auscher 1997) – Brown root rot [Subplenodomus drobnjacensis (Bubák) Gruyter, Aveskamp, and Verkley 2012] (Japan, Kondo et al. 2014) – Curvularia leaf blotch (Curvularia sp.) (USA, Jones and Harbaugh 1995) – Myrothecium leaf spot and blight (Myrothecium roridum Tode) (The Netherlands, Ludeking and Arkesteijn 2011) – Penicillium root rot (Penicillium sp.) (Japan, Tomita et al. 2004) – Phytophthora root rot and stem blight (Phytophthora spp.) (Japan, Uematsu et al. 1996) – Phyllosticta leaf spot (Phyllosticta sp.) (USA, Daughtrey 2000) – Sclerophoma stem blight (Sclerophoma eustomis Taubenhaus and Ezekiel) (USA, Taubenhaus and Ezekiel 1935) – Sclerotium stem blight [Sclerotium rolfsii Sac. (teleomorph: Athelia rolfsii (Curzi) C.C. Tu and Kimbr.) (USA, Japan, McGovern et al. 2000b; Tomita et al. 2004) – Sclerotinia stem rot [Sclerotinia sclerotiorum (Lib.) de Bary] (Wolcan et al. 1996) – Stemphylium blight (Stemphylium lycopersici Wollr.) (Venezuela, Cedeño et al. 2011) – White rust [Albugo swertiae (Bed. et Komm.) Wilson (Dingley 1969)] (New Zealand, McKenzie 1987) – White blister rust (Pustula centaurii (Hansf.) Thines, C. Rost et Y. J. Choi, comb. nov, MB519564) (Australia, Tasmania, Ploch et al. 2011)

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For an up-to-date list of fungi associated with lisianthus, access http://nt.ars-grin. gov/fungaldatabases/fungushost/FungusHost.cfm and enter “Eustoma” in the host name box.

3

Bacterial and Phytoplasma Diseases

3.1

Bacterial Crown Rot [Burkholderia gladioli (Severini 1913) Yabuuchi et al. 1993, comb. nov.]

3.1.1 Geographic Occurrence and Impact The disease has been reported in Poland and the USA (Schollenberger and Zamorski 2008; Seijo et al. 2002). 3.1.2 Symptoms/Signs The pathogen can infect plants of various ages ranging from small seedlings to flowering plants and causes tan veinal necrosis in the leaves, rot in the tap root, crown, and stem, and wilting and death (Fig. 13). 3.1.3 Biology and Epidemiology Burkholderia gladioli (formerly Pseudomonas gladioli) can infect a number of plant hosts including gladiolus (Gladiolus x hortulanus), onion (Allium cepa), and mushroom (Agaricus bitorquis), and based on its host range was divided into three pathovars: pv. gladioli, pv. alliicola, and pv. agaricicola, respectively (Young et al. 1996). However, Burkholderia gladioli from lisianthus has not yet been characterized to the level of pathovar. The bacterium has also been isolated from the pulmonary tissue of cystic fibrosis patients (Beringer and Appleman 2000). The disease is favored by warm temperatures (28 °C/82 °F), and the bacterium can be seed-borne (Seijo et al 2002). Whitby et al. (2000) developed a species-specific PCR

Fig. 13 Bacterial crown rot symptoms: crown and stem rot and veinal necrosis (left) wilting and stem collapse (right) (R. J. McGovern)

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assay that distinguishes between B. gladioli and a number of other closely related species such as B. caryophylli which causes similar symptoms in lisianthus.

3.1.4 Management • Cultural practices – Use pathogen-free seed, and avoid unnecessarily wounding plants. Rapidly remove infected plants.

3.2

Burkholderia Wilt [Burkholderia caryophylli (Burkholder 1942) Yabuuchi et al. 1993, comb. nov.]

3.2.1 Geographic Occurrence and Impact This disease has occurred in Japan (Furuya et al. 2000). 3.2.2 Symptoms/Signs The symptoms are similar to those caused by Ralstonia solanacearum (see next disease). The pathogen causes root rot and yellowing, wilting, and browning of the foliage through destruction of the water-conducting tissue; a cross section of infected stems reveals a typical tan to yellow-brown discoloration of the cortical tissue. The lower leaves are affected first. 3.2.3 Biology and Epidemiology Burkholderia caryophylli (formerly Pseudomonas caryophylli) also causes rot in carnation, onion, and statice (Limonium sinuatum) (Ballard et al. 1970; Jones and Engelhard 1984; Palleroni 1984). Shao et al. (2011) developed a real-time fluorescent PCR assay for specific detection of B. caryophylli. 3.2.4 Management • Cultural practices – Rapidly remove infected plants.

3.3

Ralstonia Wilt (Ralstonia solanacearum (Smith 1896) Yabuuchi et al. 1996, comb. nov.)

3.3.1 Geographic Occurrence and Impact Bacterial wilt caused by Ralstonia solanacearum (formerly Pseudomonas solanacearum) is one of the most important bacterial diseases of cultivated plants and has been reported to occur in hundreds of plant species representing more than 40 families (Buddenhagen and Kelman 1964). This disease of lisianthus has occurred in Japan and Taiwan (Chao et al. 1995; Tomita et al. 2004). 3.3.2 Symptoms/Signs Disease symptoms include wilting of lower leaves and then the entire plant, followed by browning of the foliage and plant death. In cross section, infected stems exhibit a

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characteristic brown discoloration in cortical tissue; infected stems also feel hollow when pressed.

3.3.3 Biology and Epidemiology Five groups (races) of Ralstonia solanacearum have been described based on host range (Buddenhagen 1986; Buddenhagen et al. 1962). Thus far, all strains of the bacterium in Taiwan have been classified as race 1, including the strain that infects lisianthus (Lee et al 2001). Race 1 infects many solanaceous crops, ornamentals, and weeds and has a very high temperature optimum (35–37  C/95–98  F). The bacterium can survive in the soil in association with plant debris (Sugawara and Ishii 2009). In carnation the pathogen has been reported to move from one cutting to another in irrigation water during propagation, but plant to plant transmission appeared to be slow and limited to neighboring plants (Bradbury 1973). Lee et al. (2001) have developed a method to detect race 1 strains of R. solanacearum by PCR. 3.3.4 Management • Cultural practices – Sugawara and Ishii (2009) recommended the avoidance of successive plantings of lisianthus, rapid removal of infected plants, disinfestation of soil and equipment, and improvement of drainage.

3.4

Phytoplasma Diseases

3.4.1 Geographic Occurrence and Impact Diseases of lisianthus caused by phytoplasmas have been reported to occur in Brazil and Israel (Rivas et al. 2000; Weintraub et al. 2007). However, it is likely that additional reports of phytoplasma infection of lisianthus will be forthcoming given the global distribution of both host and pathogen. 3.4.2 Symptoms/Signs Symptoms caused by phytoplasmas may include yellowing of foliage, stunting and proliferation of shoots (witch’s brooms), and the abnormal development of floral parts into leafy structures (phyllody) and of green coloration in flowers (virescence). 3.4.3 Biology and Epidemiology Phytoplasmas are prokaryotes that lack cell walls and are obligate plant parasites vectored by phloem-feeding insects in the Cicadellidae family and Fulgoroidea superfamily such as leafhoppers and planthoppers, respectively. Based on 16S rDNA analysis, the phytoplasma detected in lisianthus and those detected in other plants in Israel including apricot (Prunus sp.), cyclamen (Cyclamen sp.), grape (Vitis vinifera), papaya (Carica papaya), and pepper (Capsicum annuum) have been placed in the Stolbur phytoplasma group (Weintraub et al. 2007). Six confirmed leafhopper vectors and one suspected planthopper vector of phytoplasmas occur in Israel (Weintraub et al. 2007).

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3.4.4 Management Management of phytoplasma diseases, like those caused by viruses, is based on monitoring and reduction of insect vector populations through chemical or biological means, vector exclusion by physical barriers such as fine mesh screening, vector disorientation and repellence in field-grown crops through the use of reflective mulches, and elimination of, and avoidance of growing near, alternate hosts especially established, infected crops.

4

Viral Diseases

4.1

Bromoviridae

4.1.1 Geographic Occurrence and Impact Viruses in the Bromoviridae commonly cause major worldwide crop losses. Two members of this group have been reported in lisianthus: Cucumber mosaic virus (CMV, genus Cucumovirus) and Tobacco streak virus (TSV, genus Ilarvirus) (de Freitas et al. 1996a; Fujinaga et al. 2006; Gera and Cohen 2007; Hahm et al. 1998; Providenti 1985; Rivas et al. 2000). The geographic occurrence of these viruses is indicated in Table 1. CMV was detected at an incidence of 8.6% in Japan (Nagano Prefecture) (Fuginaga et al. 2006) and 10% in alpine production areas in the ROK in combination with a Fabavirus, Broad bean wilt virus (Hahm et al. 1998). 4.1.2 Symptoms/Signs Symptoms caused by plant viruses, including the Bromoviridae, can be variable and depend on the host, environment, virus strain, and coinfection with other viruses; different Bromoviridae genera may produce similar symptoms. Symptoms caused by the Bromoviridae in lisianthus are indicated in Table 1. 4.1.3 Biology and Epidemiology Viruses in the Bromoviridae are acquired and spread by aphids (Hemiptera: Aphididae) or thrips (Thysanoptera: Thripidae) in a noncirculative (nonpersistent) manner; these viruses are mouthpart-borne and do not enter the insect’s hemolymph. Spread of TSV-infected pollen by thrips has been demonstrated (Klose et al. 1996). Seed transmission of CMV has been reported in many plant species at incidences of less than 1% up to 50% (Garcia-Arenal and Palukaitis 2008). Seed transmission of TSV in various hosts has been observed at incidences of 0.7–90.6% (Kaiser et al. 1982). 4.1.4 Management Virus management, including species in the Bromoviridae, is based on monitoring and reduction of vector populations through chemical and/or biological measures, vector exclusion by physical barriers such as fine mesh screens, vector disorientation in field-grown crops through the use of reflective mulches, and elimination of, and

Tobacco streak virus (TSV)

Viruses Cucumber mosaic virus (CMV)

Occurrence in lisianthus Brazil, Israel, Japan, the ROK, the USA Brazil

Frankliniella occidentalis, F. schultzei Microcephalothrips abdominalis, Thrips parvispinus, T. tabaci, and by seed

Transmission >80 aphid species including Aphis gossypii and Myzus persicae and by seed

Table 1 Bromoviridae infecting lisianthus

A wide range of vegetables, ornamentals, and weed species

Other natural hosts A very large number of vegetable, ornamental, and weed species Irregularly shaped, necrotic streaks or ring spots on leaves; flower size reduction and premature senescence

Symptoms in lisianthus Leaf mosaic and distortion, stunting, flower color-breaking, and malformation

References Garcia-Arenal and Palukaitis (2008), Gera and Cohen (2007), Hahm et al. (1998), Providenti (1985), and Rivas et al. (2000) de Freitas et al. (1996a), Kaiser et al. (1982), and Klose et al. (1996)

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avoidance of growing near, alternate hosts especially established, infected crops. Cross protection, prior infection with an attenuated strain of CMV, was shown to reduce subsequent infection by a severe strain of the virus in cucumber, lisianthus, petunia, and tomato (Sayama 1996). Resistance to CMV has been introduced into a wide variety of vegetables through genetic modification. Additional information on integrated disease management of viruses may be found in the introductory chapter 3 “Insect Management for Disease Control in Florists’ Crops.”

4.2

Bunyaviridae

4.2.1 Geographic Occurrence and Impact Plant viruses in the Bunyaviridae and genus Tospovirus cause extensive losses in nearly all cultivated plants. Thus far seven viruses in this genus have been detected in lisianthus including: Chrysanthemum stem necrosis virus (CSNV), Groundnut ringspot virus (GRSV), Impatiens necrotic spot virus (INSV), Iris yellow spot virus (IYSV), Lisianthus necrotic ringspot virus (LNRV), Tomato chlorotic spot virus (TCSV), and Tomato spotted wilt virus (TSWV) (Alexandre et al. 1999; de Freitas et al. 1996b; Fujinaga et al. 2006; Kritzman et al. 2000; McGovern unpublished data; McGovern et al. 1997b; Momonoi et al. 2011; Mumford et al. 2008; Shimomoto et al. 2014; Wolcan et al. 1996). Alexandre et al. (1999) found that three lisianthus plants in Brazil were co-infected with either CSNV + TCSV + GRSV or those three viruses + TSWV. The geographic occurrence of these viruses is indicated in Table 2. CSNV was observed at an incidence of 10% in a commercial greenhouse in Brazil (Duarte et al. 2014). A very low incidence (900 species in >90 monocotyledonous and dicotyledonous plant families

Not determined

Necrotic spots and ring spots in leaves (Fig. 18)

Ring spots, necrosis, leaf deformation, stunting

Necrotic spots and ring spots in leaves (Fig. 17)

Alexandre et al. (1999), de Freitas et al. (1996b), McGovern (unpublished data), Melgares de Aquilar Cormenzana (1996), Pappu et al. (2009), Veerakone et al. (2015), and Wolcan et al. (1996)

Alexandre et al. (1999)and Polston et al. (2013)

Shimomoto et al. (2014) and Zen et al. (2008)

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Fig. 14 Symptoms of CSNV: stem and foliar necrosis (Photo courtesy of Momonoi et al. (2011))

Fig. 15 Symptoms of INSV: distortion and yellow spots in newly developing leaves (left) and yellow-tan and ring spots in mature leaf (right) (R. J. McGovern)

Fig. 16 Symptoms of IYSV: stem necrosis (left) and systemic necrosis (right) (Photos courtesy of Srinivasan et al. (2011))

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Fig. 17 Symptoms of LNRV (Photo courtesy of Shimomoto et al. (2014))

Fig. 18 Symptoms of TSWV: yellow-tan ring spots (Photo courtesy of Florida Department of Agriculture and Consumer Services)

proximity of infected reservoir plants are the determining factors in Tospovirus outbreaks. Zen et al. (2008) detected IYSV in an onion field near lisianthus greenhouses in Japan and hypothesized that viruliferous T. tabaci dispersed after onion harvests spread the virus to lisianthus. Similarly, Kritzman et al. (2000) suggested that the source of IYSV outbreaks in lisianthus in Israel was infected onion crops.

4.2.4 Management • Cultural practices – These practices include the use of virus-free transplants and trap plants/crops, nonhost barrier crops, and exclusion/repellence of thrips by fine mesh screening or reflective mulches and covers. In field trials conducted in the USA (California), reflective mulches and plant covers, alone and in combination, reduced the incidence of F. occidentalis in chrysanthemum (Chrysanthemum morifolium), golden rod (Solidago sp.), and roses (Rosa x hybrid) (Newman and Robb 2010); however, reflective mulch effectiveness decreased with crop growth. • Sanitation – Removal of weeds and other host plants of tospoviruses and their vectors from greenhouses and the surrounding area is essential. Weed removal is often impractical for controlling tospoviruses with large host ranges such as TSWV and INSV in field-grown crops. However, growing new crops near existing crops infected by these viruses should always be avoided. Rouging infected plants should be rapidly carried out in a manner that will prevent further virus spread by thrips such as by bagging or insecticide application before removal.

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• Scouting – Monitoring thrips populations should be routinely done in the greenhouse and field through the use of yellow or blue sticky cards and/or tapping foliage over white paper. Virus indicator plants such as petunia (Petunia x hybrida) can be used in greenhouses to assess tospovirus inoculum pressure (Allen and Matteoni 1991). • Insecticides – Resistance management of the limited number of insecticides effective against the two major tospovirus vectors, F. occidentalis and T. tabaci, is a significant challenge (Martin et al. 2003; Reitz and Funderburk 2012; Robb et al. 1995). Therefore, insecticides cannot be viewed as a stand-alone treatment for either thrips or virus, even if resistance management regimes such as rotation and limited usage are rigorously maintained. On the other hand, the incidence of necrotic ring spot disease in greenhouse-grown lisianthus in Japan caused by IYSV was decreased when applications of the insecticide acephate were timed with the emergence of viruliferous T. tabaci (Zen et al. 2010). • Biological control – Biological control should be used preventively and not when thrips populations are high. Commonly used biocontrols for thrips management in greenhouses include predatory mites such as Neoseiulus cucumeris (formerly known as Amblyseius cucumeris) (Mesostigmata: Phytoseiidae), predatory bugs, Orius spp. (Hemiptera: Anthocoridae), and entomopathogenic nematodes such as Steinernema feltiae (Rhabditida: Steinernematidae) and Thripinema nicklewoodi (Tylenchida: Allantonematidae). In a greenhouse experiment in the UK, N. cucumeris was effective in controlling F. occidentalis on impatiens and decreased the spread and severity of TSWV (Bennison et al. 2001). Weekly foliar applications of S. feltiae reaching the growing medium reduced F. occidentalis populations, delayed TSWV infection, and reduced viral symptoms in greenhouse-grown chrysanthemums in the UK (Bennison et al. 2007). Infection of F. occidentalis by T. nicklewoodi reduced feeding, reproduction, and TSWV transmission by the insect in chrysanthemum in the USA (Texas) (Arthurs and Heinz 2003). • Resistance – Partial resistance to TSWV was identified in chrysanthemum cvs. (van de Wetering et al. 1999); similar tospovirus screening in lisianthus could provide a useful virus management strategy and starting point for a traditional resistance breeding program in this crop. Using Agrobacterium-mediated transformation, Sherman et al. (1998) produced lines of the chrysanthemum cv. Polaris resistant to TSWV. Peng et al. (2014) produced transgenic tobacco plants (Nicotiana benthamiana Domin.) that were resistant to multiple tospoviruses including GNRV, INSV, IYSV, and TSWV. A method for transformation of lisianthus was developed and the TSWV nucleoprotein was introduced, suggesting that genetic engineering of this crop for tospovirus resistance is feasible (Semeria et al. 1996). • Integrative strategies – Zen et al. (2010) reported that the combined use of acephate with reflective screening was very effective in reducing IYSV in lisianthus in greenhouses in Japan. The combination of UV-reflective mulch, the systemic acquired resistance activator acibenzolar-S-methyl, and insecticides (methamidophos, spinosad) was very effective in reducing TSWV incidence in field-grown tomato in the USA (Florida) (Momol et al. 2004).

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Additional information on integrated disease management of viruses may be found in the introductory chapter 3 “Insect Management for Disease Control in Florists’ Crops.”

4.3

Geminiviridae Tomato Yellow Leaf Curl Virus (TYLCV, Genus Begomovirus)

4.3.1 Geographic Occurrence and Impact The virus has become a limiting factor for lisianthus production in Israel; TYLCV incidences of near 100% have occurred in commercial facilities (Cohen et al. 1995). An incidence of 5% was reported in a greenhouse in the ROK (Kil et al. 2014). Two other B. tabaci-vectored begomoviruses, Ageratum yellow vein virus and Papaya leaf curl guangdong virus were reported to infect lisianthus in Taiwan (Cheng et al. 2005; Chen et al. 2016). 4.3.2 Symptoms/Signs TYLCV symptoms in lisianthus include stunting, upward curling, veinal swelling (on the lower leaf surface), and yellowing of leaves, flower bleaching, and failure to flower (Fig. 19). 4.3.3 Biology and Epidemiology TYLCV comprises a virus complex that has become a worldwide limiting factor for production of tomato (Solanum lycopersicum). The virus is transmitted circulatively Fig. 19 Symptoms of TYLCV: stunting, yellowing and upward curl of leaves, and flower bleaching in infected plant (left); healthy plant (right) (R. J. McGovern)

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and non-propagatively by the whitefly Bemisia tabaci (Gennadius) (Hemiptera: Aleyrodidae). Once TYLCV is acquired, the whitefly remains viruliferous for life. Besides lisianthus and tomato, other natural hosts of TYLCV include a number of solanaceous plants [black nightshade (Solanum nigrum), chili pepper (Capsicum chinense), Jimsonweed (Datura stramonium), petunia (Petunia x hybrida), sweet pepper (C. annuum), and tobacco (Nicotiana tabacum)] and common bean (Phaseolus vulgaris) (Díaz-Pendón et al. 2010).

4.3.4 Management Whitefly management through physical, chemical, and biological means and avoidance of locating lisianthus transplant and production facilities near established TYLCV-infected crops are essential for management of the virus. Additional information on integrated disease management of viruses may be found in the introductory chapter 3 “Insect Management for Disease Control in Florists’ Crops.”

4.4

Potyviridae

4.4.1 Geographic Occurrence and Impact Viruses in the genus Potyvirus infect most cultivated plants and weeds. A number of potyviruses cause disease in lisianthus including: Bean yellow mosaic virus (BYMV), Pepper veinal mottle virus (PVMV), Turnip mosaic virus (TuMV), and Watermelon mosaic virus (WMV). The geographic occurrence of these viruses is indicated in Table 3.

4.4.2 Symptoms/Signs The range of symptoms caused by potyviruses in lisianthus is presented in Table 3.

4.4.3 Biology and Epidemiology Potyviruses are spread by aphids (Hemiptera: Aphididae) in a noncirculative (nonpersistent) manner. It was suggested that other ornamentals, such as freesia (Freesia alba) or gladiolus, known to be hosts of BYMV, may have provided a reservoir for outbreaks of the virus in lisianthus (Lisa and Dellavalle 1987).

4.4.4 Management Uga et al. (2004) demonstrated that dwarf lisianthus plants that had been infected with an attenuated strain of BYMV (B-33) were protected from subsequent infection by a virulent isolate of the virus. Refer to the Bromoviridae Management section of this chapter and introductory chapter 3 “Insect Management for Disease Control in Florists’ Crops.”

Watermelon mosaic virus (WMV)

Pepper veinal mottle virus (PVMV) Turnip mosaic virus (TuMV)

Viruses Bean yellow mosaic virus (BYMV)

Japan

Brazil, Taiwan

Taiwan

Occurrence in lisianthus Brazil, Israel, Italy

>35 aphid species

Aphis gossypii, A. spiraecola, Myzus persicae, Toxoptera citricida 40–50 aphid species especially M. persicae and Brevicoryne brassicae

Transmission At least 19 aphid species with M. persicae being the most effective

Table 3 Potyviruses infecting lisianthus Other natural hosts 14 plant families including vegetables (legumes) and ornamentals (freesia, gladiolus, etc.) Mainly infects cultivated and weedy solanaceous plants (black nightshade, pepper, petunia, tomato, etc.) Chinese mustard (Brassica campestris ssp. chinensis), Chinese white cabbage (B. campestris ssp. chinese var communis), radish (Raphanus sativus), rape (B. campestrisi) and mustard (B. juncea), etc. >170 species in 26 mono- and dicotyledonous families including cultivated plants and weeds Chlorotic or whitish ring spots on leaves

Stunting, systemic yellow spotting

Symptoms in lisianthus Stunting, mosaic and leaf curl, flower colorbreaking Stunting, yellow blotches, and ring spots on leaves

Inoue and Kasuyama (2001)and Lecoq and Desbiez (2008)

Alexandre et al. (2005), Chao et al. (2000), and ICTVdB Management (2006b)

References Alexandre et al. (2005), Bos (2010), Gera and Cohen (2007), Lisa and Dellavalle (1987), and Swenson (1957) Cheng et al. (2009)and ICTVdB Management (2006a)

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4.5

R.J. McGovern

Secoviridae Broad Bean Wilt Virus (BBWV, Genus Fabavirus)

4.5.1 Geographic Occurrence and Impact BBWV was detected in lisianthus in Japan (Nagano Prefecture) at an incidence of 5.7% (Fujinaga et al. 2006) and at 10% in combination with CMV in alpine production areas of the ROK (Hahm et al. 1998). 4.5.2 Symptoms/Signs Mosaic 4.5.3 Biology and Epidemiology BBWV can be found worldwide and infects many economically important vegetable and ornamental crops. It is transmitted in a noncirculative manner by a number of aphids including A. gossypii and M. persicae (Belliure et al. 2009). 4.5.4 Management Refer to the Bromoviridae Management section of this chapter and introductory chapter 3 “Insect Management for Disease Control in Florists’ Crops.”

4.6

Tombusviridae

4.6.1 Geographic Occurrence and Impact Three members of the Tombusviridae infect lisianthus including Carnation mottle virus (CarMV, genus Carmovirus), Lisianthus necrosis virus [LNV (a strain of Eggplant mottled crinkle virus), genus Necrovirus], and Moroccan pepper virus (MPV, genus Tombusvirus). The geographic occurrence of these viruses is indicated in Table 4. CarMV was detected at an incidence of 20% in field-grown lisianthus in Taiwan (Chen et al. 2011). About 85% of the lisianthus surveyed in greenhouses in the Pakdasht region of Iran exhibited symptoms of MPV infection (Beikzadeh et al. 2011). 4.6.2 Symptoms/Signs The range of symptoms caused by the Tombusviridae in lisianthus is presented in Table 4. 4.6.3 Biology and Epidemiology The Tombusviridae contains virus genera that persist in, and are spread by, soil, plant debris, water, beetles, chytrid fungi such as Olpidium brassicae, and mechanically. It has been suggested that plants may become infected when their roots come in contact with sloughed-off virus-infected root tissue. Tombusviridae generally have a limited host range compared to other virus groups infecting lisianthus. The mode of infection of lisianthus by mechanically transmissible viruses with no known vectors in the outbreaks of CarMV and MPV mentioned above remains unexplained. It is

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Table 4 Tombusviridae infecting lisianthus Viruses Carnation mottle virus (CarMV)

Occurrence in lisianthus Taiwan

Transmission Mechanical, especially through asexual plant propagation

Lisianthus necrosis virus (LNV)

Japan, Taiwan

Mechanical and by the chytrid fungus Olpidium sp.

Moroccan pepper virus (MPV)

Iran, Japan

Mechanical

Other natural hosts 30 species in 15 plant families, including ornamentals such as calla lily (Zantedeschia spp.), carnation, geranium (Pelargonium hortulanum), hibiscus (Hibiscus sp.), narcissus (Narcissus sp.) Calla lily, carnation, and a number artificially inoculated ornamentals such as petunia and zinnia (Zinnia elegans)

Jimsonweed, lettuce (Lactuca sativa), pelargonium (Pelargonium zonale), pepper, tomato

Symptoms in lisianthus Systemic necrotic spots (Fig. 20)

Initially yellow spots appear on upper leaves followed by systemic necrotic leaf lesions, ring spots and distortion, tip necrosis, and flower colorbreaking (Fig. 21) Stunting, necrotic leaf spots, distortion, and necrosis of leaves and stems (Fig. 22)

References Chen et al. (2011)and Qu and Morris (2008)

Chang et al. (2007), Chen and Hsu (2002), Chen et al. (2000), and Iwaki et al. (1987)

Beikzadeh et al. (2011), Fischer and Lockhart (1977), Okhi et al. (2014), Vetten and Koenig (1983), and Wintermantel and Hladky (2013)

possible that these viruses were spread by cutting tools contaminated by use in another infected cut-flower crop or through the action of an unknown vector.

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Fig. 20 Symptoms of CarMV: systemic necrotic spots (Photo courtesy of Y.-K Chen)

Fig. 21 Symptoms of LNV: systemic yellow turning necrotic leaf lesions and ring spots, leaf distortion, and flower color-breaking (Photo courtesy of Y.-K Chen)

4.6.4 Management Refer to the introductory chapter 3 “Insect Management for Disease Control in Florists’ Crops.”

4.7

Virgaviridae

4.7.1 Geographic Occurrence and Impact Three members of this virus group infect lisianthus: Tobacco mosaic virus (TMV, genus Tobamovirus), Tomato mosaic virus (ToMV, genus Tobamovirus), and Pepper ringspot virus (PepRSV, genus Tobravirus). The geographic occurrence of these viruses is indicated in Table 5.

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Fig. 22 Symptoms of MPV: necrotic leaf spots (left); distortion and necrosis of leaves and stems (right) (Photos courtesy of Ohki et al. (2014))

Table 5 Virgaviridae infecting lisianthus Viruses Tobacco mosaic virus (TMV) Tomato mosaic virus (ToMV) Pepper ringspot virus (PepRSV)

Occurrence in lisianthus Israel

Transmission Mechanical, seed

Other natural hosts Very broad host range

Symptoms in lisianthus Stunting, leaf mosaic

Italy, Taiwan

Mechanical, seed

Very broad host range

Stunting, mosaic, and necrosis of leaves

Brazil

Mechanical

Some vegetable species and ornamentals such as Gerbera jamesonii, Gloxinia sylvatica

Stunting, mosaic, and necrotic line patterns in leaves, flower color-breaking

4.7.2 Symptoms/Signs The symptoms of these viruses are presented in Table 5.

References Gera and Cohen (2007) Jan et al. (2003) and Lisa and Gera (1995) Rivas et al. (2000)

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4.7.3 Biology and Epidemiology The Virgaviridae is a relatively new family of six rod-shaped plant virus genera including Tobamovirus and Tobravirus (Adams and Antoniw 2009). These viruses are transmitted mechanically and in some cases by seed, are resistant to degradation, and may persist in the soil. 4.7.4 Management Use virus-free seed. Promptly rogue infected plants and have workers disinfest their hands before resuming regular duties. Disinfest cutting tools between plants. Areas where a high virus incidence has been observed should be scheduled last for cultural activities.

5

Nematode Diseases

5.1

Root Knot (Meloidogyne spp.)

5.1.1 Geographic Occurrence and Impact Root knot in lisianthus has been reported in Italy, Israel, and the USA (California and Florida) (Elad et al. 2014; Russo and di Vito 2005; Schochow et al. 2004; Wang and McSorley 2004). 5.1.2 Symptoms/Signs Yellowing of foliage, stunting, flower delay and number reduction, and root galls (Fig. 23). 5.1.3 Biology and Epidemiology Root knot in lisianthus can be caused by three species of Meloidogyne (Tylenchida: Heteroderidae): M. hapla, M. incognita, and M. javanica. Root-knot nematodes begin as eggs in root tissue or the soil and pass through four juvenile stages (J1–J4) before reaching maturity. The J2 stage is the only one capable of infecting roots, where they produce giant cells in the host in which they feed, enlarge, and Fig. 23 Symptoms of rootknot nematode (R. J. McGovern)

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undergo their final developmental stages. The mature females become globose and produce eggs, while the adult males migrate from the host and become free-living. Root-knot nematodes are primarily spread by water, equipment contaminated with infested soil, and in infected plant material in the case of vegetatively propagated crops. Based on reproduction on cotton and/or tobacco, M. incognita was divided into four races (Taylor and Sasser 1978). Wang and McSorley (2004) reported that lisianthus “Avila Rose Rim” and “Echo Pink” were relatively poor hosts of M. incognita races 1 and 2. Schochow et al. (2004) found that lisianthus was a better host for M. javanica than for M. incognita and a poor host for M. hapla; but that all three species reduced flower number per plant.

5.1.4 Management Prevention of nematode dissemination through disinfestation of equipment is critical. Disinfestation of soil in florists’ crops production sites has utilized fumigation, chemical drenches, and heat treatment by steam or soil solarization. Refer to chapter 7 “Soil/Media Disinfestation for Management of Florists’ Crops Diseases.” 5.1.5 Additional Nematode Pathogens The following nematode pathogen of lisianthus has also been reported: Needle nematode (Longidorus spp.) (Israel, Elad et al. 2014)

References Adams MJ, Antoniw JF (2009) Virgaviridae: a new family of rod-shaped plant viruses. Arch Virol 154:1967–1972. doi:10.1007/s00705-009-0506-6 Aegerter BJ, Greathead AS, Pierce LE, Davis RM (2002) Mefenoxam-resistant isolates of Pythium irregulare in an ornamental greenhouse in California. Plant Dis 86(6):692. doi:10.1094/ PDIS.2002.86.6.692B. Accessed 22 Oct 2016 Alexandre MAV, Duarte LML, Rivas EB, Chagas CM (1999) Mixed infections by Tospovirus species in ornamental crops in São Paulo State, Brazil. Summa Phytopathol 25:353–356 Alexandre MAV, Vagueiro Seabra P, Borges Rivas E, Lembo Duarte LM, Galleti SR (2005) Vírus, viróides, fitoplasmas e espiroplasmas detectados em plantas ornamentais no período de 1992 a 2003. Rev Bras Hortic Ornam Campinas 11(1):49–57 (in Portugese) Alfieri SA, Langdon KR, Kimbrough JW, El-Gholl NE, Whelburg C (1994) Diseases and disorders of plants in Florida. Florida Department of Agriculture and Consumer Services, Division of Plant Industry Bulletin No. 14. Gainesville, FL Allen WR, Matteoni JA (1991) Petunia as an indicator plant for use by growers to monitor for thrips carrying the tomato spotted wilt virus in greenhouses. Plant Dis 75:78–82 Aloj B, Scalcione M, Nanni B, Marziano F (1990) Studies of a new phytopathogen in Italy: Peronospora of Eustoma (Lisianthus) russelianum. Annali della Facoltà di Scienze Agrarie della Università degli Studi di Napoli, Portici. 24:45–52 (in Italian, English abstract) Arthurs S, Heinz KM (2003) Thrips parasitic nematode Thripinema nicklewoodi (Tylenchida: Allantonematidae) reduces feeding, reproductive fitness, and Tospovirus transmission by its host, Frankliniella occidentalis (Thysanoptera: Thripidae). Environ Entomol 32(4):853–858 Auscher R (1997) Implementation of integrated pest management in Israel. Phytoparasitica 25 (2):119–141

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Bag S, Schwartz HF, Cramer CS, Havey MJ, Pappu HR (2014) Iris yellow spot virus (Tospovirus: Bunyaviridae): from obscurity to research priority. Mol Plant Pathol. doi:10.1111/mpp.12177 Ballard RW, Palleroni NJ, Doudoroff M, Stanier RY (1970) Taxonomy of the aerobic pseudomonads: Pseudomonas cepacia, P. marginata, P. alliicola and P. caryophylli. J Gen Microbiol 60:199–21 Beikzadeh N, Peters D, Hassani-Mehraban A (2011) First report of Moroccan pepper virus on Lisianthus in Iran and worldwide. Plant Dis 95(11):1485. doi:10.1094/PDIS-04-11-0342. Accessed 22 Oct 2016 Belliure B, Gómez-Zambrano M, Ferriol I, La Spina M, Alcácer L, Debreczeni DE, Rubio L (2009) Comparative transmission efficiency of two Broad bean wilt virus 1 isolates by Myzus persicae and Aphis gossypii. J Plant Pathol 91(2):475–478 Bennison J, Maulden K, Barker I, Morris J, Boonham N, Smith P, Spence N (2001) Reducing spread of TSWV on ornamentals by biological control of western flower thrips. In: Marullo R, Mound L (eds) Thrips and tospoviruses proceedings of the 7th International Symposium on Thysanoptera pp 215–219 http://www.ento.csiro.au/thysanoptera/Symposium/Section7/33Bennison-et-al.pdf. Accessed 22 Oct 2016 Bennison J, Maulden K, Tomiczek M, Morris J, Barker I, Boonham N, Spence N (2007) Use of entomopathogenic nematodes for the management of western flower thrips and tospoviruses. J Insect Sci 7(28):4. doi:10.1673/031.007.2807 Beringer PM, Appleman MD (2000) Unusual respiratory bacterial flora in cystic fibrosis: microbiologic and clinical features. Curr Opin Pulm Med 6:545–550 Bertoldo C, Gilardi G, Spadaro D, Gullino ML, Garibaldi A (2015) Genetic diversity and virulence of Italian strains of Fusarium oxysporum isolated from Eustoma grandiflorum. Eur J Plant Pathol 141(1):83–97 Booth C, Waterston, JM (1964) Fusarium avenaceum. CMI descriptions of pathogenic fungi and bacteria no. 25. CAB International Bos L (2010) Legume viruses. In: Mahy BWJ, van Regenmortel MHV (eds) Desk encyclopedia of plant and fungal virology. Elsevier/Academic, Amsterdam, pp 418–425 Bradbury JF (1973) Pseudomonas caryophylli. CMI descriptions of pathogenic fungi and bacteria no. 373. CAB International Braun U (1987) A monograph of the Erysiphales (powdery mildews). Beih Nova Hedwigia 89:1–700 Braun U (1995) The powdery mildews (Erysiphales) of Europe. Nord J Bot 16(2):121–232 Buddenhagen IW (1986) Bacterial wilt revisited. In: Persley GJ (ed) Bacterial wilt disease in Asia and the South Pacific, PRO13. Proceedings of an International Workshop, PCARRD, Los Banos, Philippines. Australia Centre for International Agricultural Research, Canberra, Australia pp 126–143 Buddenhagen IW, Kleman A (1964) Biological and physiological aspects of bacterial wilt caused by Pseudomonas solanacearum. Annu Rev Phytopathol 2:203–230 Buddenhagen IW, Sequeira L, Kelman A (1962) Designation of races in Pseudomonas solanacearum. Phytopathology 52:726 Cabrera MG, Vobis G, Alvarez R (2009) First record of powdery mildew caused by Oidium sp. (Erysiphaceae) on Eustoma grandiflorum in Argentina. Australas Plant Dis Notes 4:37–38 Cedeño L, Briceño A, Fermín G, Domínguez I, Pino H, Quintero K (2007) First record of Colletotrichum acutatum on lisianthus (Eustoma grandiflorum). Fitopatol Venez 20:41–43 Cedeño L, Rodríguez LA, Quintero K (2009) Primer reporte en Venezuela de mildiú polvoriento causado por Leveillula (Oidiopsis) taurica en lisiantus Fitopatol. Venez 22:23–24 (in Spanish) Cedeño L, Carrero C, Ruíz R, Fermín G, Pino H, Quintero K (2011) First report of Stemphylium solani in Lisiantus. Fitopatol Venez 24:38–41 Chang YS, Ko WH, Chen CC, Chen YK (2007) Characterization of a calla lily infecting isolate of Lisianthus necrosis virus. Plant Pathol Bull 16:149–156 Chao YC, Liang WJ, Huang LL, Ho WC, Tsai CH (1995) Bacterial wilt of Texas blue bell (Eustoma grandiflorum L.). Plant Pathol Bull (Taiwan) 4:193–195

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Reitz SR, Funderburk J (2012) Management strategies for western flower thrips and the role of insecticides. In: Perveen F (ed) Insecticides – pests engineering. InTech, Rijeka, pp 355–384, http://cdn.intechopen.com/pdfs-wm/28269.pdf. oV Riley DG, Joseph SV, Srinivasan R, Diffie S (2011) Thrips vectors of tospoviruses. J Integ Pest Manag 1(2):1–10. doi:10.1603/IPM10020 Rivas EB, Galleti SR, Duarte LML, Seabra PV, Alexandre MAV (2000) Virus and phytoplasm diseases of lisianthus. Summa Phytopathol 26(2):257–262 Robb KL, Newman J, Virzi JK, Parella MP (1995) Insecticide resistance in the western flower thrips. In: Parker BL, Skinner M, Lewis T (eds) Thrips biology and management. Plenum Press, New York, pp 341–346 Russo G, di Vito M (2005) Damages caused by the root-knot nematodes Meloidogyne hapla on lisianthus in Apulia. Inf Fitopatol 55(3):34–35 Sacoto Bravo HH (2009) Biological control of Pythium sp. and Fusarium oxysporum in growing lisianthus (Eustoma grandiflorum), four products using three types of substrates. Thesis (Ingeniero Agrónomo) Escuela Superior Politécnica de Chimborazo. http://dspace.espoch.edu. ec/bitstream/123456789/678/1/13T701%20SACOTO%20HORACIO.pdf (in Spanish). Accessed 22 Oct 2016 Sakata Ornamentals (2012) Lisianthus. http://www.sakataornamentals.com/_ccLib/attachments/ plants/Lisianthus+Cut+Flower. Accessed 22 Oct 2016 Sakata Seed America (2015a) Piccolo 2 Lisianthus. http://www.sakataornamentals.com/plantname/ Lisianthus-Piccolo-2. Accessed 22 Oct 2016 Sakata Seed America (2015b) Piccolo 3 Lisianthus. http://www.sakataornamentals.com/plantname/ Lisianthus-Piccolo-3. Accessed 22 Oct 2016 Salles LA, Sosa DA, Valerio A (2001) Alternatives for the replacement of methyl bromide in Argentina. In: Labrada R, Fornasari L (eds) Global report on validated alternatives to the use of methyl bromide soil fumigation, vol 166, Plant production and protection paper. FAO, Rome, pp 3–11, http://www.fao.org/docrep/004/Y1809E/y1809e02.htm#P155_2791. Accessed 22 Oct 2016 Sato T, Uematsu S, Mizoguchi H, Kiku T, Miura T (1997) Anthracnose of prairie gentian and loquat caused by Colletotrichum acutatum. Ann Phytopathol Soc Jpn 63:16–20 Sato T, Moriwaki J, Misawa T (2013) Molecular re-identification of strains of Colletotrichum acutatum species complex deposited in the NIAS Genebank and morphological characteristics of its member species. JARQ 47(3):295–305, http://www.jircas.affrc.go.jp. Accessed 22 Oct 2016 Satou M, Kobayashi M, Kanto T, Sugawara K, Yamada M, Ishiwata M (2013) Suppression of anthracnose of Russell Prairie Gentian by UV-B irradiation. Ann Rep Kanto-Tosan Plant Protect Soc 2013(60):79–81. doi:10.11337/ktpps.2013.79 (in Japanese). Accessed 22 Oct 2016 Sayama H (1996) Viral resistant tomato seedling production using attenuated cucumber mosaic virus (in Japanese). Plant Protect 50:20–25 Schochow M, Tjosvold SA, Ploeg AT (2004) Host status of lisianthus ‘Marachi Lime Green’ for three species of root-knot nematodes. HortSci 39(1):120–123 Schollenberger M, Zamorski C (2008) New bacterial disease of lisianthus. Prog Plant Prot 2:520–523 Seijo TE, McGovern RJ, Morrison RH (2000) Use of a Rotorod spore sampler to examine potential airborne dispersal of Fusarium avenaceum causing crown and stem rot of lisianthus. Phytopathology 90(6 Supplement):S128, http://www.apsnet.org/members/divisions/south/meetings/ Pages/2000MeetingAbstracts.aspx. Accessed 22 Oct 2016 Seijo TE, McGovern RJ, Dickstein ER, Harbaugh BK (2002) Bacterial crown rot of lisianthus caused by Burkholderia gladioli. Online. Plant Health Prog. doi:10.1094/PHP-2002-0520-01HN Semeria L, Ruffoni B, Rabaglio M, Genga A, Vaira AM, Accotto GP, Allavena A (1996) Genetic transformation of Eustoma grandiflorum by Agrobacterium tumefaciens. Plant Cell Tissue Org Cult 47:67–72

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Shao X-L, Gan Q-H, Li Y, Zhao W-J, Wu X-H, Su Z-P (2011) Detection of Burkholderia caryophylli by TaqMan real-time fluorescent PCR. Acta Phytopathol Sin 41(1):24–30 Sherman JM, Moyer JW, Daub ME (1998) Tomato spotted wilt virus resistance in chrysanthemum expressing the viral nucleocapsid gene. Plant Dis 82:407–414 Shew HD, Lucas GB (eds) (1991) Compendium of tobacco diseases. APS Press, St. Paul Shimomoto Y, Kobayashi K, Okuda M (2014) Identification and characterization of Lisianthus necrotic ringspot virus, a novel distinct tospovirus species causing necrotic disease of lisianthus (Eustoma grandiflorum). J Gen Plant Pathol 80:169–175. doi:10.1007/s10327-014-0503-9 Shinners L (1957) Synopsis of the genus Eustoma (Gentianaceae). Southwest Natl 2:38–43 Shpialter L, Rav David D, Dori I, Yermiahu U, Pivonia S, Levite R, Elad Y (2009) Cultural methods and environmental conditions affecting gray mold and its management in lisianthus. Phytopathology 99:557–570 Spadotti DMA, Leão EU, Rocha KCG, Pavan MA, Krause-Sakate R (2014) First report of Groundnut ringspot virus in cucumber fruits in Brazil. New Dis Rep 29:25. doi:10.5197/ j.2044-0588.2014.029.025. Accessed 22 Oct 2016 Srinivasan R, Sundaraj S, Pappu HR, Diffie S, Riley DR, Gitaitis RD (2012) Transmission of Iris yellow spot virus by Frankliniella fusca and Thrips tabaci (Thysanoptera: Thripidae). J Econ Entomol 105(1):40–47. doi:10.1603/EC11094. Accessed 22 Oct 2016 Stegmark R (1994) Downy mildew on peas (Peronospora viciae f. sp. pisi). Agronomie EDP Sci 14 (10):641–647 Stovold G (1998) Fungicide resistance in isolates of Botrytis from ornamentals and development of control strategies. Horticultural Research & Development Corporation, New South Wales, Australia, Report NY 306. http://www.ngia.com.au/Attachment?Action=Download&Attach ment_id=1237. Accessed 22 Oct 2016 Strandberg J (2003) Colletotrichum leaf and flower blight of lisianthus. University of Florida-IFAS, Mid-Florida Research and Education Center Sugawara T, Ishii T (2009) Bacterial wilt of Eustoma grandiflorum. National Agriculture and Food Research Organization, Japan, http://www.naro.affrc.go.jp/flower/kakibyo/plant_search/ta/ lisianthus/post_290.html (in Japanese). Accessed 22 Oct 2016 Swenson KG (1957) Transmission of Bean yellow mosaic virus by aphids. J Econ Entom 50 (60):727–731 Taubenhaus JJ, Ezekiel WN (1935) Fusarium crown and root rot, and sclerophoma stem blight, of the Texas Bluebell. Bull Torrey Bot Club 62(9):503–510 Taylor AL, Sasser JN (1978) Biology, identification, and control of root-knot nematodes (Meloidogyne species). North Carolina State University Graphics, Raleigh Tomita Y, Chiba T, Ogawara T, Nagatsuka H (2004) Occurrence of several diseases during Russell Prairie gentian cultivation in Ibaraki prefecture. Bull Hortic Instit Ibaraki Agric Center 12:28–38 (in Japanese) Toppe B (2005) Fungal diseases in Eustoma grandiflorum – occurrence and control. Grønn Kunnskap 9(2):70–75, http://www.bioforsk.no/ikbViewer/Content/18614/toppe2.pdf (In Norwegian). Accessed 22 Oct 2016 Truter M, Wehner FC (2004) Crown and root infection of lisianthus caused by Fusarium solani in South Africa. Plant Dis 88(5):573. doi:10.1094/PDIS.2004.88.5.573A Uematsu S, Okubo H, Suzui T, Shiota A, Chiba T (1996) First report of phytophthora rot of eustoma grandiflorum caused by phytophthora spp. Ann Phytopathol Soc Jpn 62(3):266, http://ci.nii.ac. jp/naid/110002733867/ (in Japanese). Accessed 22 Oct 2016 Uga H, Kobayashi YO, Hagiwara K, Honda Y, Omura T (2004) Selection of an attenuated isolate of Bean yellow mosaic virus for protection of dwarf gentian plants from viral infection in the field. J Gen Plant Pathol 70:54–56. doi:10.1007/s10327-003-0091-6 USDA-NASS (2012) Floriculture: U.S. Summary. 8 pp http://www.nass.usda.gov/Statistics_by_ State/Ohio/Publications/Reports_by_Title/flor2012.pdf . Accessed 22 Oct 2016 van de Wetering F, Posthuma K, Goldbach R, Peters D (1999) Assessing the susceptibility of chrysanthemum cultivars to tomato spotted wilt virus. Plant Pathol 48:693–699

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van der Burg AMM, De Kreij C (2003) Voorkomen uitval bij Lisianthus: Onderzoek op 3 praktijkbedrijven. Wageningen, Praktijkonderzoek Plant & Omgeving B.V. PPO rapport nr.420002. http://www.tuinbouw.nl/sites/default/files/documenten/00021689.pdf (in Dutch). Accessed 22 Oct 2016 van der Wurff A, Hamelink R (2007) Middelentoets uitval Lisianthus: Vergelijkende effectiviteitsproef van middelen tegen Fusarium avenaceum en Myrothecium roridum in Lisianthus (Eustoma sp.) cv. Picolo White Wageningen UR Glastuinbouw (in Dutch). http://edepot.wur.nl/ 272656. Accessed 22 Oct 2016 Veeracone et al (2015) A review of the plant virus, viroid, liberibacter and phytoplasma records for New Zealand. Aust Plant Pathol 44:463–514. doi:10.1007/s13313-015-0366-3 Vetten HJ, Koenig R (1983) Natural infection of tomato and pelargonium in Germany by a tombusvirus originally described from pepper in Morocco. Phytopathol Z 108:215–220 Vrind TA (2005) The Botrytis problem in figures. Acta Hortic 669:99–102, http://www.actahort. org/books/669/669_11.htm. Accessed 22 Oct 2016 Wang K-H, McSorley R (2004) Host status of several cut flower crops to the root-knot nematode, Meloidogyne incognita. Nematropica 35(1):37–44 Webster CG, Frantz G, Reitz SR, Funderburk JE, Mellinger HC, McAvoy E, Turechek WW, Marshall SH, Tantiwanich Y, McGrath MT, Daughtrey ML, Adkins S (2015) Emergence of groundnut ringspot virus and tomato chlorotic spot virus in vegetables in Florida and the Southeastern United States. Phytopathology 105(3):388–398. doi:10.1094/PHYTO-06-140172-R Wegulo SN, Vilchez M (2004) Evaluation of fungicides for control of Botrytis blight of lisianthus. Fungicide Nematicide Tests 61:OT030 Wegulo SN, Vilchez M (2007) Evaluation of lisianthus cultivars for resistance to Botrytis Cinerea. Plant Dis 91:997–1001 Weintraub PG, Zeidan M, Spiegel S, Gera A (2007) Diversity of the known phytoplasmas in Israel. Bull Insectol 60(2):143–144 Whitby PW, Pope LC, Carter KB, LiPuma JJ, Stull TL (2000) Species-specific PCR as a tool for the identification of Burkholderia gladioli. J Clin Microbiol 38:282–285 Wintermantel WM, Hladky LL (2013) Complete genome sequence and biological characterization of Moroccan pepper virus (MPV) and reclassification of Lettuce necrotic stunt virus as MPV. Phytopathology 103:501–508 Wolcan SM (2005) Occurrence of Pseudocercospora eustomatis on Eustoma grandiflorum in Argentina. Australas Plant Pathol 34(4):617–618 Wolcan S, Ronco L, Dal Bo E, Lori G, Alippi H (1996) First report of diseases on Lisianthus in Argentina. Plant Dis 80:223. doi:10.1094/PD-80-0223A Wolcan S, Lori G, Ronco L (2001a) First report of Fusarium solani causing stunt on lisianthus. Plant Dis 85(4):443, http://dx.doi.org/10.1094/PDIS.2001.85.4.443C. Accessed 22 Oct 2016 Wolcan SM, Lori GA, Ronco L, Mitidieri AF, Fernandez R (2001b) Enanismo y podredumbre basal de Eustoma grandiflorum y su relación con la densidad de Fusarium solani en el suelo. Fitopatol Bras 26:710–714, http://www.scielo.br/pdf/fb/v26n4/8177.pdf (in Spanish). Accessed 22 Oct 2016 Yang HC, Hsieh TF (1998) Salisb Plant Protect Bull (Taichung) 40(1):37–48 Yoshimatsu H (1993) Root and foliage rot of Eustoma grandiflorum caused by Rhizoctonia solani Kuhn. Ann Phytopathol Soc Jpn 59(3):284 (in Japanese) Young JM, Saddler GS, Takikawa Y, De Boer SH, Vauterin L, Gardan L, Gvozdyak RI, Stead DE (1996) Names of plant pathogenic bacteria 1864–1995. Rev Plant Pathol 75:721–763 Yunis H, Elad Y (1989) Survival of Botrytis cinerea in plant debris during summer in Israel. Phytoparasitica 17:13–21 Zen S, Okuda M, Fuji S, Iwanami T (2008) The seasonal occurrence of viruliferous Thrips tabaci and the incidence of Iris yellow spot virus disease on lisianthus. J Plant Pathol 90(3):511–515 Zen S, Nakashima S, Tashiro N, Okuda M, Fuji S (2010) Control of lisianthus necrotic ringpot disease caused by Iris yellow spot virus using reflective net and insecticide application adjusted

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to the immigrating periods of viruliferous onion thrips, Thrips tabaci Lindeman. Jpn J Phytopathol 76:17–20, https://www.jstage.jst.go.jp/article/jjphytopath/76/1/76_1_17/_pdf. Accessed 22 Oct 2016 Zulfiqar M, Brlansky RH, Timmer LW (1996) Infection of flower and vegetative tissues of citrus by Colletotrichum acutatum and C. gloeosporioides. Mycologia 88:121–128

Diseases of Orchid Prasartporn Smitamana and Robert J. McGovern

Abstract

Orchids are monocotyledonous plants that belong to the Orchidaceae family which has a diverse range of habitats from tropical to temperate zones. Orchids grow at different elevations from sea level to the very high mountainous level of the Himalayas. Due to great variation, orchids can be classified in many genera and can be infected with many of the same fungi, bacteria, viruses, and nematodes as other plants. In this chapter, economically important diseases of orchids and their management are described. Keywords

Botrytis • Phytophthora • Pythium • Fusarium • Rhizoctonia • Sclerotium • Acidovorax • Dickeya • Burholderia • Viruses • Foliar Nematodes

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-Like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Black Rot (Phytophthora and Pythium spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Shoot Rot or Top Rot [Phytophthora nicotianae (syn. Phytophthora parasitica); Phytophthora cactorum] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Botrytis Spot (Botrytis cinerea) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Petal Blight (Curvularia eragrostidis, Alternaria alternata) . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Sclerotium Stem Rot (Sclerotium rolfsii) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2 2 2 5 6 7 8

P. Smitamana (*) Agricultural and Industrial Clinic Co. Ltd., Chiang Mai, Thailand Department of Plant Pathology, Faculty of Agriculture, Chiang Mai University, Chiang Mai, Thailand e-mail: [email protected] R.J. McGovern NBD Research Co., Ltd., Lampang, Thailand, e-mail: [email protected] # Springer International Publishing AG 2016 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florist's Crops Diseases, Handbook of Plant Disease Management, DOI 10.1007/978-3-319-32374-9_21-1

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2.6 Black Leg/Dry Rot and Slow Decline (Fusarium spp. Rhizoctonia solani) . . . . . . . . . . 2.7 Leaf Spots (Fusarium, Phyllosticta, Guignardia, Septoria, Cercospora spp.) . . . . . . . 2.8 Anthracnose [Colletotrichum gloeosporioides (syn. Glomerella cingulata)] . . . . . . . . . 2.9 Rust (Sphenospora kevorkianii) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Bacterial and Phytoplasma Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Leaf Spot (Acidovorax avenae subsp. cattleyae) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Soft Rot [Dickeya chrysanthemi (syn. Pectobacterium chrysanthemi); D. dieffenbachiae] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Black Spot [Burkholderia gladioli (syn. Pseudomonas gladioli)] . . . . . . . . . . . . . . . . . . . . 3.4 Phyllody (“Candidatus Phytoplasma” sp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Viral Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Nematode Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 Foliar Nematodes (Aphelenchoides spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Introduction

Orchid belongs to the Orchidaceae which is a highly diverse family and can be found worldwide, but most species are found in the tropics, especially in the tropical mountains (Dressler 1993). Orchidaceae is possibly the largest family of the flowering plants and consists of 899 genera that include 71,391 species, but only 27,135 are accepted species names (The Plant List 2013). Among the orchids, 73% are epiphytes (Atwood 1986) and show stereotypical flower structure and remarkably diverse plant structure (Benzing 1986a, b). Orchids are economically important ornamental crops as potted plants and cut flowers but also provide raw materials for the fragrance and pharmaceutical industries. In some countries, orchids serve as the major source of income for growers. The commercial production of orchid began in natural habitats but has been adapted for large-scale industrial production. This change has made the environment conducive for the outbreak of many diseases. Due to vegetative propagation, the planting materials are the major source of inoculum, and pathogens are easily distributed, especially viruses. Major diseases of orchids and their control measures are listed below.

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Fungal and Fungus-Like Diseases

2.1

Black Rot (Phytophthora and Pythium spp.)

Geographic occurrence and impact. Black rot is a common disease found in all orchid genera, especially after heavy rainfall or when in highly humid environments for extended periods, such as occur in poorly ventilated, crowded collections. Black rot is caused by one or both pathogens belonging to Phytophthora and Pythium spp. Fusarium can frequently occur as a secondary pathogen. Pythium ultimum, Phytophthora cactorum, and Phytophthora palmivora are commonly reported as the major pathogens (Hine 1962; Burnett 1974; Uchida 1994; Uchida and Aragaki

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1991a; Orlikowski and Szkuta 2006; Cating et al. 2013). Phytophthora multivesiculata was found in Cymbidium in New Zealand (Ilieva et al. 1998; Hill 2004). Black rot is a common disease in all nurseries worldwide, especially in the tropics and subtropics, including Australia, Bulgaria, China, New Zealand, Poland, Taiwan, and the USA (Hawaii) (Burnett 1974; Chen and Hsieh 1978; Hall 1989; Duff 1993; Uchida 1994; Ilieva et al. 1998; Hill 2004; Orlikowski and Szkuta 2006; Tao et al. 2011). Symptoms/signs. In seedlings and young plantlets, water soaked lesions and discoloration occurs at the base of the plant and roots during periods of high humidity. This condition can lead to damping off. In monopodial type orchids, e.g., Vanda, Phalaenopsis, and Rhynchostylis, the disease usually starts on new leaves as brown to black lesions (Figs. 1a, b and 2b).

Fig. 1 Black rot lesions surrounded by a yellowish region on Vanda (a) and Rhyncostylis (b) orchids; infected root velamen has dried out (b) (P. Smitamana)

Fig. 2 Black rot of Catteleya pseudobulb (a) and Vanda leaves (b) on which the fungal mycelia are visible under high moisture conditions (a, P. Smitamana; b, courtesy of Kamjaipai 1983)

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The pathogens rapidly invade downward into the stem and roots. The infected shoot can easily be pulled out and, if unattended, the whole plant will die. In the sympodial type orchids, e.g., Cattleya, Dendrobium, and Oncidium, a light yellow-brown discoloration on the pseudobulbs is observed. After the disease progresses, softening and rotting of the pseudobulbs which turn brown to black occurs (Fig. 2a). The infected roots become darkly discolored as the velamen rots. Leaf symptoms first appear on the underside as small, water-soaked, irregular spots that are yellowish brown in color, then turn brown or black with a yellowish margin. Under high temperature and humidity, the lesions rapidly enlarge and become soft with ooze if pressed. If the humidity decreases, the lesions dry and turn black into which other pathogens can easily invade. Pathogens can also infect the leaves via wounds. In general, root symptoms initially appear as brown to black rot of root tips; the stem and/or pseudobulbs may rapidly rot. Infected roots can also be stunted, lose the velamen, and dry out. In Cymbidium, a dry rot of leaves can occur that has alternating horizontal light- and dark-brown zebra-like stripes. The pseudobulb tissue usually becomes wet and brown-black (Ilieva et al. 1998). Biology and epidemiology. Phytophthora and Pythium are fungus-like, but not true fungi, and are favored by high humidity. Both species spread rapidly if free water is available giving rise to zoospores that swim freely to, penetrate, and infect plant tissue (Agrios 2005). Following infection, mycelia spread rapidly through the infected tissues. The lesions rapidly expand and exhibit brown-black necrotic regions. Motile zoospores produced in the infected lesions are spread easily in irrigation water and are splash-dispersed on uninfected tissue during watering (Uchida 1994). Management. • Cultural practices – Carefully check newly acquired plants to be sure that they are free from disease. Separate new stock from other plants and monitor them for disease symptoms for least 6 weeks before moving them into the nursery. Elevating the plants 60–90 cm above the ground to avoid splashing from the soil below or keeping them on a solid surface can also help prevent infections. Growers should maintain good ventilation through adequate plant spacing to reduce duration of leaf wetness. • Sanitation – If early symptoms of black rot are found, immediately excise infected tissue with a clean knife and discard it in order to prevent the spread of the disease on that plant or to other plants. Application of a fungicide (fosetyl-al, mefenoxam, etc.) may be effective when used immediately after. Use only clean water for watering orchids. Deep well water is safer than surface water unless surface water is disinfested. Discard or burn severely infected plants to prevent the spread of the disease. • Fungicides and biocontrols – Use suitable fungicides (fosetyl-al, mefenoxam, promocarb hydrochloride, trifloxystrobin, etc.) as a preventative measure, particularly during hot humid periods. The highest rate of the harpin protein, which induces systemic acquired resistance, significantly reduced black rot in Vanda

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caused by P. palmivora compared to the control; Bacillus subtilis + Trichoderma harzianum and etridiazole + quintozene were ineffective (Maketon et al. 2015). Growers should follow all label instructions.

2.2

Shoot Rot or Top Rot [Phytophthora nicotianae (syn. Phytophthora parasitica); Phytophthora cactorum]

Geographic occurrence and impact. This is a common disease found mostly on Vanda (upper stem) and Cattleya (leaf shoot) and is favored by high humidity and temperature, and poor ventilation due to high plant densities. This disease is caused by Phytophthora nicotianae, which can cause the death of the whole plant (Vanda); or in the case of Cattleya caused by Phytophthora cactorum, which causes rot of the shoots, leaves, pseudobulbs, rhizomes, and flower buds. Like black rot, shoot rot is commonly found in humid and high temperature environments especially in the tropical and subtropical zones: Thailand, USA (Florida, Hawaii), Poland, and Australia (Daly et al. 2013; Kamjaipai 1983; McMillan et al. 2009; Orlikowski and Szkuta 2006; Uchida 1994; Uchida and Aragaki 1991a, b). Symptoms/signs. A symptom of top rot includes the death of new leaves, which turn dark brown. This discoloration advances down the stem similar to the symptom caused by black rot. This disease may also start at the base of the stem and spread upward resulting in the same dark brown stem discoloration. Shoot rot was detected on Cattleya orchids in Darwin, Australia, and in USA (Florida) causing the rapid death of new side shoots, turning them almost black. The pathogen will spread back along the rhizomes to the next shoot causing the same symptoms. This disease has been reported on only Cattleya and Vanda but may affect others. Biology and epidemiology. As stated before, Phytophthora is fungus-like and its infection and spread are favored by wet conditions. Phytophthora cactorum is a homothallic oomycete that produces a sexual spore, an oospore, for survival under unfavorable environments. In addition, this fungus also asexually produces survival spores called chlamydospores as well as sporangia. The pathogen spreads by means of the pear or lemon shaped motile zoospores produced by either oospores or sporangia under wet conditions. After encysting and germination, zoospores can penetrate the plant directly or enter through wounds to cause infection. Management. • Sanitation and fungicides – Immediately cut the infected top with a clean knife, and discard severely infected plants to prevent the spread of the pathogen, and spray the plants with a fungicide (fosetyl-al, mefenoxam, etc.). Use suitable fungicides (see Sect. 2.1 above) as a preventative measure, particularly during hot humid periods, following label instructions. Improve ventilation to reduce leaf wetness. • Biocontrol – Certain biocontrol products containing Streptomyces lydicus, Bacillus subtilis, Trichoderma species, and Gliocladium virens applied prior to the

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disease occurring in the growing area have been reported to help prevent some soilborne and aerial diseases (Bottom 2016). Spraying infected plants with a spore suspension of Trichoderma harzianum or T. viride, or in the case of Vanda directly injecting the solution into the infected shoot, may be helpful (Smitamana, unpublished data).

2.3

Botrytis Spot (Botrytis cinerea)

Geographic occurrence and impact. Botrytis spot is a fungal disease affecting many types of orchids, mostly during cool, damp weather and in nurseries with poor air circulation. Phalaenopsis and Cattleya are particularly susceptible to Botrytis cinerea infection, but the disease may be found in a wide range of orchid genera. This disease has been reported in the tropics and the temperate regions including the USA (Florida, Hawaii) and Thailand (Jones 2003; Kamjaipai 1983; Uchida and Aragaki 1991b). Symptoms/signs. Older flowers are highly susceptible to infection. Brown necrotic or pale spots on orchid flowers are typical symptoms. The spots may enlarge and increase in number as the infection progresses and may be surrounded by a yellowish or pale pink margin depending on the background color of the flowers. Under cool and high moisture conditions, fungal mycelial webbing may be visible. Biology and epidemiology. Botrytis cinerea is an airborne plant pathogen commonly found in nurseries. Hyaline conidia of the fungus are produced on grey, branching conidiophores. Sclerotia, highly resistant survival structures, can be produced on dying infected tissue. In the temperate zone, B. cinerea overwinters in dead and dying plant material and begins producing and dispersing spores during cool, damp weather in the spring and autumn. Spores are distributed by wind, rain, irrigation water, or any mechanical action. Temperatures in the range of 18–25  C/ 64–77  F and wet plant surfaces or at least 92% RH are conducive factors for infection, and the pathogen can quickly proliferate, infecting the surrounding healthy plant tissue in less than 14 h (Agrios 2005; Sumbali 2005). Management. • Cultural practices – Growers should avoid wetting the flowers as much as possible. In tropical nurseries, the growers always spray or water the whole nonflowering plants early in the morning to make sure that the plants will be dry in the late afternoon or by nightfall. Water that remains on petals or leaves after a rain or watering promotes fungal growth. Avoid damp environments and poor air circulation that promote pathogen growth. Facilitate good air circulation through proper plant spacing and decrease the humidity during cool, damp weather by heating and venting greenhouses in temperate climates, especially when plants are in the blooming stage. • Sanitation – Remove any plant debris, fallen flowers, leaves, and infected tissue from the growing area in closed containers to reduce the infection potential and the possibility of spreading the fungus. Burning or burying all affected plant

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Fig. 3 Petal blight of Dendrobium flowers: water-soaked lesions turning brown or rust colored on white flower petals (a); and whitish yellow lesions on “Madame Pompadour” flowers (b). (Photos courtesy of Kamjaipai 1983)

tissue is recommended. Eliminate unwanted residual ornamental crops and weeds in the growing area. Make sure that newly introduced plants are free of any disease; normally keep them in an isolated area for at least 2 weeks before bringing them into the nursery. • Fungicides – Spray preventively with fungicides such as chlorothalonil, thiophanate methyl, iprodione, or vinclozolin. Resistance to thiophanate methyl and vinclozolin has been widely reported in B. cinerea; therefore, rotation of these fungicides with those of different modes of action is essential.

2.4

Petal Blight (Curvularia eragrostidis, Alternaria alternata)

Geographic occurrence and impact. This disease is found mostly in the Dendrobium hybrids whose flowers are most susceptible to the pathogens. Petal blight has been reported in Australia, Thailand, and the USA (Burnett 1957, 1965; Daly et al. 2013; Kamjaipai 1983). Symptoms/signs. Small brown spots develop on the flower petals, then the spots enlarge and merge to form necrotic lesions; under high moisture conditions or after rainfall these spots will rot (Fig. 3a, b). Biology and epidemiology. Alternaria and Curvularia belong to the fungal family Dermatiaceae which produces both dark hyphae and conidia. No organized fruiting bodies are formed, and the majority of these groups are saprophytic but some are plant pathogens (Sumbali 2005). Conidia of Alternaria are typically club or pear shaped and multicellular with both transverse and longitudinal cross walls. Conidia are easily detached and carried by air currents (Agrios 2005). Curvularia conidia are curved, clavate, broadly fusiform, obvoid, or pyriform with three or more transverse septa, pale or dark brown, with the end cells paler than the others (Ellis 1977). Both Alternaria and Curvularia can survive in plant debris, infected, or dry tissue, and the conidia are easily spread by air currents or water drops splashed off the orchid plants during overhead irrigation. Management.

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• Cultural practices – Keep orchids in bloom under solid rooves in order to protect them from rain. Avoid damp environments caused by incorrect watering practices and poor air circulation that promote pathogen infection. • Fungicides and biocontrols – The highest rate of the harpin protein, which induces systemic acquired resistance, significantly reduced rust spot in Dendrobium flowers caused by Curvularia lunata, compared to the control. Formulations of Bacillus subtilis + Trichoderma harzianum and mancozeb were intermediate in effectiveness (Maketon et al. 2015).

2.5

Sclerotium Stem Rot (Sclerotium rolfsii)

Geographic occurrence and impact. This disease is found mostly in Ascocenda, Brassidium, Cymbidium, Dendrobium, Neofinetia, Phaius, Paphiopedilum, Phalaenopsis, and Vanda orchids. Stem rot is commonly found in the tropics especially in nurseries that do not have roofs to protect the foliage from rainfall. Pots with poor drainage, such as those used to grow Paphiopedilum, are conducive to the disease. Stem rot has been reported in both the tropics and temperate zones including Australia, India, Korea, Thailand, and the USA (Florida) (Bag 2004a, b; Cating et al. 2009a; Cating et al. 2013; Daly et al. 2013; Han et al. 2012; Kamjaipai 1983; Tara et al. 2003). Symptoms/signs. The symptoms of this disease usually start on the lower shoots, pseudobulbs, and leaves which turn yellow, grow poorly, and then rapidly collapse (Fig. 4a). Rotting of the stem (Fig. 4c), roots, and leaf tissue gradually occurs, and in severe infections the entire plant wilts and dies. White fluffy mats of fungal mycelia and small yellow cream to brown spherical, mustard seed-size bodies called sclerotia appear on infected tissue (Fig. 4d). On Dendrobium plants, the pathogen can infect the leaves and basal part of pseudobulbs on which white mycelia are found; or in certain cases, the plants can be infected from the top causing rapid collapse and rotting of shoot apices (Fig. 4b). On Phaius flavus and Paphiopedilum venustum the disease may be found on the base of the pseudobulbs and collar region, respectively, and causes rot and leaf yellowing or collapse, which spreads upward, until the entire plant turns brown to black and dies. Diagnostically characteristic white mycelia and small sclerotia are found on the infected areas. Biology and epidemiology. Sclerotium rolfsii is characterized by small cream to dark brown sclerotia produced on septate, white fluffy mycelia with hyphal clamp connections (Sumbali 2005). Sclerotia can be spread by water and act as sources of infection. The fungus has a wide host range and is favored by a warm, moist environment. In the temperate zone, the pathogen can overwinter in the sclerotia and starts infecting new plants when the temperatures become warm. Sclerotia can survive in soil for many years. Management. Managing Sclerotium stem rot is quite difficult because of the pathogen’s wide host range and long survival in soil via sclerotia. Infected plants should be immediately destroyed, and an appropriate fungicide such as PCNB

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Fig. 4 Infected Vanda basal shoot with white mycelia and small sclerotia (a), young Dendrobium plant with severe leaf rot and white mycelia (b), severely infected Vanda showing dry rot of stem (c), and sclerotia on the infected stem and leaves (d) (Photos courtesy of Kamjaipai 1983)

should be applied when the disease is observed. Using biocontrol agents, such as Trichoderma spp., as an alternative measure for preventing and controlling the pathogen is also recommended.

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2.6

Black Leg/Dry Rot and Slow Decline (Fusarium spp. Rhizoctonia solani)

Geographic occurrence and impact. Fusarium solani, F. oxysporum, F. subglutinans, and F. proliferatum have been associated with the disease; in addition, Rhizoctoniasolani has also been involved in some areas. Susceptible orchids include Dendrobium, Cattleya, Phalaenopsis, Vanda, Rhyncostylis, and Vanilla. Black leg, dry rot, and slow decline have been reported in the tropics and temperate zones including Australia, Brazil, Indonesia, Malaysia, Taiwan, Thailand, the USA (Florida) (Daly et al. 2013; Dehgahi et al. 2014; Kamjaipai 1983; Pinaria et al. 2010; Su et al. 2010; Pedroso-de-Moraes et al. 2011; Wedge and Elmer 2008). If uncontrolled, Fusarium rot can affect up to 100% of an orchid crop (Kawate and Sewake 2014). Fusarium is considered a quarantine pest. Symptoms/signs. This disease is mostly found in orchids grown in damp environments or waterlogged pots especially after heat stress and heavy fertilization (Wedge and Elmer 2008). Plantlets after transferring from tissue culture vessels to community trays or pots are most susceptible to infection. Infected plantlets show water soaked root tips followed by leaf yellowing in whole plantlets. In mature plants, the symptoms are mostly observed after transplanting when the pathogens can easily infect the plants through wounds. Infected plants show discoloration of the root tips and a gradual dry rot. Pseudobulbs become spongy and discolored, successive new shoots tend to get smaller in height, and stem thickness is also reduced. The leaves will yellow and drop off, one by one, until none are left and the plant dies (Fig. 5). Biology and epidemiology. Fusarium and Rhizoctonia are both soilborne pathogens. Fusarium at first produces colorless mycelia but with age the mycelia color will change to cream, pale yellow, pale pink, or purplish. Fusarium produce three kinds of asexual spores: microconidia which have 1–2 cells; curved macroconidia which have 3–5 cells, and 1–2 cell, thick-walled resistive chlamydospores (Agrios 2005). The microconidia and macroconidia are dispersed easily by air currents and spread with rain and irrigation water. Fusarium can also be spread by fungus gnats. Rhizoctonia produces sterile colorless mycelia which will change to yellowish or light brown with age. Hyphae consist of long cells, with branching at an approximate right angle to the main hypha with slight hyphal constrictions at the junction. Under certain conditions, Rhizoctonia produces sclerotia which function as survival structures (Agrios 2005). The fungus can overwinter as mycelia or sclerotia in or on infected plants or by contaminating planting materials and containers, e.g., coconut husks and pots. The fungus spreads by rain or irrigation water. Management. • Resistance – Brassocattleya “Orquidacea’s Melody” and Brassocattleya “Orquidacea’s Rare Bird,” were resistant and moderately resistant, respectively, to Fusarium wilt caused by Fusarium oxysporum f. sp. Cattleyae (Pedroso-deMoraes et al. 2011).

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Fig. 5 Primary symptoms of Vanda plantlet rot caused by Fusarium spp. include root tip necrosis in the community tray (a); yellowish leaves and leaf drop, dark brown roots, and plantlet death (b); Dendrobium roots showing brown discoloration with white fungal mycelia, the root system totally dried out and the pseudobulb turning brown (c); Paphiopedilum showing decline symptoms including yellowish leaves and brown and rotten collars caused by Rhizoctonia sp. from contaminated coconut husks used as the planting medium (d) (P. Smitamana)

• Cultural practices – Growers should inspect newly introduced plants and keep them in an area removed from general production for 4–6 weeks to observe the health of the plants and make sure that they are free from any diseases. Transplant uninfected pseudobulbs into pathogen-free media and new pots. Adjust the pH of the irrigation water and medium to 6.0–6.5 to help reduce infection by Fusarium. • Sanitation – For slightly infected Vanda, Vanilla, and Rhyncostylis with good aerial roots, cut above the diseased portion using a sterile cutting tool, pot in a fresh medium, remove and discard or burn infected tissue, and apply an appropriate fungicide or biocontrol (see below). • Fungicides and biocontrols – Use biocontrol agents, e.g., Trichoderma harzianum, by incorporation into fresh media in order to prevent contamination by pathogens. If chemical control is needed, apply cyprodinil + fludioxonil, chlorothalonil, azoxystrobin, or thiophanate methyl when early symptoms are observed, being sure to rotate different classes of fungicides. Prochloraz and tebuconazole were very effective in reducing Fusarium bulb and root rot in Cymbidium goeringii (Jee et al. 2003). Quaternary ammonium salts, sodium hypochlorite, and other materials should be used to disinfest benches and

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containers (see chapter “▶ Sanitation in Florists’ Crops Disease Management” for more information).

2.7

Leaf Spots (Fusarium, Phyllosticta, Guignardia, Septoria, Cercospora spp.)

Geographic occurrence and impact. Leaf spots of orchids can be caused by many fungi. Symptoms in the diseased orchid are varied based on the causal agent and host. Many orchids are susceptible to these pathogens, e.g., Brassolaeliocattleya, Cattleya, Cymbidium, Dendrobium, Epidendrum, Laelia, Laeliocattleya, Odontoglossum, Oncidium, Phalaenopsis, and Vanda. Leaf spots of orchids are found worldwide: Australia, Brazil, Japan, the Netherlands, Thailand, and the USA (Hawaii) (Daly et al. 2013; Ichikawa and Aoki 2000; Jones 2003; Kamjaipai 1983; Silva and Pereira 2007; Silva et al. 2008; Uchida and Aragaki 1980; Uchida 1994). Leaf spots are common in most nurseries and can lead to serious losses in shipping containers due to the high moisture and low light conditions which favor the growth and infection of the pathogens. Symptoms/signs. Fusarium spp. – Two spot types; yellow and black are caused by F. subglutinans and F. proliferatum, respectively, on Cymbidium. The yellow spot symptoms start as water soaked patches on the leaves then the lesions enlarge with sunken centers and turn reddish-brown surrounded with a yellowish swelling without a definitive boarder. In the case of black spot, small black speckles are found at the early stage, then the lesions enlarge to form irregular, angular black spots; some yellow halos are found in certain Cymbidium cultivars. Phyllosticta spp. – Symptoms may start anywhere on the leaf, flower, or pseudobulb. Tiny yellow and slightly sunken spots are initially observed on the leaves. As the lesions enlarge, they become more sunken with round or oval shapes (Fig. 6a). The fungal growth within the tissue turns the lesion tan or brownish as the disease develops. Severely infected leaves may drop prematurely and a black web-like pattern formed by the fungus is observable (Fig. 6b). Spot symptoms on flowers are small, faint, and commonly blue or lavender on purple cultivars. Guignardia spp. – Symptoms start as tiny, dark purple, elongated lesions which can be found on either leaf surface. With age, the lesions enlarge and run parallel to the veins and form elongated purple streaks. Large irregular lesions are formed by the merging of the spots, and the center of the lesions turns tan. Black fruiting bodies of the fungus develop in the affected area and can be seen as tiny raised black spots. Cercospora spp. – Cercospora attacks the underside of the leaf and induces small yellow spots. After the disease progresses, yellow green areas are observed on the top surface of the leaf (Fig. 6c). Subsequently, the spots enlarge and form irregular patterns with slightly sunken areas which turns brown to black. The necrotic lesions

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Fig. 6 Leaf spots caused by Phyllosticta sp. in Vanda: sunken lesions (a); merging lesions with black fungal growth in the leaf tissue (b); yellow green area with brown spots on Dendrobium leaf caused by Cercospora sp. (c) (P. Smitamana)

can enlarge to cover the whole leaf area and severely infected leaves may drop prematurely. Septoria spp. – Septoria can infect both sides of the leaf. Primary symptoms start from small sunken yellow lesions which gradually enlarge and form dark brown to black, circular, or irregular lesions. Lesions may merge to form irregular areas on the leaf which may prematurely drop. Biology and epidemiology. Phyllosticta belongs to the Sphaeropsidales and produces asexual spores in pycnidia, small, black globose, elongate, or cup-like fruiting bodies. Pycnidia may be superficial or immersed in the tissue. Conidia are hyaline, single-celled, globose, or ovoid (Sumbali 2005). High light levels and dry conditions inhibit the growth and spread of the fungus, whereas low-light and high humidity, or rainy weather, promote Phyllosticta outbreaks (Jones 2003). Guignardia is the perfect stage of Phyllosticta and belongs to the Ascomycetes, and produces eight single celled ascospores in perithecia (Silva and Pereira 2007). Cercospora belongs to the Dermatiaceae and produces dark colored hyphae and long slender hyaline or dark, straight to curved, multicellular conidia on short dark conidiophores. Clusters of conidiophores arise from the plant surface through the stoma and form conidia on the growing tips (Agrios 2005). No fruiting bodies are formed (Sumbali 2005). Septoria belongs to the Sphaeropsidales like Phyllosticta and produces long, slender, hyaline, generally curved conidia with one or more septa (Sumbali 2005).

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Conidia of these fungi are easily detached from the conidiophores, and ascospores discharge from perithecia into the air and are carried great distances. Conidia are splashed from older diseased leaves to other leaves nearby and infect new plants. Management. • Cultural practices – Good sanitation with adequate air movement is important to control these diseases. Discard severely infected plants. Collect all fallen leaves, old or dry pseudobulbs, and sheaths from pots, benches, and the ground and discard or burn them. Water the orchids early in the day to allow leaves to dry before dark. • Fungicides – Spray with mancozeb, chlorothalonil, or another effective fungicide to kill the fungal spores and reduce the inoculum level (see chapter “▶ Fungicides and Biopesticides for Florists’ Crops Disease Management” for more information).

2.8

Anthracnose [Colletotrichum gloeosporioides (syn. Glomerella cingulata)]

Geographic occurrence and impact. This is a disease that commonly affects Dendrobium, Vanda, Cattleya, Phalaenopsis, and Ceologyne orchids. Anthracnose in orchids has been reported in Australia, India, Thailand, and the USA (Florida) (Bottom 2016; Chowdappa et al. 2014; Daly et al. 2013; Jadrane et al. 2012; Kamjaipai 1983; McMillan 2011; Prapagdee et al. 2008; Prapagdee et al. 2012). This disease is considered only a minor pathogen and is usually a result of some type of injury to the leaf, whether it is mechanical, chemical, or insect damage. Nevertheless, anthracnose can result in reduced plant quality and growth; disease development may be sporadic as it is affected by levels of pathogen inoculum and environmental conditions. Symptoms/signs: Colletotrichum infects the aerial portion of the plant of which leaves are most often attacked. Infected leaves show brown discolorations which are irregularly shaped sunken lesions. Leaf tips turn brown and the affected area gradually enlarges toward the base. With age, the lesions become dark brown or light gray patches, and sometimes brownish black concentric rings of fruiting bodies or numerous dark bands develop across the leaf. Flower infection is characterized by water soaked spots and black or brown pustules which are usually raised and occur on the underside of old sepals and petals. The spots may merge and cover the entire flower. In white flowered Phalaenopsis, the infected petals show anthracnose-like, sunken lesions which are surrounded by a ring of green tissue. Biology and epidemiology. Colletotrichum gloeosporioides is a member of the Melanconiaceae and is the imperfect stage of the ascomycete Glomerella cingulata. The conidia are produced in acervuli and are single celled, hyaline, ovoid, cylindrical, typically elongated with round ends and slightly narrower in the middle. Large brown or black setae are formed among the conidiogenous cells (Agrios 2005; Sumbali 2005). The conidia are disseminated by air currents, wind, rain, or irrigation water. The pathogen is most active in warm weather when light is low and moisture

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Fig. 7 Typical symptom of anthracnose on the black orchid (Ceologyne pandurata): the early symptom showing a brownish water-soaked area starting at the leaf tip (a); later stage lesions showing dark concentric rings on the upper leaf surface (b) and lower leaf surface (c) (P. Smitamana)

is high. Colletotrichum can survive or overwinter on plant residue in soil or on infected plants. Anthracnose is common after orchids pass through a stress period, e.g., cold or hot temperature, mechanical injuries, or chemical damage. Management. • Cultural practices – Good sanitation and air movement, lower temperatures (if possible), and increased light may help reduce the spread and severity of this disease. • Fungicides and biocontrols – Alternation of protectant fungicides like mancozeb and systemic fungicides such as thiophanate methyl is recommended. The bacterium Bacillus subtilis was shown to reduce anthracnose caused by C. gloeosporiodes in Paphiopedilum concolor in Thailand (Kuenpech and Akarapisan 2014). Application of culture filtrates of the bacterium Streptomyces hygroscopicus prevented anthracnose on orchid (Propagdee et al. 2008).

2.9

Rust (Sphenospora kevorkianii)

Geographic occurrence and impact. Sphenospora kevorkianii has been found infecting orchids in Brazil and in the USA (Linder 1944; Pereira et al. 2002; Pereira and Silva 2009). Known hosts are Cyrtopodium punctatum, Epidendrum difforme, E. secundum, E. xanthinum, Notylia lyrata, Pleurothallis mentigera, Prescottia sclerophylla, Stanhopea graveolens, and Zygostates lunata which are epiphytes found in tropical forests. Symptoms/signs. Rust fungi are very easy to identify from their rust colored spore masses on leaves and stems. The symptoms start out as flecks or spots and grow into bumps, and most commonly appear on the underside of leaves.

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Biology and epidemiology. Sphenospora kevorkianii produces uredospores in ovoid, ellipsoid, or subspherical uredinia on the lower leaf surface (Pereira et al. 2002). The uredospores are easily detached and dispersed by wind, rain, and irrigation water. Management. Cut off and discard or burn the infected parts. Discard or burn severely infected plants. Reducing nitrogen fertilizer application may help. If needed, spray with appropriate systemic fungicides (see chapter “▶ Fungicides and Biopesticides for Florists’ Crops Disease Management” for more information).

3

Bacterial and Phytoplasma Diseases

3.1

Leaf Spot (Acidovorax avenae subsp. cattleyae)

Geographic occurrence and impact. This disease has been reported in Australia, China, India, Italy, Korea, Taiwan, Poland, and the USA (Florida) (Borah et al. 2002; Cating and Palmeteer 2011; Ding et al. 2010; Hseu et al. 2012; Kim et al. 2015; Li et al. 2009; Lin et al. 2015; Pulawska et al. 2013; Scortichini et al. 2005; Stovold et al. 2001). Acidovorax avena is a pathogen of quarantine importance and causes serious losses to orchid production especially in Cattleya, Cypripedium, Dendrobium, Oncidium, Phalaenopsis, and Vanda. Symptoms/signs. Symptoms usually start as small brown soft water-soaked spots. The spots turn black forming cavities in the parenchyma which quickly expand over the entire leaf. Eventually, the pathogen invades the growing point of the plant causing death (Fig. 8a, b). Phalaenopsis, due to its succulent leaves, is generally reported in many countries as the most susceptible genus. Biology and epidemiology. Acidovorax avenae subsp. cattleyae is a gramnegative rod shaped, nonfluorescent bacterium, which moves using a single polar flagellum. The bacteria are easily spread by rain, irrigation, and contaminated tools commonly used in the nurseries. High moisture and temperature promote the growth of bacteria and accelerate the progress of disease development. Management. Management of bacterial orchid diseases is challenging and must rely on an integrated approach. • Cultural practices – Avoid overhead watering to minimize the dispersal of the pathogen in nurseries or water early in the morning to avoid long hours of leaf wetness. If the disease is found, reducing the use of nitrogen fertilizer and increasing potassium fertilizer has been associated with disease suppression. • Sanitation – Growers should discard severely infected plants. Trim and discard diseased areas from slightly infected plants and keep them in an area removed from production for monitoring. Early detection of disease outbreaks is essential. Apply a bactericide as soon as the disease is observed (see below). Clean tools after each use by soaking in a 10% bleach solution. • Bactericides – If the infection becomes widespread, growers may dip plants in a labeled quaternary salt material, orthophenylphenol, or natriphene, or spray with

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Fig. 8 Leaf spot of Phalaenopsis caused by Acidovorax avenae: brownish water soaked area which begins at a mechanical injury (a) and the soft rot of leaf (b) (P. Smitamana)

a copper compound. Formulations of hydrogen peroxide may help reduce the disease. If antibiotic use is allowed, streptomycin, tetracycline, or a mixture of both may be sprayed. Small-scale testing for phytotoxicity is recommended. However, reliance on chemical control alone for management of bacterial diseases of orchids is most often ineffective.

3.2

Soft Rot [Dickeya chrysanthemi (syn. Pectobacterium chrysanthemi); D. dieffenbachiae]

Geographic occurrence and impact. This disease mainly affects Oncidium, Vanda, Tolumnia, Cattleya, Dendrobium, Phalaenopsis, and Miltoniaorchids and has been reported in China, Taiwan, and the USA (Florida) (Cating et al. 2008; Cating et al. 2009b; Li et al. 2009; Cating et al. 2009b; Hseu et al. 2012; Keith and Sewake 2009; Zhou et al. 2012). This is an important quarantine disease that affects the export and import of orchids in many countries. Symptoms/signs. The symptoms are typical of other bacterial soft rots and start with water-soaked tissue at the infection site. On Oncidium, the initial spots are commonly found at the base of the stem, and the lesions expand and become elongated. Gradually, the stem and leaf tissues become soft and watery. Bacterial ooze can be observed by cutting the edge of infected stem and leaf tissues and dipping them in a beaker or Petri dish containing water; bacterial streaming can be observed from the cut tissue by eye or under a stereomicroscope. Infected Phalaenopsis plants initially show water-soaked, pale-to-dark brown pinpoint spots on leaves; some may be surrounded by a yellow halo. Under high humidity and temperature, the spots expand rapidly and extend over the leaf which turns a light tan color with a darker brown border in a few days. A typical odor of rotten tissue can be detected. When the bacteria invade the stem, plant death follows. In Miltonia, Cattleya, and Oncidium, symptoms start with small water-soaked lesions which become a dark brown to black color and gradually expand throughout the leaves which subsequently rot. Some Dendrobium cultivars appear to be resistant

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Fig. 9 Symptoms of orchid viruses. CymMV: small, discolored Dendrobium flower (a), twisted shoot with yellow streaks (b); ORSV: distorted Dendrobium flower with color-breaking (c), mosaic and yellow patches in Dendrobium (d); OFV: yellow concentric rings in Phalaenopsis leaf (e); INSV: chlorotic patches and necrotic ringspots and distortion in Oncidium leaves (a-e, P. Smitamana; f, courtesy of S. Koike)

to this disease and only small water-soaked spots are produced. In the susceptible types, the spots expand and coalesce to form larger necrotic areas; after that, the infected leaves dry out. If the bacteria invade the pseudobulbs, they turn soft and watery. In Vanda, infected areas show water soaked spots, some are brown color, and the leaves are rapidly rotted.

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Biology and epidemiology. Dickeya spp. are gram-negative bacteria, facultative, anaerobic rods with peritrichous flagella. Infection by the pathogen normally starts in cutting wounds or other mechanical injuries. Drops of bacterial ooze from the infected leaves or stems are dispersed by rain, irrigation, and wind. The bacteria enter and colonize plants mostly at the basal part of leaves or pseudobulbs. Management. Management of soft rot is the same as for Acidovorax leaf spot listed above.

3.3

Black Spot [Burkholderia gladioli (syn. Pseudomonas gladioli)]

Geographic occurrence and impact. Burkholderia gladioli was first characterized as the causal agent of leaf and corm disease in gladiolus. This pathogen is now known to cause serious losses in many plants especially orchids, e.g., Phalaenopsis, Cattleya, Dendrobium Oncidium, Odontioda, and Miltonia. This pathogen is found in tropical and subtropical zones of such countries as China, the USA (Hawaii), Indonesia, Taiwan, and Thailand (Chuenchitt et al. 1983; Hseu et al. 2012; Joko et al. 2014; Kamjaipai 1983; Keith et al. 2005; Lee et al. 2013; Takahashi et al. 2004; Uchida 1995; You et al. 2016). Symptoms/signs. Infected leaves initially show small dark green to brown spots surrounded by a water-soaked area. Sometimes only small brown spots surrounded by yellow halos, usually circular, or in rare cases irregularly shaped occur. Under high humidity or after rain, the wet rot area rapidly expands and causes soft rot of the whole leaf, especially the succulent leaves of Phalaenopsis. The bacteria can invade the stem or pseudobulbs and sometimes cause defoliation. Under low humidity, the infected leaf may dry and crack in the center of the lesion. However, if the humidity increases, bacterial ooze can be seen on the cracks and the rotten tissues. Symptoms on the young plants spread rapidly and cause the whole plant to die. Biology and epidemiology. Burkholderia gladioli is a gram-negative rod that may be straight or slightly curved. It is aerobic, catalase positive, urease positive, and non-sporeforming. Like other phytopathogenic bacteria, it is easily spread by water splash from rain, overhead sprinklers, and wind-blown water. Management. Management of soft rot is the same as for Acidovorax leaf spot listed above.

3.4

Phyllody (“Candidatus Phytoplasma” sp.)

Geographic occurrence and impact. Only a Sarcochilus hybrid in Australia has been reported to be infected by a phytoplasma (Gowanlock et al. 1998). Symptoms/signs. The infected Sarcochilus hybrid exhibited phyllody, the abnormal development of floral parts into leafy structures, and stunting. Biology and epidemiology. Phytoplasmas are bacteria-like microorganisms lacking cell walls, but bounded by a unit membrane. Phytoplasmas are found located in the sieve tubes of phloem. Transmission of phytoplasmas is by phloem-feeding

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insects in the Cicadellidae family and Fulgoroidea superfamily such as leafhoppers and planthoppers, respectively. Management. Avoid introducing diseased plants into nurseries. Remove and burn symptomatic plants.

4

Viral Diseases

Geographic occurrence and impact. Viruses are among the most damaging pathogens of orchids because of their debilitating effects on flower production and quality and extreme difficulty to control in a vegetatively propagated crop. Orchids can be infected by a very wide range of viruses. The most common and economically important orchid viruses worldwide are Cymbidium mosaic virus (CymMV) and Odontoglossum ringspot virus (ORSV), also known as Tobacco mosaic virus-orchid strain (TMV-O); and these two viruses are often found in mixed infections. Other orchid viruses of note include: Orchid fleck virus (OFV), Impatiens necrotic spot virus (INSV), Vanilla mosaic virus (VanMV), and Calanthe mild mosaic virus (CalMMV). The geographic occurrence and orchid host range of these viruses are indicated in Table 1. Orchids have also been reported to host a large number of other viruses in the following genera: Carmovirus – Carnation mottle virus (CarMV) Closterovirus – Dendrobium vein necrosis virus (DVNV) Cucumovirus – Cucumber mosaic virus (CMV) Nepovirus –Tomato ringspot virus (ToRSV) Potyvirus – Bean yellow mosaic virus (BYMV); Ceratobium mosaic virus (CerMV); Clover yellow vein virus (ClYVV); Colombian datura virus (CDV); Dasheen mosaic virus (DsMV); Dendrobium mosaic virus (DeMV); Diurus virus Y (DVY); Habenaria mosaic virus (HaMV); Pecteilis mosaic Virus (PcMV); Phalaenopsis chlorotic spot virus (PhCSV); Spiranthes mosaic virus 2 (SpiMV2); Sarcochilus virus Y (SVY); Spiranthes mosaic virus 3 (SpiMV3); Turnip mosaic virus (TuMV); Vanilla necrosis virus (VanNV) Rhabdovirus – Colmanara mosaic virus (CoIMV); Dendrobium ringspot virus (DRV) Tobravirus – Tobacco rattle virus (TRV) Tombusvirus – Cymbidium ringspot virus (CymRSV); Lisianthus necrosis virus (LNV): Tomato bushy stunt virus (TBSV) Tospovirus – Capsicum chlorosis virus (CaCV); Tomato spotted wilt virus (TSWV)

Symptoms/signs. Orchids can show variation in viral symptoms that depend on the virus, virus strain, mixed viral infections, and orchid variety. In addition, different viruses can cause similar symptoms in the same orchid, making diagnosis of virus infection by symptoms alone unreliable. Common symptoms of virus infection of orchids are presented in Table 1.

Geographic occurrence Argentina, Brazil, China, Colombia, Cook Islands, Costa Rica, French Polynesia, Guam, India, Indonesia, Japan, New Caledonia, Netherlands, Niue, Palau, Puerto Rico, Reunion Island, Samoa, Singapore, Taiwan, Thailand, Tonga, USA, Vanuatu, Venezuela

Argentina, Brazil, China, Colombia, Cook Islands, Costa Rica, Fiji, French Polynesia, India, Indonesia, Japan, Netherlands, New Zealand, Niue, Puerto Rico, Reunion Island, Singapore, Taiwan, Thailand, Tonga, USA, Vanuatu, Venezuela

Virus, genus Cymbidium mosaic virus (CymMV), Potexvirus

Odontoglossum ringspot virus (ORSV), (also known as TMV-orchid strain and TMV-O), Tobamovirus

Table 1 Selected orchid viruses

Mechanical

Transmission Mechanical

A wide range including: Aerides, Bulbophyllum, Calanthe, Cattleya, Cymbidium, Dendrobium, Oncidium, Phalaenopsis, and Vanilla

Orchid hosts A wide range including: Aranthera, Arachnis, Calanthe, Cattleya, Cymbidium, Dendrobium, Gromatophyllum, Phalaenopsis, Phaius, Oncidium, Rynchostylis, Vanda, and Vanilla

Chlorotic or necrotic sunken lesions on leaves and flowers. In highly susceptible varieties, flowers may be smaller, distorted, and exhibit color breaking (Fig. 9c, d). Root tip necrosis is also a common symptom

Symptoms Chlorotic mosaic patterns, seen clearly on the youngest leaves in many orchid varieties, to black necrotic streaks, spots, or rings and sunken patches on Cattleya orchids. Flower size reduction with abnormal coloration and twisted young shoots with chlorotic streaks are common symptoms on young Dendrobium shoots (Fig. 9a, b)

(continued)

References Baker et al. 2007a; Cánovas et al. 2016; Davis and Ruabete 2010; Elliot et al. 1996; Farreyrol et al. 2001; Freitas-Astua et al. 1999; Gara et al. 1996; Hu et al. 1993; Inouye and Gara 1996; Jensen 1952; Kamjaipai 1983; Sherpa et al. 2003; Singh et al. 2007; McMillan et al. 2006; Sutrabutra 1989; Tanaka et al. 1997; Wong et al. 1994; Zheng et al. 2010; Zhou et al. 2004 Cánovas et al. 2016; Davis and Ruabete 2010; Elliot et al. 1996; Farreyrol 2001; FreitasAstua et al. 1999; Hu et al. 1993; Inouye and Gara 1996; Kamjaipai 1983; McMillan et al. 2006; Pearson et al. 2006; Sherpa et al. 2006; Sutrabutra 1989; Tanaka et al. 1997; Thomson and Smirk 1967; Wong et al. 1994; Zhou et al. 2004

Diseases of Orchid 21

Mechanical; (Myzus persicae) Mechanical; aphid

Japan French Polynesia, Reunion Island

Calanthe, Phalaenopsis, and Tetragonia expansa Vanilla

Phalaenopsis Vanilla

Mechanical Mechanical; green peach aphid (Myzus persicae)

Taiwan Cook Islands, Fiji, French Polynesia

Phalaenopsis chlorotic spot virus Potyvirus Vanilla mosaic virus (VanMV), Potyvirus Calanthe mild mosaic virus (CalMMV), Potyvirus Cucumber mosaic virus Cucumovirus

Orchid hosts Cymbidium, Maxillaria, Renanthera, Phalaenopsis

Cymbidium, Dendrobium, and Phalaenopsis and many other ornamentals Phalaenopsis

China, Taiwan, USA Taiwan

Impatiens necrotic spot virus (INSV), Tospovirus Capsicum chlorosis virus (CaCV), Tospovirus

Transmission False spider mite (Brevipalpus californicus)

A number of thrips species including the Western flower thrips (Frankliniella occidentalis) Mechanical

Geographic occurrence Australia, Brazil, Costa Rica, Denmark, Germany, Japan, Korea, USA

Virus, genus Orchid fleck virus (OFV), Dichorhavirus

Table 1 (continued)

Chlorotic spots Chlorotic and necrotic flecks, mosaic and deformation in leaves; dieback of shoots Mild leaf mosaic leaf and flower breaking Severe stunt, conspicuous stem and leaf deformation

Symptoms Yellow flecks and yellow or necrotic ringspot lesions on leaves (Fig. 9e). In Dendrobium, Miltonia, Odontoglossum, Oncidium, and Paphiopedilum, the chlorotic areas often with necrotic centers or rings Large chlorotic or necrotic ringspots on leaves; leaf yellowing and distortion (Fig. 9f) Chlorotic spots with concentric necrosis

Gara et al. 1998 Farreyrol et al. 2001; Farreyrol et al. 2010

Bakardjieva et al. 1998; Baker et al. 2007b; Koike and Mayhew 2001; Zhang et al. 2010; Zheng et al. 2008a Zheng et al. 2008a Zheng et al. 2008b Davis and Ruabete 2010

References Brunt et al. 1997; Dietzgen et al. 2014; Freitas-Astúa et al. 2002; Kondo et al. 2003; Kitajima et al. 2001; Kubo et al. 2009

22 P. Smitamana and R.J. McGovern

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Biology and epidemiology. As obligate pathogens, viruses need a living host for their survival and replication. However, the molecular structure of Tobamoviruses, such as ORSV, makes them resistant to degradation for long periods on tools and in soil when they are released from host tissues. A number of important orchid viruses, such as CymMV, ORSV, VanMV, and CalMMV, are mechanically transmissible in plant sap. This characteristic is very important for virus spread in a vegetatively propagated crop such as orchid. In addition, potyviruses including VanMV and CalMMV are vectored in a nonpropagative (nonpersistent) manner on the stylets of aphids such as Myzus persicae. INSV, a tospovirus, is vectored in a circulative and propagative manner (it enters and replicates within the hemolymph of the insect) by a number of thrips species including the Western flower thrips, Frankliniella occidentalis. Only the first and early second larval stages are able to acquire tospoviruses, and only immature thrips that acquire these viruses or adults derived from such immatures are vectors. Adult thrips remain viruliferous for life; but tospoviruses are not transovarial. INSV also infects a large number of other ornamentals which can serve as reservoirs of the virus. OFV is spread by the false spider mite, Brevipalpus californicus. Management. Once infected by a virus, unlike with other pathogens, nothing short of complicated and exacting tissue culture procedures can rid plants of viruses, and even these extreme measures are not always successful. Therefore, management of orchid viruses must be based on prevention of infection by using virus-free propagative material and good sanitation practices. Isolate suspect plants and new introductions and monitor for symptoms for 6 weeks or until proven by a diagnostic test (ELISA, PCR, etc.) to be virus-free. Promptly discard or destroy by burning all confirmed diseased plants. Use only sterile cutting tools; soak the tools in a 10% bleach solution or saturated trisodium phosphate solution for 10 min; or use disposable razor blades when dividing plants and cutting flowers. Wear latex gloves when handling plants and discard those gloves when finished. Always change to new gloves when handling a new plant. Disinfest containers by first washing with soap to remove residual organic matter, then soak in a 20% bleach solution for 1 h, after that soak them for 1 h in a quaternary ammonium salt solution as per label instructions. Exclude insect-vectored viruses by using fine mesh screening, pesticides, and/or biological controls. Refer to the chapter “▶ Insect Management for Disease Control in Florists’ Crops” for additional information.

5

Nematode Diseases

5.1

Foliar Nematodes (Aphelenchoides spp.)

Geographic occurrence and impact. A number of Aphelenchoides spp. including A. besseyi, A. fragariae, and A. ritzemabosi have been reported to attack orchids in Singapore, Thailand, and the USA (Florida) (Esser et al. 1988; Kawate and Sewake 2014; Latha et al. 1999). Because warm, wet greenhouse environments are conducive to foliar nematode infection, the occurrence of these nematodes in orchids is

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undoubtedly more widespread. This disease affects many orchids including Cattleya, Cymbidium, Dendrobium, Oncidium, Paphiopedilum, and Vanda. Losses from uncontrolled foliar nematodes have been estimated at up to 90–95% in potted orchid production in Hawaii (Kawate and Sewake 2014). Symptoms/signs. Foliar nematodes infect immature buds and prevent flower formation and development on Vanda; diseased buds abscise or adhere and become blackened. Symptoms on Dendrobium leaves include yellow-green blotches which become brown as the lesions age. On Oncidium, browning of foliar buds and dark streaks in pseudobulbs can occur (Uchida and Sipes 1998). Biology and epidemiology. A film of water enables foliar nematodes to move up stems and over leaf surfaces to seek new infection sites made by wounds or natural openings such as stomates. Infection and symptom expression are, therefore, enhanced by warm, wet conditions. The Aphelenchoides spp. that attack orchids have a very broad host range that includes many ornamentals and other cultivated crops. Adults and 4th stage juveniles feed on mesophyll and epidermal tissue within leaves (endoparasitism). These tissues collapse and turn brown. Females lay eggs within green leaf tissue. Foliar nematodes also feed externally on stems, buds, and flowers (ectoparasitism). Adults and 4th stage juveniles can survive in association with living and desiccated plant tissues for long periods. Foliar nematodes can be disseminated by splashing water, infected plant propagative material, and debris (Kohl 2011). Management. Prevention of nematode introduction and good sanitary practices are the keys to managing foliar nematodes. Isolate suspicious plants and new introductions to monitor for symptom expression. Discard plant debris and infected plants. All surfaces coming in contact with plants (pots, stakes, tools, etc.) should first be cleaned with soap and water and surface-disinfested with a 10% solution of household bleach for 10 min after each use (Uchida and Sipes 1998). Growers should avoid wetting plant surfaces as much as possible. Increasing air circulation through increased plant spacing will help to dry plants. Only propagate from clean material. Hot water treatment of Vanda cuttings for 15 min at 46  C (115  F) or 5–10 min at 49  C (121  F) eliminated nematodes without injuring the plants (Uchida and Sipes 1998).

References Agrios GN (2005) Plant pathology, 5th edn. Elsevier Academic Press, San Diego. 922 p Atwood JT Jr (1986) The size of the Orchidaceae and the systematic distribution of epiphyticorchids. Selbyana 9:171–186 Bag TK (2004a) Two new orchid hosts of Sclerotium rolfsii from India. Plant Pathol 53:255 Bag TK (2004b) Occurrence of orchid wilt (Sclerotium rolfsii Sacc.) in Cymbidium and its hybrids. Sci Cult 70:287–288 Bakardjieva N, Denkova S, Hristova D (1998) Tomato spotted wilt virus on ornamental species in Bulgaria. Biotechnol Biotechnol Equip 12(2):49–52. http://www.tandfonline.com/doi/abs/10. 1080/13102818.1998.10818987. Accessed 8 Oct 2015

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Singh MK, Sherpa AR, Hallan V, Zaidi AA (2007) A potyvirus in Cymbidium spp. in northern India. Aust Plant Dis Notes 2:11–13 Stovold GE, Bradley J, Fahy PC (2001) Acidovorax avenae subsp. cattleyae (Pseudomonas cattleyae) causing leafspot and death of Phalaenopsis orchids in New South Wales. Australas Plant Pathol 30:73–74 Su JF, Lee YC, Chen CW, Hsieh TF, Huang JH (2010) Sheath and root rot of Phalaenopsis caused by Fusarium solani. Acta Hortic 878:389–394. doi:10.17660/ActaHortic.2010.878.49 Sumbali G (2005) The fungi. Alpha Science International, 298 p Sutrabutra T (1989) Virus and virus-like of important plants in Thailand. Funny Publishing, Bangkok. 310 p (in Thai) Takahashi Y, Takahashi K, Watanabe K, Kawano T (2004) Bacterial black spot caused by Burkholderia andropogonis on Odontoglossum and intergeneric hybrid orchids. J Gen Plant Pathol 70:284–287 Tanaka S, Nishii H, Ito S, Kameya-Iwaki M, Sommartya P (1997) Detection of Cymbidium mosaic potexvirus and Odontoglossum ringspot tobamovirus from Thai orchids by rapid immunofilter paper assay. Plant Dis 81:167–170 Tao YH, Ho HH, Wu YX, HE YQ (2011) Phytophthora nicotianae causing Dendrobium blight in Yunnan Province, China. Int J Plant Pathol 2:177–186 Tara BP, McMillan RT, Graves WR (2003) Sclerotium rolfsii Southern blight of Brassidium hybrid orchid. Proc Fla State Hort Soc 116:195–196 The Plant List (2013) Version 1.1. Published on the Internet; http://www.theplantlist.org/1.1/ browse/A/Orchidaceae/. Accessed 4 Apr 2016 Thomson AD, Smirk BA (1967) An unusual strain of tobacco mosaic virus from orchids. N Z J Bot 5:197–202. doi:10.1080/0028825X.1967.10428740. Accessed 4 Mar 2016 Uchida JY (1994) Diseases of orchids in Hawaii. Plant Dis 78:220–224 Uchida J (1995) Bacterial diseases of Dendrobium, Research extension series, vol 158. Institute of Tropical Agriculture and Human Resources. University of Hawaii Uchida JY, Aragaki M (1980) Nomenclature, pathogenicity, and conidial germination of Phyllostictina pyriformis. Plant Dis 64:786–788 Uchida JY, Aragaki M (1991a) Phytophthora diseases of orchids in Hawaii, Research extension series, vol 129. College of Tropical Agriculture and Human Resources. University of Hawaii Uchida JY, Aragaki M (1991b) Fungal diseases of Dendrobium flowers, Research extension series, vol 133. College of Tropical Agriculture and Human Resources. University of Hawaii Uchida JY, Sipes BS (1998) Foliar namatodes on orchids in Hawaii. University of Hawaii at Manoa, PD-13, 7 pp. http://www.ctahr.hawaii.edu/oc/freepubs/pdf/PD-13.pdf Wedge D, Elmer H (2008) Fusarium wilt of orchids. Int Commer Orchid Growers Organ (ICOGO) Bull 2:9–10 Wong SM, Chng CG, Lee YH, Tan K, Zettler FW (1994) Incidence of cymbidium mosaicand Odontoglossum ringspot viruses and their significance in orchid cultivation in Singapore. Crop Prot 13:235–239 You Y, Lü FB, Zhong RH, Chen HM, Li HP, Liu JM, Zhang JH (2016) First report of bacterial brown spot in Phalaenopsis spp. caused by Burkholderia gladioli in China. Plant Dis PDIS-0915-0963-PDN. http://apsjournals.apsnet.org/doi/abs/10.1094/PDIS-11-15-1373-PDN. Accessed 25 Mar 2016 Zhang Q, Ding YM, Li M (2010) First report of impatiens necrotic spot virus infecting Phalaenopsis and Dendrobium orchids in Yunnan Province, China. Plant Dis 94:915–915 Zheng YX, Chen CC, Yang C, Yeh SD, Jan FJ (2008a) Identification and characterization of a tospovirus causing chlorotic ringspots on Phalaenopsis orchids. Eur J Plant Pathol 120:199–209 Zheng YX, Chen CC, Chen YK, Jan FJ (2008b) Identification and characterization of a potyvirus causing chlorotic spots on Phalaenopsis orchids. Eur J Plant Pathol 121:87–95 Zheng YX, Shen BN, Chen CC, Jan FJ (2010) Odontoglossum ringspot virus causing flower crinkle in Phalaenopsis hybrids. Eur J Plant Pathol 128:1–5

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Zhou G, Chen X, Li M, Zhou J, Tang T, Feng S, Guo L, Zhang W (2004) Identification and detection of two major viruses infecting orchids by molecular technique. Virol Sin 19:149–152 Zhou JN, Lin BR, Shen HF, Pu XM, Chen ZN, Feng JJ (2012) First report of a soft rot of Phalaenopsis aphrodita caused by Dickeya dieffenbachiae in China. Plant Dis 96:760–760

Diseases of Proteaceae Brett A. Summerell

Abstract

The Proteaceae are a southern hemisphere family of plants found predominantly in southern Africa and Australia. Included in the family are Protea, Leucospermum, and Leucodendron from southern Africa and Banksia, Telopea, and Grevillea from Australia all of which are important cut flower crops. All of these genera are susceptible to a range of diseases, especially root rot diseases caused by Phytophthora, Armillaria, and Fusarium which can limit production and leaf spot diseases which both limit production and impair the aesthetics of the cut flower. Management of these diseases is complex and includes chemical, biological, cultural, and physical control, as well as host resistance. Keywords

Phytophthora spp • Leaf spots • Fusarium spp • Mycosphaerella leaf blotch • Integrated disease management

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-Like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Leaf Spot Diseases (A Broad Range of Fungal Species) . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Botryosphaeria Leaf Blight and Stem Canker (Neofusicoccum sp.) . . . . . . . . . . . . . . . . . 2.3 Botrytis Seedling and Flower-Head Blight (Botrytis cinerea Pers.: Fr.) . . . . . . . . . . . . . 2.4 Colletotrichum Anthracnose Disease Complex/Colletotrichum Tip Dieback (Colletotrichum spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Elsinoe Scab Disease (Elsinoe spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 Pyrenophora Blight (Pyrenophora spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7 Phytophthora Root and Collar Rot and Sudden Death Disease (Phytophthora spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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B.A. Summerell (*) Royal Botanic Gardens and Domain Trust, Sydney, NSW, Australia e-mail: [email protected] # Her Majesty the Queen in Right of Australia 2017 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florist's Crops Diseases, Handbook of Plant Disease Management, DOI 10.1007/978-3-319-32374-9_22-1

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2.8 Pythium Root Rot (Pythium spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.9 Armillaria Root Rot (Armillaria sp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.10 Rhizoctonia Root Rot (Rhizoctonia sp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.11 Fusarium Wilt (Fusarium oxysporum Schlect.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.12 Ilyonectria Black Foot Rot [Ilyonectria spp. (formerly Cylindrocarpon spp.)] . . . . . 3 Bacterial and Phytoplasma Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Pseudomonas Leaf Spot [Pseudomonas syringae Pathovar “Pv. Proteae”] . . . . . . . . . . 3.2 Phytoplasma Diseases: Witch’s Brooms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Viral Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Introduction

The Proteaceae is one of the most iconic groups of plants found in the southern hemisphere. It is widespread, reflecting its Gondwanic heritage (Barker et al. 2007), and is highly speciose adapting to diverse ecosystems throughout southern Africa, Australia, and to a lesser extent South America (Rebelo 2001; Weston and Barker 2006). Members of the family are also found in New Zealand, New Caledonia, Fiji, and Vanuatu, and there are a small number of species in Southeast Asia (Weston and Barker 2006). There are approximately 80 genera in the family, and these can range from ground cover trailing species through to very large rainforest trees (Rebelo 2001; Weston and Barker 2006). Of the 80 genera only a relatively small number are grown for cut flower production, mainly species in Protea, Leucodendron, Leucospermum, Banksia, Telopea, and Grevillea. There are other species in some of the other genera that are cultivated, but these are rare and commercially insignificant despite being of interest from a plant pathological perspective. All of the species grown are essentially woody shrubs or small trees, perennial, and clonally propagated through cuttings or, occasionally, tissue culture (Crous et al. 2013). Due to the nature of the species, orchards are long lived, and hence ongoing disease management is extremely important and in some circumstances quite problematic (Greenhalgh 1981; Crous et al. 2013). Where data are available, specific information on the management of diseases of Proteaceae is presented in the following sections. General management strategies may be found in the introductory chapters on integrated disease management.

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Fungal and Fungus-Like Diseases

2.1

Leaf Spot Diseases (A Broad Range of Fungal Species)

2.1.1 Geographic Occurrence and Impact These diseases have been reported in all regions where species of Proteaceae are grown and cultivated and in all regions where these plants grow naturally (Crous et al. 2000, 2008; Taylor et al. 2001a, b). They are a major constraint to cultivation of

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Fig. 1 Phyllosticta leaf spot on Telopea sp. (left); Mycosphaerella leaf spot on Protea cynaroides (right) (Photos courtesy BA Summerell)

Proteaceae as they both reduce growth and reduce the esthetic quality of the cut flowers.

2.1.2 Symptoms/Signs Early symptoms on leaves consist of small necrotic spots which expand and coalesce potentially resulting in large areas of the leaf turning necrotic and dead. In some cases the fruiting structures of the fungus may be evident in the leaf spots as small black pinhead-sized dots. There are very subtle differences between some of the leaf spots caused by different fungal species, but generally the symptoms overlap considerably and generally the control approaches are consistent (Fig. 1). 2.1.3 Biology and Epidemiology The fungal species of major concern are Teratosphaeria, Batcheleromyces, Brunneosphaerella, Coleroa, Pestalotiopsis (Swart et al. 1999b), Phyllosticta, and Ramularia. Many of these species were formerly referred to under the broad heading of Mycosphaerella spp., and the older literature will refer to these pathogens by this name (Taylor and Crous 2000; Crous et al. 2009). A comprehensive account of the fungal species causing these diseases is provided in Crous et al. (2013). Generally all of these fungi are favored by warm, humid conditions that allow sporulation and subsequent infection, although some species of Teratosphaeria appear to require a period of lower temperature (c. 15  C/59  F) in order for the infection process to occur. Infection propagules are a mixture of ascospores and conidia, with the latter often produced from pycnidia. It is presumed that persistence of the fungi occurs via ascocarps and mycelia in host tissue and trash in plantations. 2.1.4 Management • Cultural practices – Reduction of relative humidity and the duration of plant tissue wetness are thought to assist in reducing opportunities for infection, and consequently drip irrigation is likely to be beneficial when used.

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• Sanitation – Removal of senescing and dead plant tissue that provide a ready substrate for persistence of these fungi is likely to be important for the management of these diseases. • Fungicides – Standard fungicide applications for the control of leaf spots in other florists’ crops are likely to be effective depending on whether the fungicide is registered for use on Proteaceous crops (for additional information, refer to Chap. 6, Fungicides and Biopesticides for Florists’ Crops Disease Management).

2.2

Botryosphaeria Leaf Blight and Stem Canker (Neofusicoccum sp.)

2.2.1 Geographic Occurrence and Impact Cankers and dieback of Proteaceae are among the most important and persistent diseases that afflict all of the cultivated Proteaceae around the world. Affected plants lose productivity, and the speed of the dieback can vary considerably depending on environmental conditions. Diagnosis is difficult because non-symptomatic plants can harbor the pathogen for some time without any obvious signs (Swart et al. 2000). 2.2.2 Symptoms/Signs Typically symptoms include cankers on limbs and branches of the bushes with a resultant dieback of the affected part of the plant. In some situations the whole plant may be affected and will die. There are indications, based on observations on Proteaceae but also from extrapolation from other perennial species, that pruning wounds and other damage facilitates entry into the plant. 2.2.3 Biology and Epidemiology There are a number of species that have been associated with these diseases on a range of Proteaceae, and the taxonomy of this group is complex. Many are referred to as Neofusicoccum species, but Diplodia seriata and Lasiodiplodia theobromae are also included in this group of pathogens on members of this plant family (Denman et al. 2003; Crous et al. 2006). All are considered wound pathogens that infect via pruning wounds, tears, and cracks and insect injury (there are a number of stem borers that affect Proteaceae species). Sporulation occurs in late spring and early summer depending on rainfall/irrigation frequency, and rain splash appears to be the most frequent method of dispersal. After infection the fungus may colonize parts of the plant and remains apparently dormant for some time causing no obvious symptoms. Stress events like drought stress or frosts appear to allow the fungus to begin colonizing the plant and eventually increase biomass and cause the characteristic symptoms. 2.2.4 Management • Cultural practices – Cultivation practices that reduce the opportunities for the plant to be wounded are highly recommended. Given that the nature of the cultivation and harvesting requires cutting and wounding perennial woody material, this can be especially problematic in the Proteaceae. This includes ensuring

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that crop and plantation architecture are designed to allow rapid drying of cut surfaces and training harvesters to ensure that cuts are clean, cutting implements are clean, and stem bases and trunks are not damaged in harvest, weed control, and other management routines. Practices that minimize the plants from being stressed are also critical. This can include irrigation frequency and management and use of mulches to prevent soil drying out and insect management. • Sanitation – Removal of senescing and dead plant tissue that provide a ready substrate for these fungi to persist is important for the management of the disease. It is also important to ensure that dead and dying nonproductive plants are removed quickly and hygienically. Dead material, especially woody material, needs to be removed from the plantation or burnt to ensure there is no new inoculum available for reinfection. This may include woody material from other unrelated plant material that can potentially harbor the pathogen. • Fungicides – Effective management of Botryosphaeria was reported using a range of fungicides (benomyl, captan, captab, mancozeb) (Benic and Knox- Davies 1983; von Broembsen and van der Merwe 1990). In South African field trials using Protea magnifica, the best control was achieved by treatments of prochloraz alternated with mancozeb (Denman et al. 2004). The occurrence of cankers was also significantly reduced by applications of bitertanol and fenarimol. However, the labeling of these materials for use on Proteaceae must be determined. • Resistance – There is no information on the relative tolerance or resistance of the Proteaceae to these pathogens.

2.3

Botrytis Seedling and Flower-Head Blight (Botrytis cinerea Pers.: Fr.)

2.3.1 Geographic Occurrence and Impact Botrytis blight is a ubiquitous problem for the production and postharvest shelf life of florists’ crops. The pathogen occasionally attacks seedlings and the flowers of various Proteaceae when the conditions are humid and warm enough to favor both infection and proliferation of the fungus. 2.3.2 Symptoms/Signs Botrytis blight, also known as gray mold, is characterized by prolific gray, fuzzy fungal growth consisting of masses of hyphae, conidiophores, and conidia. On seedlings the fungus can cause a damping off and attack cuttings during propagation. On flowers the infection will initially begin on the styles and perianths and eventually attack the whole bloom (Fig. 2). Senescing and abscising flowers and damaged tissue are very susceptible to B. cinerea, and the fungus commonly uses such infection sites to invade healthy tissue; the nectar of Proteaceae flowers is produced in significant quantities and can provide a substrate to assist in proliferation of the fungus. New shoots are also susceptible to infection, and the symptoms can be evident as a shepherd’s crook appearance which will eventually die-off. In extreme circumstance the whole bloom can be covered by a mass of grey Botrytis spores.

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Fig. 2 Botrytis blight on Telopea sp. (Photo courtesy BA Summerell)

2.3.3 Biology and Epidemiology The fungus can survive as a pathogen or saprophyte in a wide range of plant hosts including many florists’ crops (Daughtrey et al. 2000). Mycelium of the fungus was found to survive through the summer in plant debris in Israel (Yunis and Elad 1989), and some isolates produce small, black sclerotia (Ellis and Waller 1974). Conidia of B. cinerea are easily spread by air currents and water splash. All above-ground plant parts (stems, leaves, and flowers) are susceptible to B. cinerea, which is favored by cool, moist conditions. Conidia of the pathogen must remain wet for 5–8 h for infection by B. cinerea to take place (Jewett and Jarvis 2001). 2.3.4 Management Components of the integrated management of Botrytis blight include: • Cultural practices – Reduction of RH and the duration of plant tissue wetness are critical for Botrytis blight control (Daughtrey et al. 2000) and avoidance of overhead irrigation. • Sanitation – Removal of senescing and dead plant tissue that provide a ready substrate for Botrytis sporulation is important for the management of the disease. • Fungicides – Standard fungicide applications for the control of Botrytis in other florists’ crops are likely to be effective depending on whether the fungicide is registered for use on Proteaceous crops.

2.4

Colletotrichum Anthracnose Disease Complex/Colletotrichum Tip Dieback (Colletotrichum spp.)

2.4.1 Geographic Occurrence and Impact The disease has been reported to occur in all areas wherever members of the Proteaceae are grown. It is more problematic in nursery situations, particularly in

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young plants, where both the environmental conditions favor the fungus, and the tissue of the younger plants are more susceptible to infection.

2.4.2 Symptoms/Signs Colletotrichum tip dieback is an important disease of Proteas in South Africa and Australia but only occurs in those regions where the environmental conditions favor the pathogen. Anthracnose on Proteas is typical of these types of diseases with symptoms that include leaf necrosis, sunken lesions, and cankers on stems and petioles which in some circumstances can girdle the stem. This can cause death above that point and may abort the flower. In extreme circumstances it may extend into the whole plant causing death of the plant. It is not uncommon for the disease to be quite prevalent in nursery situations causing damping off and seedling blight syndromes, and this can be difficult to control. 2.4.3 Biology and Epidemiology The specific identity of the species of Colletotrichum that are causing these diseases is poorly understood. In South Africa it was shown that C. acutatum and C. gloeosporioides isolates originating from Protea were the primary pathogens associated with Colletotrichum leaf necrosis, while C. acutatum was the primary cause of anthracnose and stem necrosis on the cvs. tested (Lubbe et al. 2006). It is likely that given both the geographic spread and the range of host plants, a complex of species of Colletotrichum is responsible for the disease. Conidia, produced in diseased leaves, flowers, and stems, appear to be spread by rain or irrigation water especially in nursery situations. This occurs very frequently when extended periods of warm, humid, and wet weather occur. It is likely that the fungus persists on infected plant material as acervuli and reinfection is initiated from conidia produced there. 2.4.4 Management Components of the integrated management of Colletotrichum diseases in Proteaceae include: • Cultural practices – Reduction of RH and the duration of plant tissue wetness are likely to reduce disease severity (Lubbe et al. 2006). • Sanitation – Removal of infected, senescing, and dead plant tissue that provides a ready substrate for sporulation is critical for effective management of the disease. • Fungicides – There were reports of effective management of the disease using a range of fungicides (e.g., benomyl, captan, mancozeb), but their labeling may be problematic.

2.5

Elsinoe Scab Disease (Elsinoe spp.)

2.5.1 Geographic Occurrence and Impact Present on Proteaceae crops in South Africa, the USA (Hawaii), and Australia (Fosberg 1993). In some species, especially Leucospermum and Leucodendron,

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Fig. 3 Elsinoe scab on Protea magnifica x obtusifolia (Photo courtesy BA Summerell)

the disease can cause significant disease and crop losses. The scabby lesions can also reduce the esthetics and visual appearance on a range of species thus affecting the value per flower.

2.5.2 Symptoms/Signs Symptoms begin as small black specks on leaves, flowers, and the stem which expand to produce the distinctive scabby lesions. The scabs are more prevalent on leaves but can also be problematic on flowers. The lesions can coalesce and expand to disfigure large areas of the leaves. In some species there can be distortion of the stem and splitting of the stem may occur (Fig. 3). 2.5.3 Biology and Epidemiology There are a number of Elsinoe species involved in this disease, the most important being E. leucospermi, E. proteae, and E. protearum (Swart et al. 2001). Relatively little is known about the biology of these organisms, but it appears that infection only occurs on young actively growing tissue. The infection process is favored by rain or irrigation and moderate temperatures and is more prevalent on plants grown under shade cloth, presumably because leaf moisture persists longer. Microsclerotia may be formed by some species and may provide inoculum for the fungus to persist, but it is likely that small scabs on existing stems provide opportunities for the fungus to persist between favorable environmental conditions. 2.5.4 Management • Fungicides – Fungicides (including prochloraz, captafol, and mancozeb) were effective for control of scab by spraying young active growth when conditions were favorable for infection in South Africa. However, availability of these fungicides may be limited, and alternatives have not been tested. Certain

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fungicides containing copper are registered for control of citrus scab caused by Elsinoe fawcettii in Australia, but their labeling for Proteaceae must be determined. • Resistance – Cultivars and seedlings differ in susceptibility to the scab pathogens and even within species, there can be substantial variation in susceptibility to the disease amongst cultivars. • Sanitation – Sanitation pruning is essential for control of this disease. Pruning should be focused on late summer when lesions are most evident and infection minimal; pruning should aim to remove stems and other material containing scab lesions.

2.6

Pyrenophora Blight (Pyrenophora spp.)

2.6.1 Geographic Occurrence and Impact Present on Proteaceous crops in South Africa, the USA (Hawaii), and Australia and also reported in New Zealand, Canary Islands, and Madeira. Mostly associated with Leucospermum and Protea but also recorded on Leucodendron. 2.6.2 Symptoms/Signs The disease results in a severe blight of new leaves, shoots, and flowers as they are produced in growth flushes. Initial lesions on leaves are yellow with a red margin, and these expand and cause the blight. The lesions become necrotic with age (Fig. 4). 2.6.3 Biology and Epidemiology The disease is most commonly associated with Pyrenophora biseptata, with P. leucospermi also reported. Relatively little is known about the biology of these organisms, but it appears that infection only occurs on young actively growing tissue and that like other Pyrenophora pathogens is favored by moist conditions. 2.6.4 Management • Fungicides – Information on fungicides for control of this disease is not available. • Resistance – Leucospermum species are most susceptible, and there appears to be some variability in sensitivity of different species and cultivars. • Sanitation – Sanitation pruning is essential for control of this disease. Pruning should aim to remove stems and other material containing scab lesions.

2.7

Phytophthora Root and Collar Rot and Sudden Death Disease (Phytophthora spp.)

2.7.1 Geographic Occurrence and Impact The disease is caused by a number of Phytophthora spp. but most especially P. cinnamomi and P. nicotianae but also by P. citricola, P. citrophthora, and P. cryptogea in South Africa and P. multivora in Australia (Burgess et al. 2017).

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Fig. 4 Pyrenophora blight of Leucospermum sp. (Photo courtesy BA Summerell)

Of these species P. cinnamomi is the most prevalent and the most destructive and is a pathogen of Proteaceae wherever they are cultivated, and indeed in many of the areas where they occur naturally.

2.7.2 Symptoms/Signs Symptoms include brown root rot, stunting, yellowing and purpling, and wilting of foliage – it is not uncommon for many species of Proteaceae to die from the impact of the disease – hence the description “sudden death disease” is provided. Since these symptoms overlap with those caused by other soilborne pathogens, a correct diagnosis is necessary for management. The symptoms will reflect the nature of the disease – root rot symptoms include necrosis and rot of the root system and depending on the susceptibility of the species will be restricted to the extremities of the root system or will advance through the root system eventually killing the plant. Infection at the collar will cause a collar rot that may eventually girdle the stem, also resulting in death of the plant. Infection of seedlings can result in damping off, often killing the young plants (Figs. 5 and 6).

2.7.3 Biology and Epidemiology Although the genus Phytophthora is fungus-like, it belongs to a different kingdom, the Chromalveolata. Phytophthora spp. are readily and rapidly disseminated as motile zoospores through irrigation, surface water, and rain; diseases incited by Phytophthora are favored by saturated soil. Long-term survival is by chlamydospores and sexually produced oospores. Phytophthora, depending on the species, has a broad host range. Infection can occur in any part of the root system and occurs via zoospores that encyst on the root system and penetrate the roots.

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Fig. 5 Phytophthora root rot and blight on Banksia grandis seedling (left); yellowing of leaves of Telopea sp. caused by Phytophthora root rot (right) (Photos courtesy BA Summerell)

Fig. 6 Purpling of the foliage of Protea sp. caused by Phytophthora root and collar rot (left); yellowing and dieback of the foliage and unthrifty growth of Protea sp. resulting from root and collar rot caused by P. cinnamomi (right) (Photos courtesy BA Summerell)

2.7.4 Management Refer to the introductory chapters on integrated disease management. • Cultural practices – The use pathogen-free plant propagative material and media. In the field, plant in well-drained soil. Carefully monitor irrigation practices to avoid saturated soil and refrain from using flood irrigation which may spread propagules of the pathogen. Control weeds to increase air circulation and drying

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of plant surfaces especially the collar region. Control insects which may create infection courts for the fungus. • Sanitation – Promptly remove and discard/burn infected plants. Avoid using untreated surface water which may contain inocula of Phytophthora. Disinfest tools, equipment, and shoes before traveling to other Protea production areas. Refer to Chap. 8, Sanitation in Florists’ Crops Disease Management, for information on disinfestation practices. • Fungicides – Potassium phosphonate (phosphite)-based fungicides are widely used to reduce the impact of this disease on plantations. These can be applied via stem injection or via foliar applications as either a preventative or a curative. • Resistance – In research conducted in South Africa with potted Leucospermum, the cv. Spider was tolerant to root and crown rot caused by P cinnamomi even when tested against a highly virulent isolate under disease-conducive conditions (Swart and Denman 2000). The researchers suggested the integration of tolerance, when known, with fungicide use.

2.8

Pythium Root Rot (Pythium spp.)

2.8.1 Geographic Occurrence and Impact Pythium vexans was reported to cause disease in Proteaceae in the USA (California (Raabe and Hürlimann 1972), and P. debaryanum and P. spinosum were isolated from these plants in New Zealand (Robertson 1973). There are also a number of reports of Pythium spp. isolated from the root systems from a number of different species of Proteaceae in both South Africa and Australia, but little detailed information is available on the nature of the specific species of Pythium or the extent of disease caused. It is presumed that a number of different species are capable of causing damping off and root disease in nursery situations. 2.8.2 Symptoms/Signs The root systems become discolored and rotted, and infected seedlings rapidly wilt and may cause pre- and postemergence damping-off. 2.8.3 Biology and Epidemiology The genus Pythium is closely related to Phytophthora and is often recovered in association with species of Phytophthora. It is disseminated as zoospores presumably through irrigation, surface water, and rain, and long-term survival is enabled by chlamydospores and sexually produced oospores that may persist in potting mixes and soil. 2.8.4 Management Following the management practices indicated above for Phytophthora should help to control Pythium.

Diseases of Proteaceae

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Armillaria Root Rot (Armillaria sp.)

2.9.1 Geographic Occurrence and Impact Armillaria root rot has been recorded on members of the Proteaceae in South Africa, Zimbabwe, Australia, New Zealand, and the USA. As the species of Proteaceae that are grown are woody perennials, the disease has time to develop and eventually causes significant problems in individual specimens. The disease is only sporadically considered an issue in cut flower plantations. 2.9.2 Symptoms/Signs Symptoms typically are dieback and death of part or all of the plant (Fig. 7). Diagnostic symptoms include splitting and cracking of the stem at ground level; white fungal growth may be obvious if the bark at ground level is lifted off. In severe cases the stem bases may become soft and decayed. Fruiting bodies of the fungus may appear on rooted wood. 2.9.3 Biology and Epidemiology Different species of Armillaria cause disease in different parts of the world. In South Africa A. gallica and A. mellea have been reported on Leucodendron and Protea species and appear to be incursions from the Northern Hemisphere. In Australia a broad range of Proteaceae species, both in natural ecosystems and cultivated circumstances, have been infected by the indigenous fungus, Armillaria luteobubalina. The fungus persists as mycelium on root systems and is the key inoculum for infecting plants; the fungus can persist on those root systems for extended period depending on the size of the piece of infected root or crown material. Infection occurs where an infected (or infested) root system comes into

Fig. 7 Dieback on Protea cynaroides caused by Armillaria (left); signs of Armillaria on wood: white fungal mycelium and black rhizomorphs (arrow) (center); fruiting bodies (mushrooms) of Armillaria luteobubalina on rotted roots and crown (right) (Photos courtesy BA Summerell)

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contact with uninfected root material. In the case of A. mellea and A. gallica, rhizomorphs may extend through the soil to infect root material; this does not happen with A. luteobubalina. The mycelium of the fungus colonizes the root system eventually killing that part of the roots and may eventually colonize and girdle the stem or crown of the plant thus killing the plant. The basidiomata are produced annually, usually in late autumn, but, while diagnostic, play a relatively little role in initiating new infections, but under some circumstances the basidiospores can initiate a new infection. Disease spread occurs via movement of the mycelium along the root system moving from plant to plant, through transplant or planting of infected material, or via the use of infested woody material as mulch.

2.9.4 Management Once infected it is difficult to control the fungus in the plant. However in some circumstances, it is possible to expose infected areas by removing soil, and this allows the area to dry suppressing the growth of the fungus. • Sanitation – Control depends on the removal of infected material from an area. This can be achieved by removal of as much of the root system as possible which will effectively result in much of the fungal material being removed thus stopping the infection cycle.

2.10

Rhizoctonia Root Rot (Rhizoctonia sp.)

2.10.1 Geographic Occurrence and Impact Rhizoctonia root, crown, and stem rot is a poorly understood pathogen on Proteaceae but is believed to be common, particularly in a nursery situation. There is effectively no information of the nature of which species of Rhizoctonia or which of the anastomosis groups may be involved; however it is presumed that Rhizoctonia solani is involved. The disease has been reported in most regions where Proteaceae species are cultivated and across the broad range of Proteaceae species. 2.10.2 Symptoms/Signs The most often reported symptom is seedling rot and wirestem symptoms that may result in the death of the seedling. The pathogen may also cause pre- and postemergence damping-off. 2.10.3 Biology and Epidemiology Rhizoctonia solani has a broad host range that appears to include most of the Proteaceae that are cultivated. The disease appears to be more active and prevalent at warm temperatures, and the fungus effectively colonizes and survives as hyphae and sclerotia in plant debris, soil, and other growing media.

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2.10.4 Management There has been little research on control measures for this disease, but it is presumed that standard control measures for the disease in other crops would be effective.

2.11

Fusarium Wilt (Fusarium oxysporum Schlect.)

2.11.1 Geographic Occurrence and Impact Fusarium wilt has been recorded on a number of South African species in the Proteaceae, most notably on Protea, Leucospermum, and Leucodendron and has been reported as the second most important root disease after Phytophthora root rot in South Africa (Swart et al. 1999a). The disease has also been reported in Colombia (Martínez Granja et al. 2014), and there are no substantial reports of Australian species of Proteaceae being affected by Fusarium wilt, although F. oxysporum may often be isolated from the roots of a range of Australian Proteaceae (Summerell et al. 2011; Summerell unpublished data).

2.11.2 Symptoms/Signs Plants that are infected will display early symptoms of leaf blackening which may coalesce resulting in leaf and shoot death. Most typically new shoot growth will wilt. Necrosis can be observed in the roots and will extend into the phloem and xylem with vascular discoloration evident (Fig. 8).

2.11.3 Biology and Epidemiology The etiology of the disease is typical of Fusarium wilt. Unassisted spread is slow but can easily be moved via movement of soil or in irrigation water. The pathogen can persist in soil as chlamydospores and in root tissue for extended periods of time. Typically infection occurs via roots and moves through the vascular system blocking the movement of water through the plant.

2.11.4 Management • Sanitation – The use pathogen-free plant propagative material and media. Disinfest tools, work surfaces and containers before reuse as Fusarium can survive for long periods even on such inert surfaces. Promptly remove and discard/burn infected plants. • Soil fumigation – Soil fumigation is reported to be effective in reducing the amount of inoculum in the soil, but little work has been done in determining effectiveness in disease control. • Fungicides – No fungicides are available for control of this disease.

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Fig. 8 Fusarium wilt. Note brown discolored streak on the stem of the infected plant on the left (arrow) (Photo courtesy of Swart et al. 1999a)

Fig. 9 Ilyonectria black foot rot: decline of newly planted cuttings in a field bed (left); root and black foot rot with vascular discoloration in the stem (right) (Photos courtesy of Lombard et al. 2013)

2.12

Ilyonectria Black Foot Rot [Ilyonectria spp. (formerly Cylindrocarpon spp.)]

2.12.1 Geographic Occurrence and Impact The disease has been reported in South Africa and Australia on Protea, Leucospermum, and Telopea (Summerell et al. 1990). A range of species of Ilyonectria have been reported; this genus was previously better known as Cylindrocarpon. Similar diseases are common in a range of perennial woody species, especially in grapevines. 2.12.2 Symptoms/Signs Disease symptoms include root, collar, stem, and crown rot, usually associated with blackening of these parts that slowly spreads up the stem of the infected plant (Fig. 9).

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2.12.3 Biology and Epidemiology This pathogen is predominantly a problem of seedlings, particularly in nurseries and propagation facilities. If the plant is infected in this situation and then planted out, the disease can persist in older plants in plantations. In these situations the symptoms may not be obvious until an infected plant is stressed in some way and then symptoms can include death of the plant. 2.12.4 Management Management of this disease has not received much attention, but it can be expected that nursery hygiene and ensuring that diseased plants are not used for establishing plantations is essential. 2.12.5 Additional Fungal Diseases The following fungal diseases of Proteaceae have also been reported: – – – – – – – –

Calonectria root rot (Calonectria pauciramosa) (South Africa, Crous 2002) Ceratocystis canker (Ceratocystis albifundis) (South Africa, Crous et al. 2000) Diaporthe stem canker (Diaporthe saccarata) (Mostert et al. 2001) Epicoccum brown stem canker (Epicoccum sorghi) (South Africa, Crous et al. 2013). Phloesporella canker (Phloesporella protearum) (Zimbabwe, Taylor and Crous 2000) Charcoal rot (Macrophomina phaseolina) (South Africa, Canary Islands, Espino et al. 2011, Knox-Davies et al. 1987) Rosellinia root rot (Rosellinia necatrix) (Canary Islands, Madeira Island, Espino et al. 2011, Moura and Rodrigues 2001)

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Bacterial and Phytoplasma Diseases

3.1

Pseudomonas Leaf Spot [Pseudomonas syringae Pathovar “Pv. Proteae”]

3.1.1 Geographic Occurrence and Impact The disease has been reported in Australia and the UK (Wimalajeewa et al. 1983) on Protea cynaroides. 3.1.2 Symptoms/Signs The pathogen can infect seedlings as well as older plants, and this can retard growth and reduce the esthetic quality of the cut flowers. The spots start as water-soaked lesions that expand, turn necrotic, and can be surrounded by a bright red halo. The entire leaf surface may become affected and wither and die.

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3.1.3 Biology and Epidemiology Little is known of the disease, but it is presumed that spread of the pathogen occurs as a result of rain splash and movement of infected plant material. Disease development is, like many bacterial diseases, dependent on wet and humid conditions. 3.1.4 Management • Fungicides – Copper sprays are reported to protect foliage against infection.

3.2

Phytoplasma Diseases: Witch’s Brooms

3.2.1 Geographic Occurrence and Impact Diseases of Proteaceae caused by phytoplasmas have been reported to occur in South Africa (Wieczorek and Wright 2003) and observed in Australia (Summerell unpublished data). The disease has been reported in species of Aulax, Leucodendron, Leucospermum, Protea, Serruria, and Vexatorella in South Africa and observed in species of Banksia, Grevillea, and Hakea in Australia. 3.2.2 Symptoms/Signs Symptoms caused by phytoplasmas may include yellowing of foliage, stunting, and proliferation of shoots (“witch’s brooms”), the abnormal development of floral parts into leafy structures (phyllody), and of green coloration in flowers (virescence). 3.2.3 Biology and Epidemiology Phytoplasmas are prokaryotes that lack cell walls and are obligate plant parasites vectored by, in this case, a range of mites. Species in the genera Aceria, Proctolaelaps, and Oxycarenus were reported on Proteas in South Africa, but little is known as to whether these are the same species on other South African Proteaceae or associated with species in Australia. 3.2.4 Management Management of phytoplasma diseases, like those caused by viruses, is based on monitoring and reduction of arthropod vector populations through chemical or biological means, vector exclusion by physical barriers such as fine mesh screening, vector disorientation and repellence in field-grown crops through the use of reflective mulches, and elimination of, and avoidance, of growing near, alternate hosts especially established, infected crops.

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Viral Diseases

No viral diseases of Proteaceae have been reported.

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References Barker NP, Weston PH, Rutschmann F, Sauguet H (2007) Molecular dating of the ‘Gondwanan’ plant family Proteaceae is only partially congruent with the timing of the break-up of Gondwana. J Biogeogr 34:2012–2027 Benic LM, Knox-Davies PS (1983) Scab of Leucospermum cordifolium and other Proteaceae, caused by an Elsinoë sp. Phytophylactica 15:95–107 Burgess TI, White D, McDougall KM, Garnasc J, Dunstan WA, Catala S, Carnegie AJ, Worboys S, Cahill D, Vettraino A-M, Stukely MJC, Liew ECY, Paap T, BoseC T, MiglioriniC D, Williams B, Brigg F, Crane C, Rudman T, GESJ H (2017) Distribution and diversity of Phytophthora across Australia. Pac Conserv Biol 23:1–13 Crous PW (2002) Taxonomy and pathology of Cylindrocladium (Calonectria) and allied genera. APS Press, Saint Paul Crous PW, Summerell BA, Taylor JE, Bullock S (2000) Fungi occurring on Proteaceae in Australia: selected foliicolous species. Australas Plant Pathol 29:267–278 Crous PW, Slippers B, Wingfield MJ, Rheeder J, Marasas WFO, Philips AJL, Alves A, Burgess T, Barber P, Groenewald JZ (2006) Phylogenetic lineages in the Botryosphaeriaceae. Stud Mycol 55:235–253 Crous PW, Summerell BA, Mostert L, Groenewald JZ (2008) Host specificity and speciation of Mycosphaerella and Teratosphaeria species associated with leaf spots of Proteaceae. Persoonia 20:59–86 Crous PW, Summerell BA, Carnegie AJ, Wingfield MJ, Hunter GC, Burgess TI, Andjic V, Barber PA, Groenewald JZ (2009) Unravelling Mycosphaerella: do you believe in genera? Persoonia 23:99–118 Crous PW, Denman S, Taylor JE, Swart L, Bezuidenhout C, Hoffman L, Palm ME,Groenewald JZ (2013) Cultivation and diseases of Proteaceae: Leucadendron, Leucospermum and Protea, 2nd edn. CBS Biodiversity Series, Vol 2. CBS-KNAW Fungal Biodiversity Centre, Utrecht, pp 1–360 Daughtrey ML, Wick RL, Peterson JL (2000) Botrytis blight of flowering potted plants. Plant Health Progress. doi:10.1094/PHP-2000-0605-01-HM Denman S, Crous PW, Groenewald JG, Slippers B, Wingfield BD, Wingfield MJ (2003) Circumscription of Botryosphaeria species associated with Proteaceae based on morphology and DNA sequence data. Mycologia 95:294–307 Denman S, Crous PW, Sadie A, Wingfield MJ (2004) Evaluation of fungicides for the control of Botryosphaeria protearum on Protea magnifica in the Western Cape Province of South Africa. Australas Plant Pathol 33:97–102 Ellis MB, Waller JM (1974) Sclerotinia fuckeliana. CMI descriptions of pathogenic fungi and bacteria no. 431. CAB International, Wallingford Espino AI, Guzman MN, Carrascosa F (2011) Enfermedades fúngicas de proteas en Canarias. Granja. Revista Agropecuaria, Cabildo de Gran Canaria No. 18. http://anuariosatlanticos. casadecolon.com/index.php/GRANJA/article/view/9826 Forsberg L (1993) Protea diseases and their control. Queensland Government, Department of Primary Industries, Brisbane Greenhalgh FC (1981) Diseases of Proteaceous plants. In: Mathews P (ed) The growing and marketing of proteas: 30–39. Report of the first international conference of protea growers, Melbourne, 4–8 October Jewett T, Jarvis W (2001) Management of the greenhouse microclimate in relation to disease control: a review. Agronomie 21:351–366 Knox-Davies PS, van Wyk PS, Marasas WFO (1987) Diseases of Protea, Leucospermum and Leucadendron recorded in South Africa. Phytophylactica 19:327–337

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Lombard L, Bezuidenhout CM, Crous PW (2013) Ilyonectria black foot rot associated with Proteaceae. Australas Plant Pathol 42:337–349. doi:10.1007/s13313-012-0188-5 Lubbe CM, Denman S, Lamprecht SC, Crous PW (2006) Pathogenicity of Colletotrichum species to Protea cultivars. Australas Plant Pathol 35:37–41 Martínez Granja E, Reyes Benitez S, Sanjuanello D. (2014) Effect of antagonists and plant extracts in the control of Protea Wilt (F. oxysporum). Am J Plant Sci 05(21):9 pp. Article ID: 50856. doi:10.4236/ajps.2014.521336 Mostert L, Kang JC, Crous PW, Denman S (2001) Phomopsis saccharata sp. nov., causing a canker and die-back disease of Protea repens in South Africa. Sydowia 53:227–235 Moura MF, Rodrigues PF (2001) Fungal diseases on proteas identified in Madeira Island. Acta Hortic 545:265–268. doi:10.17660/ActaHortic.2001.545.34 Raabe RD, Hürlimann JH (1972) Control of Pythium root rot in carnations. Calif Agric 26:4–5 Robertson GI (1973) Occurrence of Pythium spp. in New Zealand soils, sands, pumices, and peat, and on roots of container-grown plants. N Z J Agric Res 16:357–365 Rebelo T (2001) SASOL proteas: a field guide to the proteas of southern Africa. Fernwood Press, Vlaeberg Summerell BA, Burgess LW, Nixons PG (1990) Crown and stem canker of waratah caused by Cylindrocarpon destructans. Australas Plant Pathol 19:13–15. doi:10.1071/APP9900013 Summerell BA, Leslie JF, Liew ECY, Laurence MH, Bullock S, Petrovic T, Bentley AR, Howard CG, Peterson SA, Walsh JL, Burgess LW (2011) Fusarium species associated with plants in Australia. Fungal Divers 46:1–27 Swart L, Denman S (2000) Chemical control of Phytophthora cinnamomi in potted Leucospermum plants. Australas Plant Pathol 29:230–239 Swart L, Denman S, Lamprecht SC, Crouse PW (1999a) Fusarium wilt: a new disease of cultivated Protea in southern Africa. Australas Plant Pathol 28:156–161 Swart L, Taylor JE, Crous PW, Percival K (1999b) Pestalotiopsis leaf spot disease of Proteaceae in Zimbabwe. S Afr J Bot 65:239–242 Swart L, Crous PW, Petrini O, Taylor JE (2000) Fungal endophytes of Proteaceae, with particular emphasis on Botryosphaeria proteae. Mycoscience 41:123–127 Swart L, Crous PW, Kang J-C, Mchau GRA, Pascoe IA, Palm ME (2001) Differentiation of species of Elsinoë associated with scab disease of Proteaceae based on morphology, symptomatology, and ITS sequence phylogeny. Mycologia 93:365–379 Taylor JE, Crous PW (2000) Fungi occurring on Proteaceae. New anamorphs for Teratosphaeria, Mycosphaerella and Lembosia, and other fungi associated with leaf spots and cankers of Proteaceous hosts. Mycol Res 104:618–636 Taylor JE, Crous PW, Palm ME (2001a) Foliar and stem fungal pathogens of Proteaceae in Hawaii. Mycotaxon 78:449–490 Taylor JE, Crous PW, Swart L (2001b) Foliicolous and caulicolous fungi associated with Proteaceae cultivated in California. Mycotaxon 78:75–103 von Broembsen SL, van der Merwe JA (1990) Canker and die-back of cut-flower proteas caused by Botryosphaeria dothidea: epidemiology and control. Acta Hortic 264:133 Weston PH, Barker NP (2006) A new suprageneric classification of the Proteaceae, with an annotated checklist of genera. Telopea 11:314–344 Wieczorek AM, Wright MG (2003) PCR detection of phytoplasma from witches’ broom disease on Protea spp. (Proteaceae) and associated arthropods. Acta Hortic 602:161–166 Wimalajeewa DLS, Hayward AC, Greenhalgh FC (1983) A bacterial leaf spot of (King protea). Ann Appl Biol 102(2):339–344 Yunis H, Elad Y (1989) Survival of Botrytis cinerea in plant debris during summer in Israel. Phytoparasitica 17:13–21

Diseases of Rose Jay W. Pscheidt and Tatiana Gomez Rodriguez

Abstract

Roses are one of the most popular flowering plants in the world and tend to have a few common disease problems such as black spot, Botrytis blight, downy mildew, powdery mildew, and rust. There are a dozen different fungal diseases, four bacterial problems, three nematodes, and many viral diseases that occur on roses. Several are root or root crown-related diseases, but most are concerns for canes, leaves, and especially flowers. Cultural management of these diseases is effective especially when combined with genetic resistance. Chemical management of important problems is useful for favored, susceptible rose cultivars. Keywords

Rose • Black spot • Botrytis blight • Downy mildew • Powdery mildew • Rust • Diseases • Disease management • Rosarian

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Oomycete Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Armillaria Root Rot (Armillaria mellea (Vahl: Fr.) P. Kumm. 1871) . . . . . . . . . . . . . . . 2.2 Black Mold (Thielaviopsis thielavioides (Peyronel) A.E. Paulin, T.C. Harr. & McNew 2002) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Black Spot (Diplocarpon rosae F.A. Wolf 1912) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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J.W. Pscheidt (*) Department of Botany and Plant Pathology, Oregon State University, Corvallis, OR, USA e-mail: [email protected] T.G. Rodriguez GR Chia SAS, Centro Empresarial Centro Chia Oficina 304 Chia, Cundinamarca, Bogotá, Colombia e-mail: [email protected] # Springer International Publishing AG 2016 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florist's Crops Diseases, Handbook of Plant Disease Management, DOI 10.1007/978-3-319-32374-9_23-1

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Botrytis Bud and Twig Blight (Botrytis cinerea Pers.: Fr. 1794) . . . . . . . . . . . . . . . . . . . . Brand Canker (Coniothyrium wernsdorffiae Laubert 1905) . . . . . . . . . . . . . . . . . . . . . . . . . Brown Canker (Cryptosporella umbrina (Jenk.) Jenk. & Wehm. 1935) . . . . . . . . . . . . Cercospora Leaf Spot (Pseudocercospora puderi B.H. Davis ex Deighton 1976) (Passalora rosicola (Pass.) U. Braun 1995) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.8 Common Canker (Paraconiothyrium fuckelii (Sacc.) Verkley & Gruyter 2012) . . . . 2.9 Cutting Rots (Cylindrocladium scoparium Morg. 1892 and Fusarium spp.) . . . . . . . 2.10 Downy Mildew (Peronospora sparsa Berk. 1862) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.11 Myrothecium Gall (Myrothecium sp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.12 Petal Spots (Ghost Spot) (Many Fungi Possible Such as Bipolaris, Botrytis, Cercospora, and/or Cladosporium) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.13 Powdery Mildew (Podosphaera pannosa (Wallr.: Fr.) de Bary 1870) . . . . . . . . . . . . . . 2.14 Replant Disease (Many) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.15 Rust (Phragmidium mucronatum (Pers.) Schltdl. 1824 and Many Other Species.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.16 Spot Anthracnose (Elsinoe rosarum Jenk. & Bitanc. 1957) . . . . . . . . . . . . . . . . . . . . . . . . . 2.17 Verticillium Wilt (Verticillium dahliae Kleb. 1913) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.18 Miscellaneous Fungal and Oomycete Diseases of Minor Importance or Considered Rare . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Bacterial and Phytoplasma Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Bacterial Cane Blight (Pseudomonas syringae – Undetermined Pathovar) . . . . . . . . . . 3.2 Bacterial Leaf Spot and Blast (Pseudomonas syringae pv. morsprunorum ) . . . . . . . . . 3.3 Crown Gall (Agrobacterium radiobacter (Beijerinck and Van Delden 1902) Conn 1942 or Also Known as Rhizobium radiobacter (Beijerinck and Van Delden 1902) Young et al. 2001) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4 Miscellaneous Bacterial Diseases of Minor Importance or Considered Rare . . . . . . . . . 4 Viral Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Rose Leaf Curl (A Virus Is Suspected) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Rose Mosaic [Prunus necrotic ringspot virus (PNRSV), Apple mosaic virus (ApMV), Arabis mosaic virus (ArMV), and Strawberry latent ringspot virus (SLRSV)] . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Rose Rosette (Rose Rosette Virus) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4 Rose Spring Dwarf (Rose Spring Dwarf-Associated Virus) . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5 Rose Yellow Mosaic (Rose Yellow Mosaic Virus) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6 Miscellaneous Viral or Virus-Like Diseases of Minor Importance or Considered Rare . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Nematode Diseases (Meloidogyne hapla, Pratylenchus Penetrans, P. Vulnus, and Xiphinema Diversicaudatum) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Introduction

Roses are one of the most popular flowering plants in the world grown as cut flowers in greenhouses, as whole plants in nurseries or in backyards. There tend to be a few common problems in each growing situation, such as black spot, Botrytis blight, downy mildew, powdery mildew, and rust. Some problems have become troublesome during the 2010s including downy mildew and rose rosette. Major regional differences occur, such as rose replant disease, which is of concern in Europe but not recognized as a problem in North America. Powdery mildew, downy mildew, rust, and Botrytis blight are more problematic under greenhouse and tropical weather conditions found

Diseases of Rose

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Fig. 1 Hydroponic production of roses in Sabana Bogotá. Left – 8-week-old plants in rice hull substrate. Right – same plants at 4 months old (Photo courtesy of Tatiana Gomez Rodriguez)

in production houses in Central and South America. Disease management and handling increases commercial production costs and limits rose production. In Colombia, pest management has improved by switching from traditional production in the ground with drip irrigation to hydroponics. This change has resulted in better area usage, nutrition, and soil pest management, ultimately leading to better rose quality and productivity. Production takes place most commonly under plastic cover (in greenhouses) with varying degrees of environmental control. Although rose is a perennial, in this production system, it only has a useful life of 8–15 years. About 6.5 plants/m2 or 380 plants per bed are used in a double row of plastic containers with nutrient film technique (NFT) subirrigation. In the hydroponic system, beds are covered by white plastic mulch or raw rice husks to improve soil moisture and facilitate weed management. It takes approximately 750–820 degrees days (equivalent to 70–80 calendar days), depending on the cultivar of rose, from the time of pruning to the first flower crop (Fig. 1). An important consideration before implementing a disease management plan is the ultimate goal of growing roses. A backyard, non-gardener grower of a simple rose bush will have very different goals and concerns than the rosarian growing specific bushes for competitions or the nurseryman growing bare-root plants, plants in containers or cuttings. The grower’s tolerance for various diseases will differ radically among these groups. We will cover many different management tactics. Choose the tactics most applicable for your situation.

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Fungal and Oomycete Diseases

2.1

Armillaria Root Rot (Armillaria mellea (Vahl: Fr.) P. Kumm. 1871)

Geographic occurrence and impact. Worldwide, an occasional problem depending on the history of the growing area.

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Symptoms/signs. Aboveground symptoms are general and not specific. Diagnostic mycelial fans can be observed beneath the bark of the root crown of infected plants. Dig down about 15 cm (6 in.) below the soil line, and use a pocketknife to remove thin layers of bark from the root collar. Mycelial fans are thick, white layers of the fungus that adhere to the inner root bark and/or the wood beneath the bark. The fungus also makes black, shoestring-like structures called rhizomorphs, which are occasionally found within the bark and/or extending into surrounding soil. Rhizomorphs may look like roots on the outside but have an entirely different structure when cut open in cross section. Biology and epidemiology. The fungus, infects roots, killing the cambium and decaying the underlying xylem. Occasionally found on rose, this root pathogen is found in native vegetation where it occurs on the roots of many forest tree species and agronomic and ornamental hosts. The host range includes over 500 species of woody plants. The fungus spreads slowly, below ground, from root to root, and can survive on woody host roots for long periods. When infected plants are removed, infected roots that remain below ground serve as a source of inoculum for other plants placed in the same location. Infection occurs when rose roots come in direct contact with partially decayed tree roots and are colonized by mycelium. Infection can also occur when roots contact rhizomorphs that grow out from partially decayed roots and through the soil. Management • Cultural practices – Roses that are vigorous, as a result of favorable climate and sound cultural practices usually are more tolerant. Water deeply when watering is needed. Avoid surface watering, especially wetting the root crown and trunk root area. If using drip irrigation, move drip-line emitters away from the root crown and in between plants. Remove severely infected bushes and destroy them, being careful to remove small roots from the soil. Permanently removing soil from around the root crown and main root area has been effective in tree fruits grown in California and Australia and may be of benefit for managing infected bushes elsewhere (Schnabel et al. 2012).

2.2

Black Mold (Thielaviopsis thielavioides (Peyronel) A.E. Paulin, T.C. Harr. & McNew 2002)

Geographic occurrence and impact. Pacific Northwest of North America – rare, unknown how important in other regions. Fungus found in South America (Brazil), North America, Europe, Australia, Malaysia, and Indonesia. Symptoms/signs. A white, granular, fungal growth that spreads rapidly over cut and bruised areas of stock and scion. Soon the growth turns black. Biology and epidemiology. The fungus grows on the ends of “Manetti” cuttings (Rosa chinensis var. manetti) and on wounds left by disbudding, resulting in poor callus formation and, subsequently, poor root formation (Milbrath 1946). It develops under bud shields on field-budded “Manetti,” thus preventing callus formation and causing the bud to die. The fungus also may damage rose roots if the plants are in

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storage too long before planting or if plants are overwatered after planting. The chief infection source is fungal spores in the dust and debris of storage houses where the plants are handled and cuttings prepared for planting. Management • Cultural practices – Practice strict sanitation in greenhouses, storage bins, and storage houses. Wash all dust and debris from the walls and floors before disinfecting storage house and bins. Use only uncontaminated stock for planting.

2.3

Black Spot (Diplocarpon rosae F.A. Wolf 1912)

Geographic occurrence and impact. Worldwide. Repeated defoliation from this disease leads to low vigor, inferior blooms, and high susceptibility to winter injury. Very common on plants grown outdoors. Symptoms/signs. Circular black spots, frequently with fringed (diffuse or feathery) margins, on the leaf. Yellowing and defoliation are common in susceptible cultivars. Bare stems with few leaves attached near the top is a frequent symptom. In wet weather, spots may become very severe and coalesce to make large irregular spots covering a third of the leaf surface. In cases of severe infections, similar appearing lesions also form on the stems. May be confused with other fungal leaf spotting diseases (Fig. 2). Biology and epidemiology. The fungus overwinters on living or dead plant tissue that was infected in the previous growing season. Newly emerging leaves are most susceptible to this fungus. Spores produced on the old plant material are splashed on young plant tissue by rains or by watering. After conidia land on a leaf, it takes at least 9 h of leaf wetness for the spore to infect. Once the fungus is established in plant tissues, fungal fruiting bodies (acervuli) form in the spots in 11–30 days. A new crop of spores is produced and spreads to healthy portions of the plant. The cycle of infection is ready to begin again within 10–18 days. Rainy periods in any season encourage black spot, as long as the temperatures are between 10  C and 26  C (50–80  F). Leaves infected with black spot produce the plant hormone ethylene. In leaves, high ethylene content leads to leaf drop. As a result, rose plants infected with black spot are defoliated early and look bare. Leaves may drop after only one or two lesions. Dropped leaves around the base of the plant then serve as a reservoir of fungal spores, re-infecting the plant during subsequent rains or irrigation. The natural genetic variability of the fungus means roses found resistant in one region may be susceptible in another region due to the presence of different fungal strains. Also, resistant roses may become susceptible after a few years due to changes in the local fungal population. Management • Cultural practices – Plant cultivars known to be resistant in your area. Note that the cultivar Carefree Spirit was the first landscape shrub to survive for 2 years in

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Fig. 2 Black spot leaf symptoms (left) and stem lesions with severe defoliation (right) (Photo courtesy of Jay W. Pscheidt)

All-American Rose Selections (AARS) tests without any fungicide and voted a winner. Avoid dense plantings and shaded areas. Avoid overhead watering that keeps plants wet for extended periods of time. Rake up and burn all leaves in a rose planting at season’s end. Prune canes back to two buds if canes are infected. Remove and destroy diseased canes before bud break. • Fungicides – Focus applications at the beginning of the season starting at bud break. Apply frequently in wet weather and occasionally in drier periods (1- to 2-week intervals). May become very difficult to manage once the disease gets started. Almost every fungicide on the market is registered in the USA for this disease on this crop, but only one or two materials are needed at any one time. Alternate or tank-mix materials with different modes of action to prevent developing resistant fungal populations. Research in Alabama, USA, indicates that monthly dormant season applications of Fungicide Resistance Action Committee (FRAC) group 3 fungicides may help reduce or delay onset of this disease (Bowen and Roark 2001). Information developed by FRAC may be accessed at: http://www.frac.info/. Bicarbonate-based materials are not recommended for use in high disease pressure regions as one will get poor control using these products (Pscheidt and Ocamb 2016). Chlorothalonil-based products may be a problem on “Knock Out” and “Double Delight” roses resulting in damaged foliage. Rosarians may not like the plant growth regulation effect (slight stunting) that may result from the use of FRAC group 3 fungicides. Always watch for warnings on labels of various products.

Diseases of Rose

2.4

7

Botrytis Bud and Twig Blight (Botrytis cinerea Pers.: Fr. 1794)

Geographic occurrence and impact. Worldwide but mostly a problem in greenhouse cut flower production. Symptoms/signs. Infected buds may fail to open, droop, and develop smooth, slightly sunken, grayish black lesions. A grayish brown mycelial growth may develop on the entire bud and extend into the stem. Petal tips may become brown, or numerous small flecks, circular brown spots, or blister-like patches may appear on petals (Fig. 5). Infected stubs or pruning wounds will develop into cane blights or cankers. Canes may be girdled, causing stem collapse. Biology and epidemiology. The fungus is ubiquitous and survives well as a saprophyte on dead plant debris as mycelium or sclerotia. Conidia from these sources or from active infections initiate new infections. Common entry points include pruning wounds including those from normal winter pruning or from removing spent flowers (Pie and De Leeuw 1991). Buds, flowers, leaves, and stems can be infected directly when the fungus produces enzymes that degrade components of the cuticle. Healthy tissue can become infected through weakened or dead tissues that are colonized by the fungus. It may also be a secondary invader attacking plants already infected with another pathogen. A moist, humid environment is ideal for pathogen sporulation and spread. Spore dispersal is stimulated by changes in relative humidity (Hausbeck and Moorman 1996). Concentrations peak in the greenhouse during irrigating, spraying pesticides, harvest, and shipping. Often the infection is imperceptible when cutting and packing flowers; however, moist conditions that occur during transport and/or storage favor disease development. Frost and cold injury to canes and buds can also contribute to disease development. Bare-root nursery stock held in moist, cold storage conditions also can develop Botrytis blight on canes and roots. Management • Cultural practices – Timely detection of the first signs of disease is essential. Collect and destroy infected plant debris and tissues from and around the flowers, stems, and leaves. Reduce greenhouse moisture by raising the temperature, venting, and/or increasing air circulation. Space plants for adequate air movement. Use drip rather than overhead irrigation. Use calcium nitrate, gypsum, or similar calcium source topdressed or incorporated into growing media. • Fungicides – Use as a foliar spray, but focus on cultural controls. Alternate or tank-mix materials with different modes of action to prevent developing resistant fungal populations. Fungicides from the following groups are effective if resistance is not a problem: FRAC groups 1, 2, 7, 9, 12, 14, and 17. Good to mix or alternate with FRAC group M3 or M5 fungicides.

2.5

Brand Canker (Coniothyrium wernsdorffiae Laubert 1905)

Geographic occurrence and impact. Commonly found in North and South America, Europe, and Russia. Canker diseases can severely impact improperly grown roses leading to entire loss of the desired scion.

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Fig. 3 Brand canker has gray lesions with a dark border that restricts water and nutrient movement in the cane (Photos courtesy of OSU Extension Plant Pathology Slide Collection (left) and by Tatiana Gomez Rodriguez (right))

Symptoms/signs. Discolored (yellow to red) spots that gradually enlarge with the center turning light brown. In cut flower production, the cut stem turns yellow to black as the pathogen advances from the cut down to the base of the stem. Brand canker develops a reddish brown or purple margin that contrasts sharply with the normal green rose cane (Fig. 3). Small black fruiting bodies (pycnidia) develop toward the center of damaged tissue. Brand canker that develops under the snow line will at first in spring be black, but then it develops as described above after being uncovered several weeks. Cankers may girdle stems, causing stems and shoots above that point to wilt and die. The disease can be easily confused with common canker (Paraconiothyrium fuckelii) or brown canker (Cryptosporella umbrina). Biology and epidemiology. The fungus infects cane wounds caused by pruning, harvesting flowers, insects, thorn scars, or scratches (Intrama 1968). Contaminated cutting knives, scissors, or pruners are a major source of the pathogen. Cool winter temperatures and winter coverings that keep canes moist also favor brand canker. Management • Cultural practices – Avoid injuring canes. Prune canes with a disinfected, sharp knife or pruner immediately above a node. Avoid leaving long stubs above a node. Frequently clean and disinfect cutting tools (refer to chapter “▶ Sanitation in Florists’ Crops Disease Management”). Cut and destroy infected canes well below the affected area. Prevent irrigation from wetting the plant and/or use drip irrigation. Use a winter protecting mulch that does not hold much water such as sand, rock pumice, or coarse bark. Use a balanced fertilizer with moderate nitrogen.

Diseases of Rose

2.6

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Brown Canker (Cryptosporella umbrina (Jenk.) Jenk. & Wehm. 1935)

Geographic occurrence and impact. The disease is worldwide both in greenhouse and outside landscapes. Canker diseases can severely impact improperly grown roses leading to entire loss of the desired scion. Symptoms/signs. Small red to purple spots appear on current season canes. Spots enlarge into whitish necrotic lesions with reddish purple boarders during the winter months. Cankers may coalesce and can be 2.5–10 cm (1–4 in. long. Small black pycnidia can be seen in the cankered area. Cankers can cause wilt in and kill distal portions of the cane. It is easily confused with other canker diseases such as brand or common canker. Biology and epidemiology. High moisture environments (such as hilling for winter protection) favor the disease (Horst and Cloyd 2007). Spores can penetrate uninjured tissue. Lesions can form after 10 days under favorable conditions. Management • Cultural practices – Cultural practices similar to common canker will be effective. Sanitation is very important. Prune canes with a sharp knife or pruner immediately above a node when first symptoms are noticed. Cut and destroy infected canes well below the affected area. Remove and destroy canes before sporulation occurs.

2.7

Cercospora Leaf Spot (Pseudocercospora puderi B.H. Davis ex Deighton 1976) (Passalora rosicola (Pass.) U. Braun 1995)

Geographic occurrence and impact. Worldwide. Misidentification of this disease on black spot resistant cultivars can lead to defoliation and loss of vigor. Symptoms/signs. Numerous tiny maroon to purple oval lesions develop randomly across the leaf surface. The center of these spots then turns tan to gray, while the margins of spots remain maroon to dark purple. Heavily spotted leaves turn yellow and are prematurely shed. Typically, leaf loss begins at the base of the canes and gradually spreads upwards through the plant canopy towards the shoot tips. In the southeast USA, symptoms get worse through the summer and into early fall until many of the leaves are prematurely lost. Leaves are most often infected but stems, pedicels, fruits, and bracts can also have symptoms. Biology and epidemiology. Damage appears to be greater on shrub and ground cover roses compared with hybrid tea and grandiflora roses. Spores produced in the spots are rain splashed to healthy foliage. Rainy weather favors the disease. Management • Cultural practices – Avoid dense plantings and shaded areas. Avoid overhead watering that keeps plants wet for extended periods of time. Rake up and burn all leaves at season’s end.

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• Fungicides – FRAC fungicide groups M5, 3, and 11 were found to be effective (Hagan and Akridge 2005).

2.8

Common Canker (Paraconiothyrium fuckelii (Sacc.) Verkley & Gruyter 2012)

Geographic occurrence and impact. Commonly found worldwide. Canker diseases can severely impact improperly grown roses leading to entire loss of the desired scion. Symptoms/signs. Discolored (yellow to red) spots that gradually enlarge with the center turning light brown. Common canker has a darker brown margin and generally starts to develop at the cane’s cut end. Small black fruiting bodies (pycnidia) develop toward the center of damaged tissue. Cankers may girdle stems, causing stems and shoots above that point to wilt and die. The disease is easily confused with brown or brand canker. Biology and epidemiology. The fungus infects cane wounds caused by pruning, insects, thorn scars, or scratches (Intrama 1968). Common canker is favored by improper pruning, especially leaving long stubs above the last node. Common canker lesions developed more rapidly on young succulent stems than on older canes. High nitrogen fertility also increased common canker size. Management • Cultural practices – Avoid injuring canes. Prune canes with a sharp knife or pruner immediately above a node. Avoid leaving long stubs above a node. Cut and destroy infected canes well below the affected area. Use a winter protecting mulch that does not hold much water such as sand, rock pumice, or coarse bark. Use a balanced fertilizer with moderate nitrogen.

2.9

Cutting Rots (Cylindrocladium scoparium Morg. 1892 and Fusarium spp.)

Geographic occurrence and impact. Western North America and South America. A problem when trying to root new plants from soft wood cuttings. Symptoms. General yellowing, wilting, and loss of lower leaves. Yellowing progresses upward from the base of the cutting. Cuttings show a basal stem rot where the base of the cutting becomes water soaked and sunken and turns dark brown. Biology and epidemiology. The Oregon State University Plant Clinic has diagnosed cutting rots many times with the association of Fusarium spp., while Chase Ag Consulting, LLC, has reported that Cylindrocladium spp. are found in cutting rots of potted miniature roses (Chase 2015). Many fungi can be spread when contaminated rooting solutions are used just before sticking cuttings in media. A few infected cuttings in a dipping solution can inoculate all the other cuttings that

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come in contact with the same solution. Disease is encouraged by warm wet conditions, which are typical when trying to root cuttings (Philley et al. 2001). Management • Cultural practices – Take cuttings from field plants above the splash zone. Rain and irrigation can splash soil particles as well as microorganisms onto plants. Do not take cuttings from obviously or subtly diseased plants. Take cuttings in the morning and/or after irrigation when they are more fully hydrated. Use sharp tools to remove cuttings, buds, or leaf tissue. Clean tools and soak in a disinfectant. Use two tools so a worker can trim or cut with one while the other soaks. When done with a batch, cultivar, etc., switch one tool for the other and allow the used one to soak. Saturate media 24 h before sticking. Make bottom cuts through nodes to help stimulate meristematic activity. Make all cuts of stems or leaves with clean sharp tools. Use disposable gloves and aprons for workers. Disinfect the storage facility with peroxide or quaternary ammonium products before cuttings come in. Disinfect the sticking area before work begins and frequently while sticking. Disinfect carts, benches, and any containers that come in contact with cuttings. Clean the floors at least once a day. Use as little misting as possible after sticking cuttings. Use only enough water to prevent cuttings from wilting. Mist should not wet the media until roots are formed. Adjust misting based on weather conditions such as sunny vs cloudy weather. Water cuttings after sticking and once on the bench. Maintain moderate media moisture while cuttings are forming callus. Start drying the media once callus has formed and reduce mist. Once roots form, dry media a little more but just enough to prevent wilting. Uniform moisture from top to bottom of media is important. Careful and frequent scouting for problems is important. Remove, diagnose, and destroy any diseased cuttings. • Fungicides – Use products containing FRAC fungicide group 12 on the stock plants the day prior to cutting selection or just after sticking cuttings into media.

2.10

Downy Mildew (Peronospora sparsa Berk. 1862)

Geographic occurrence and impact. Commonly found worldwide and of major importance. Symptoms/signs. Purplish-red to dark brown, irregular spots on leaves. Leaflets may yellow but contain “green island” areas. Major leaf veins often restrict fungal growth such that lesions become angular as they enlarge. A grayish spore mass may be on the leaf underside if humidity persists. However, the diagnostic sporulation of this fungus is not abundant. Defoliation also may occur in extreme cases. Small spots or long purplish areas may form on canes and may kill twigs. Some people may confuse this with black spot or even rust. In the early stages, it may get confused with nutrient deficiency or spray injury. In advanced stages, stem lesions may have white sporulation (Fig. 4). Biology and epidemiology. This fungus-like organism can be systemic in roses. It overwinters as mycelium in stems, root crowns, and roots or as oospores in leaf

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Fig. 4 Downy mildew can easily be confused for black spot or other problems (Photo courtesy of Melodie Putnam (left) and Tatiana Gomez Rodriguez (center and right))

debris and stems (Aegerter et al. 2002). Humidity above 85% and cool temperatures favor the disease. Infection usually occurs on young plant parts. Cuttings taken from infected stock plants will carry the disease. Infected stock may or may not show symptoms. Excess nitrogen favors disease development. Management • Cultural practices – Lower humidity in the greenhouse by raising temperatures and venting at key times of day. Rake leaves and prune out old flowers and stems. Burn or bury them or send them to a landfill. Take cuttings from plants with no history of the disease. Scout plants carefully for symptoms of the disease especially the undersides of leaves and any stock plants or plants held over from the previous year. Remove and destroy any tissue with symptoms of the disease. Entire leaves should be removed even if one leaflet has symptoms. Although hot water soaks have been effective, they have also damaged cuttings. Hot water soaks are not recommended until time and temperature durations have been investigated. Maintain adequate, not excessive, fertility. Silicon applications have been found to help strengthen tissue and minimize tissue damage in Colombia. • Fungicides – Focus on cultural control methods first. Do not use mefenoxambased products (FRAC group 4 fungicides) alone as they may not offer much control, and encourage resistance to develop. Chemical control tactics help prevent infection but may not be helpful once the disease starts. Fungicides from the following groups are effective if resistance is not a problem: FRAC groups 4, 33, and 40 and some FRAC group 11 materials.

2.11

Myrothecium Gall (Myrothecium sp.)

Geographic occurrence and impact. Tropical regions.

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Symptoms/signs. A malformation or gall occurs at a cane node where a flower stem was removed. The gall is made of cortical tissue and can resemble the gills of a fish. The gall encompasses the cut surface and the lower portion of the new stem that developed from that node. The surface of the galled tissue becomes white to gray with orange yellow dots. Several galls can occur on the same stem. Small orange/ pink fruiting bodies (sporodochia) develop toward the center of damaged tissue (Fig. 5). Biology and epidemiology. The fungus infects through pruning wounds. Contaminated cutting knives, scissors, or pruners are a major inoculum source. The fungus is favored by temperatures between 21  C and 27  C with conditions of high relative humidity > 70%. Management • Cultural practices – Avoid injuring canes. Prune canes with a disinfected, sharp knife or pruner immediately above a node. Avoid leaving long stubs above a node. Frequently clean and disinfect cutting tools (refer to chapter “▶ Sanitation in Florists’ Crops Disease Management”). Cut and destroy infected canes well below the affected area. Avoid water splashes caused by irrigation, prefer the irrigation drip. Use a winter protecting mulch that does not hold much water such as sand, rock pumice, or coarse bark.

2.12

Petal Spots (Ghost Spot) (Many Fungi Possible Such as Bipolaris, Botrytis, Cercospora, and/or Cladosporium)

Geographic occurrence and impact. Worldwide. Damage makes flowers unmarketable. Symptoms. Numerous small spots develop on petals as buds are opening through flowering. Spots may have colored or darkened margins depending on the cultivar and fungus involved (Fig. 6). Biology and epidemiology. Warm, humid/wet conditions favor disease development especially during rainy weather. Management • Cultural practices – Keep rainfall off blooms. Prune off symptomatic buds as soon as disease is noticed.

2.13

Powdery Mildew (Podosphaera pannosa (Wallr.: Fr.) de Bary 1870)

Geographic occurrence and impact. Commonly found worldwide on roses and other members of the rose family. Symptoms/signs. White, powdery, fungal growth on young leaves, shoots, and buds. When scouting, check on the underside of leaves with good lighting

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Fig. 5 Galls can be starting at pruning wounds (Photo courtesy of Tatiana Gomez Rodriguez)

Fig. 6 Botrytis sp. has infected the petals (Photo courtesy of Melodie Putnam (left) and Tatiana Gomez Rodriguez (center and right))

conditions, as this is where it may first start. Distorted growth is common especially if young tissues are infected. If the disease is severe, foliage is stunted (Fig. 7). Biology and epidemiology. The fungus overwinters in infected buds, rose leaves, twigs and branches. Infected buds grow in spring but are stunted and white with fungal conidia. Conidia are released in response to abrupt decreases in relative humidity. Wind blows conidia to healthy foliage where they start new mildew colonies. Germination and growth of conidia happen most readily on nights with high humidity or heavy dew and at temperatures near 21  C (70  F). Too much water, such as flowing water or rain, destroys spores by causing them to burst. The fungus does not grow in a leaf, but rather across the surface. Small anchor cells of the fungus, haustoria, remain in the leaf and take nourishment from the rose plant, but the main filaments and the multitudes of spores it produces are outside the leaf.

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Fig. 7 Powdery mildew symptoms are not generally seen on the flower petals, but early infections can deform flower buds and affect the bloom (Photo courtesy of Jay W. Pscheidt (left) and Tatiana Gomez Rodriguez (center and right))

Newly unfurled leaves are more susceptible to infection than mature leaves. Increasing day lengths to 20–22 h were shown to reduce conidial production on greenhouse roses while maintaining postharvest quality (Suthaparan et al. 2010). Brief exposure to red light during the dark period may also do the same. Small, black fruiting structures (chasmothecia) also allow the fungus to overwinter on plant parts and debris. Ascospores from chasmothecia initiate new infections. The disease can develop under relatively dry conditions so long as the air is humid. Plants in shade tend to have more problems because that environment favors disease development. Multiple disease cycles occur during the growing season. Climbers, ramblers, and hybrid teas are susceptible. The natural genetic variability of the fungus means roses found resistant in one location may be susceptible in another due to the presence of different fungal strains. Also, resistant roses may become susceptible after a few years due to changes in the local fungal population. Management • Cultural practices – Plant cultivars known to be powdery mildew resistant in your area. Note that the cultivar Carefree Spirit was the first landscape shrub to survive for 2 years in AARS tests without any fungicide and voted a winner. Isolate susceptible cultivars. Space plantings for good air circulation between plants. Prune canes when dormant for an open habit also for good air circulation. Remove and destroy diseased canes. Rake and destroy fallen leaves. Use a highpressure water hose to thoroughly wet all leaf and cane surfaces in the early afternoon so plants dry quickly. May need to do this two or three times a week. In Western Oregon, this reduced but did not eliminate powdery mildew. Briefly exposing plants to red light during the dark interval has some utility against powdery mildew in greenhouse production. • Fungicides – Applying a dormant spray of lime sulfur at one part lime sulfur to nine parts water may help a planting that had severe powdery mildew the year

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before. Foliar applications of potassium salts have been found in Colombia to help reduce fungal sporulation. Apply foliar fungicides during the growing season, starting in early spring when young growth first appears. May need frequent applications, depending on the fungicide, to control the disease. Almost every fungicide on the market is registered for this disease on this crop, but only one or two materials are needed at any one time. Alternate or tank-mix products with different modes of action to prevent buildup of resistant fungal populations. Rosarians may not like the plant growth regulation effect (slight stunting) that may result from the use of FRAC group 3 or group 8 fungicides. Use of sulfur, which is a very effective organic material, can also stunt shoots, and frequent handling of sprayed foliage can irritate skin. Chlorothalonil-based products may be a problem on “Knock Out” and “Double Delight” roses resulting in damaged foliage. Always watch for warnings on labels of various products. • Biological control – Various Bacillus-based products are available and may have efficacy beyond the use of water alone but will not be as effective as many synthetic materials.

2.14

Replant Disease (Many)

Geographic occurrence and impact. The problem appears to be important on roses in Europe but has not been recognized in the Pacific Northwest of North America. Symptoms. The poor growth produced in replant situations may not be noticed unless you had a comparison. Roses planted into soil that has never grown roses (virgin soil) or on fumigated or sterilized soil would produce much more luxuriant, vigorous growth than when the same rose is replanted in an area that has grown roses for many years. Biology and epidemiology. Replant diseases are complexes of biological and environmental factors that vary by geographic region (Pscheidt and Ocamb 2016). This is a real disease complex in the Pacific Northwest (PNW) of the USA for many crops in the rose family including pome fruit such as apple and pear and stone fruit such as cherry and peach. Similar problems exist for other crops such as strawberry (black root rot). In the PNW, roses planted into grounds that have grown roses before appear to grow fine. The Washington Park International Rose Test Garden in Portland OR reports that they routinely remove a thousand plants every winter and replant a thousand new ones. They have not noted a replant problem.

Management • Cultural practices – When replanting roses, follow correct planting practices. Correct any soil problems such as pH or drainage. You can also plant into new areas of your production site or rotate out of rose crops for several years before planting roses again on the same ground.

Diseases of Rose

2.15

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Rust (Phragmidium mucronatum (Pers.) Schltdl. 1824 and Many Other Species.)

Geographic occurrence and impact. Worldwide. Repeated defoliation leads to low vigor, inferior blooms, and high susceptibility to winter injury. Phragmidium americanum (Peck) Dietel 1905 – Eastern North America and Japan P. andersonii Shear 1902 – Northern North America; Asia; Europe P. bulbosum (F. Strauss) Schltdl. 1824 – Africa, Asia, Europe P. butleri Syd. & P. Syd. 1907 – Asia P. fusiforme J. Schröt. 1870 – Temperate northern hemisphere P. kamtschatkae (H.W. Anderson) Arthur & Cummins 1933 – Asia, Europe P. montivagum Arthur 1909 – Temperate North America and Asia; USSR P. mucronatum (Pers.) Schltdl. 1824 – Worldwide P. occidentale Arthur 1901 – North America P. rosae-arkansanae Dietel 1905 – Central North America P. rosae-californicae Dietel 1905 – Western North America P. rosae-pimpinellifoliae Dietel 1905 – North America; temperate Europe P. rosicola (Ellis & Everh.) Arthur 1934 – Central North America P. speciosum (Fr.) Burrill 1875 – North America P. tuberculatum J. Müll. 1885 – Asia, Africa, Europe, North and Central America Symptoms/signs. Small orange pustules (aecia) appear early in spring on both leaf surfaces. Later, the pustules enlarge and become more numerous on lower leaf surfaces (Fig. 8). Mottled and chlorotic areas may develop on upper leaf surfaces opposite the spots (uredinia) on the lower surfaces. In late summer and fall, the small pustules turn black (telia) and contain the winter spore stages of the rust. Stems occasionally are infected. Biology and epidemiology. Many species of these autoecious fungi (fungi that complete their life cycle on one plant host) are found throughout the world. They overwinter on diseased leaves and stems. Wind blows spores to healthy foliage. They germinate and infect through the stomata when leaves are wet for 2–4 h. Mild, humid weather favors disease development.

Fig. 8 Note the two different rust pustules on this leaf, orange uredia and black telia (Photo courtesy of Jay W. Pscheidt)

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Management • Cultural practices – Rake up all dead leaves and prune out infected and dead wood during the dormant season. Plant resistant cultivars. Removing infected leaves early in the season may be effective in some small plantings. • Fungicides – Focus applications during wet weather. Wetting agents will help with many of fungicides if allowed by the label. Many fungicides used for black spot will be helpful to control rust.

2.16

Spot Anthracnose (Elsinoe rosarum Jenk. & Bitanc. 1957)

Geographic occurrence and impact. Temperate regions. Repeated defoliation leads to low vigor, inferior blooms, and high susceptibility to winter injury. Symptoms/signs. At first, the appearance of red spots that vary from brown or dark purple on the upper leaf surfaces occurs (Bagsic et al. 2015). Spots up to 0.5 cm in diameter may be scattered or grouped and sometimes overlapping. Chlorosis or yellowing of the leaves may also occur. Later, spots become ashen white with a dark red margin. This tissue may fall off of the lower leaf surface, leaving a thin papery membrane or fall out entirely resulting in a shothole symptom. Symptoms occur on stems, hips, and pedicels as well as leaves. Can be easily confused with black spot or Cercospora leaf spot. Biology and epidemiology. The fungus overwinters on infected leaves and stems with conidia continuously being formed in early spring and through the summer. Spore dispersal primarily occurs with the aid of water through rain and irrigation. There were 5 races of the fungus identified in Germany (Bagsic et al. 2015). Management • Cultural practices – Remove and destroy infected plant parts. Space or prune bushes to allow for good airflow. • Fungicides – No specific chemical control has been reported. Fungicides in the FRAC group 3 used to control black spot, powdery mildew, or rust are also being used by growers to control this disease (Bagsic et al. 2015).

2.17

Verticillium Wilt (Verticillium dahliae Kleb. 1913)

Geographic occurrence and impact. Worldwide. Symptoms. At first, leaves near the growing point of young canes wilt, and lower leaves yellow (Hammett 1971). Sometimes mature leaves become necrotic between veins while the veins remain green. Defoliation progresses from the base of canes to the tip. Permanent wilt, defoliation, and death also may occur. If only a few canes are infected, they may grow normally next season or dieback. Symptoms generally are more severe in the greenhouse than in the field. The characteristic vascular discoloration in other plants is not evident in rose. Biology and epidemiology. This fungus survives a long time in soil and can infect a wide range of hosts. Rootstocks such as Rosa odorata and “Ragged Robin”

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are susceptible; R. multiflora and “Dr. Huey” have more resistance. R. chinensis var. manetti is very resistant. Under favorable growing conditions, plants may be able to tolerate infection. The fungus grows into the xylem where it colonizes the plant through mycelial growth and conidial production. Fluid movement in the xylem passively transports the conidia. Once in the xylem, this fungus partially blocks water movement and produces toxins that result in wilt symptoms. Wilting occurs under periods of water stress such as midsummer heat and drought. After diseased plant parts die, microsclerotia form and live several years in soil. Many weeds are susceptible and can help the fungus survive and disperse. Send soil samples to any of various private and public laboratories to assay for Verticillium propagules. Nurseries may wish to test individual core samples to determine the distribution in a particular field. The presence of any microsclerotia in the soil should be interpreted as a potential disease risk. Management • Cultural practices – Use resistant rootstocks. Avoid planting in old vegetable fields. In the greenhouse, use steam-sterilized soil or use a sterile soilless potting mix and/or hydroponics. Remove and destroy symptomatic or dead branches preferably before leaves fall and thus before new inoculum gets incorporated into the ground. Clean pruning equipment after use. Mulch with conifer-based products as conifers are resistant to infection. General yard waste mulch could contain infected plants and thus inoculum of the fungus.

2.18

Miscellaneous Fungal and Oomycete Diseases of Minor Importance or Considered Rare

Alternaria leaf spot – Alternaria spp. Phytophthora root rot – Phytophthora spp. have been identified on rose samples with root rot from the PNW several times by the Oregon State University (OSU) Plant Disease Clinic. Also reported from Asia, Europe, and North America. Septoria leaf spot – Septoria rosae Desm. 1831. Southern blight – Sclerotium rolfsii Sacc. 1911. Rhizoctonia root and stem rot – Rhizoctonia spp. – Japan. Note: Fungal names based on Farr and Rossman (2015).

3

Bacterial and Phytoplasma Diseases

3.1

Bacterial Cane Blight (Pseudomonas syringae – Undetermined Pathovar)

Geographic occurrence and impact. Treasure Valley of Idaho, USA (Mohan and Bijman 2010).

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Symptoms/signs. The symptoms usually start at the base of a vegetative bud or at leaf scars or wounds, as reddish-brown areas on the bark that later turn dark purple to black and necrotic (Fig. 9). The necrotic areas expand around and along the cane, often involving a major part or even the entire cane. Vegetative buds on the affected parts of the cane turn brown and dried. The surface of the necrotic areas of the bark is glossy, and the tissue beneath the epidermis is brown to dark brown and moist in the early stages. Often confused with winter injury. Biology and epidemiology. This disease is different from the various symptoms on roses previously attributed to P. syringae and/or P. syringae pv. morsprunorum (see below) in the literature. It has been an aggressive cane blight observed in the Treasure Valley of Idaho, USA, since 1996. Symptoms were common under cool, wet conditions in spring (March to May), and the level of incidence and severity of the disease varied from year to year. Several cultivars of climbing, floribunda, grandiflora, hybrid tea, hybrid perpetual, miniature, and shrub roses can show severe symptoms. Management • Cultural practices – The following is suggested in the absence of specific research to control this problem. Remove and destroy infected stems. Disinfect pruning shears before cutting more stems. Copper-based products are also recommended before fall rains begin and again when half the leaves have fallen.

3.2

Bacterial Leaf Spot and Blast (Pseudomonas syringae pv. morsprunorum )

Geographic occurrence and impact. Pacific Northwest of North America – rare. Symptoms. Dark brown, sunken spots appear on leaves, flower stalks, and calyx parts. Flower buds may die without opening. Black streaks appear on 1-year-old stems. Fig. 9 Glossy, dark purple to black, necrotic bark of canes with bacterial cane blight (Photo courtesy of Krishna Mohan)

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Biology and epidemiology. The disease has been diagnosed only four times in the PNW from 1962 to 1992. Most common in cool, wet spring weather. Management • Cultural practices – Remove and destroy infected stems. Disinfect pruning shears before cutting more stems. Copper-based products are also recommended before fall rains begin and again when half the leaves have fallen. Repeat in spring to protect new growth.

3.3

Crown Gall (Agrobacterium radiobacter (Beijerinck and Van Delden 1902) Conn 1942 or Also Known as Rhizobium radiobacter (Beijerinck and Van Delden 1902) Young et al. 2001)

Geographic occurrence and impact. Worldwide and common. Symptoms. Galls are often at or just below the soil surface in the basal or root crown region. They may frequently be on roots but less frequently on aerial plant parts such as stem and leaves. Galls are usually rounded with a rough, irregular surface (Fig. 10). They first appear as small protuberances on the plant surface. Young, actively developing galls are light green or nearly white, and the tissue is soft. As they age, galls become dark and woody. Outer portions can slough off with age. Sometimes galls have a rather smooth surface, which makes it difficult to distinguish between gall and callus growth, especially if the gall is at the plant

Fig. 10 Galls can be seen above, below ground, starting at pruning wounds or wounds on leaves (Photo courtesy of Jay W. Pscheidt (left) and Tatiana Gomez Rodriguez (center and right))

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base or at the graft or bud union. Plants can be stunted and have reduced vigor, poor foliage, and fewer blossoms. A single gall at the plant’s base may be more detrimental than several galls on canes and roots. Symptoms may not develop for over a year if infection occurs when temperatures are below 15  C (59  F). Biology and epidemiology. The bacterium enters plants through wounds, either natural or caused by pruning, grafting, mechanical injury from cultivation, heaving of frozen soils, chewing insects, or the emergence of lateral roots. Systemic populations that initiate disease do not seem to be as important in rose as they are in grape and walnut. After the bacterium enters a wound, a small piece of its DNA is transferred into the plant’s DNA. The foreign DNA transforms normal plant cells in the wounded area into tumor cells. Once transformed, tumor cells proliferate automatically. The result is a gall: a disorganized mass of hyperplastic and hypertrophic tissue. Pruning tools that cut through galls can become contaminated with the bacteria and spread them to cut surfaces of subsequently pruned plants. Galls breaking down in soil release bacteria, which can be transported by moving soil or water. In the absence of plant roots, bacterial populations gradually decrease; however, the pathogen may survive in soil for as long as 3 years. Management • Cultural practices – Use only pathogen-free nursery stock. Inspect new plants; do not plant any rose that has galls. Avoid wounding plants, especially at planting. If root pruning at planting, soak in the biocontrol agent listed below. Use plants with resistant rootstocks. Rootstocks differ widely in susceptibility. Prune off any galls on aerial parts of the plant. Disinfect pruning shears frequently. Clean shears and long soak times improve the disinfectant’s efficacy. Remove and destroy badly affected plants. Preplant soil solarization has been effective against this disease for cherry nursery stock grown in Western Oregon and may be useful for roses. Place clear plastic (anti-condensation coating) on rototilled ground, irrigated to near field capacity, from mid-July to mid-September. Solarization is more effective on sandy loam soil. The technique may help after removing diseased plants from a bed in which roses will be planted again. • Biological control – Agrobacterium radiobacter strain 84 has been used successfully with roses in Australia, New Zealand, and Spain but has not been effective in limited trials in the USA. Strain K 84 is preventive only. Agrobacterium radiobacter strain K1026 is a genetically modified strain of K84 that will help reduce the potential for development of resistant crown gall bacteria. Latent infections (symptomless) and existing galls are not controlled. A suspension of strain 84 may be used as a soak or spray. Thoroughly cover grafting wood, roots, and crown. Spray to runoff. To be effective, it must be applied a few hours after wounding.

Diseases of Rose

3.4

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Miscellaneous Bacterial Diseases of Minor Importance or Considered Rare

Hairy root – Agrobacterium rhizogenes (Riker et al. 1930) Conn 1942

4

Viral Diseases

4.1

Rose Leaf Curl (A Virus Is Suspected)

Geographic occurrence and impact. California. Symptoms/signs. Spring leaves are small, leaflets detach easily, leaf epinasty, necrosis of shoot tips, and yellow vein flecking occur. Shoots are pointed with a broad base. Plants may recover in summer but show symptoms again in fall. Biology and epidemiology. Little is known about this disease other than natural spread appears to be slow. Management • Cultural practices – Purchase stock that is clean and/or certified as tested and free of all known viruses. Remove and destroy infected plants. Heat-treat scion stock plants 4 weeks at 38  C (100  F) before grafting.

4.2

Rose Mosaic [Prunus necrotic ringspot virus (PNRSV), Apple mosaic virus (ApMV), Arabis mosaic virus (ArMV), and Strawberry latent ringspot virus (SLRSV)]

Geographic occurrence and impact. Worldwide. Symptoms/signs. Symptoms may range widely depending on time of year, temperature, and type of virus(es) infecting the plant. Characteristic symptoms include chlorotic line patterns (zigzag pattern), ringspots, and mottles in leaves sometime in the growing season (Fig. 11). There may also be yellow net and yellow mosaic symptoms. Symptoms often are evident in spring and early summer but may not be on leaves produced in summer. Vein banding may be on leaves in long hot periods. Flower distortion, reduction in flower production, flower size, stem caliper at the graft union, winter survival, and early leaf drop and increase susceptibility to cold injury have all been reported. Some infected cultivars may not show any symptoms at all. Biology and epidemiology. Several viruses are associated with the range of symptoms of rose mosaic, including Prunus necrotic ringspot virus (PNRSV), Apple mosaic virus (ApMV), Arabis mosaic virus (ArMV), and Strawberry latent ringspot virus (SLRSV). The disease does not spread naturally and has no known insect vector, but grafting transfers it to healthy plants. Transmission of ArMVand/or SLRSV could occur by Dagger nematodes, but this has not been extensively studied. Viruses can be in the rootstock or scion or both and may not show symptoms.

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Fig. 11 Each of these leaves was from the same plant with rose mosaic (Photo courtesy of Jay W. Pscheidt)

“Madame Butterfly,” “Ophelia,” and “Rapture” are highly susceptible. Some report the disease does not spread; others indicate it may spread very slowly over many years. Root grafting between infected and healthy plants can also spread the disease (Golino et al. 2011). Management • Cultural practices – Purchase stock that is clean and/or certified as tested and free of all known viruses. Remove and destroy infected plants. However, the disease will not spread unless you propagate from or onto an infected bush. Heattreat scion stock plants 4 weeks at 38  C (100  F) before grafting.

4.3

Rose Rosette (Rose Rosette Virus)

Geographic occurrence and impact. Widespread east of the Rocky Mountains of North America. Plants decline and die after a few years. Symptoms. Rose rosette symptoms are complex and variable as plants of the same cultivar may have different symptoms at the same or different location(s). Infected plants may have foliage that is bright red throughout the summer rather than just in the spring and fall. Leaves may become unusually long and thin or strapped shaped. Increased thorniness and flattening of stems (fasciation) is often but not always observed. Symptomatic foliage is often more susceptible to winter kill and/or desiccation. Multiple shoots can emerge from a single node to form a witches’ broom. Witches’ brooms are easier to recognize in the winter months after most of the foliage has fallen. Unusually large masses of distorted flower buds may also occur that, in most cases, do not open. Infection can resemble herbicide (glyphosate) injury. Infected bushes will decline and die in 3–5 years. Cane mortality is usually observed in spring when symptomatic canes fail to push out new foliage. Large commercial plantings or private rose gardens can be decimated by rose rosette if the disease is left unchecked.

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Biology and epidemiology. All cultivated roses (shrub type, hybrid tea, floribunda, grandiflora, and miniature roses) are thought to be susceptible to the disease. The Knock Out rose cultivars are as susceptible to rose rosette as other types. The virus is vectored by an eriophyid mite, Phyllocoptes fructiphilus. Although these mites do not fly, they may “balloon” in air currents, as do dust particles, and thus can be spread surprisingly long distances. The closer a healthy rose is planted to an infected rose, the more likely it is to become infected. In Tennessee, rose beds located near a source of the virus have a pronounced edge effect where roses nearest the source are more likely to become infected. Distribution of infected plants in a large rose bed will appear random if the plants were infected prior to planting or if there is a great distance between the rose planting and the inoculum source. Management • Cultural practices – Purchase stock that is clean and/or certified as tested and free of all known viruses. Roses should be inspected for symptoms before being purchased. Purchase plants from a nursery where all roses appear to be healthy. Inspected regularly for symptoms and rogue entire plants soon as possible when found. Do not just prune off symptomatic stems. Rogued plants should be bagged on site before removal. At this writing, preliminary studies have not demonstrated that miticides are effective. Also, it is not known if disinfecting pruning shears will also aid in disease management. Roses reported to be resistant include: R. setigera, R. aricularis, R. arkansana, R. blanda, R. palustris, R. carolina, and R. spinosissima.

4.4

Rose Spring Dwarf (Rose Spring Dwarf-Associated Virus)

Geographic occurrence and impact. California and Chile. Symptoms/signs. A rosetting or balling of new growth (Salem et al. 2008). Leaves are curved, very short, and show vein clearing or a netted pattern. Leaves occur on arrested shoots, which may not elongate for several months. Canes may grow in a zigzag pattern during the growing season. Biology and epidemiology. Easily transmitted through vegetative propagation techniques. The virus has been found in aphids on affected plants but is not known if they are a vector of the disease or not (Rivera and Engel 2010). Management • Cultural practices – Purchase stock that is clean and/or certified as tested and free of all known viruses. Remove and destroy infected plants. Heat-treat scion stock plants 4 weeks at 38  C (100  F) before grafting.

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4.5

Rose Yellow Mosaic (Rose Yellow Mosaic Virus)

Geographic occurrence and impact. North America. Symptoms/signs. Symptoms associated with RoYMV infection include yellow mosaic, ring mosaic, premature leaf senescence, and necrotic dark brown rings on canes. Symptoms first appeared in new growth 4–6 weeks postinoculation. Necrotic cane symptoms are observed only in the cultivar Ballerina. Symptoms persist throughout the season (Lockhart et al. 2008). Biology and epidemiology. Potyviruses such as RoYMV are known to be aphid transmitted. Management • Cultural practices – Purchase stock that is clean and/or certified as tested and free of all known viruses. Remove and destroy infected plants.

4.6

Miscellaneous Viral or Virus-Like Diseases of Minor Importance or Considered Rare

Rose flower break – England, New Zealand, and Australia – unknown causal agent Rose Streak – eastern and mid-North America and Europe – suspect Rose streak virus The following have only been associated with virus symptoms in rose and are not fully characterized; however, the four viruses commonly found in the rose mosaic complex were not present in plants infected with these viruses (Lockhart et al. 2008): Rose yellow leaf virus Rosa rugosa leaf distortion virus Rosa multiflora cryptic virus Rose chlorotic ringspot virus Rose necrotic mosaic virus

5

Nematode Diseases (Meloidogyne hapla, Pratylenchus Penetrans, P. Vulnus, and Xiphinema Diversicaudatum)

Geographic occurrence and impact. Worldwide. More of a problem on sandy soils. Symptoms/signs. Specific diseases have not been linked with specific nematodes; however, specific nematodes are associated with various root symptoms. In general, nematodes disrupt the root system, which will result in general aboveground symptoms that can include reduced vigor, poor growth or flowering, stunting, nutrient deficiencies, chlorosis, and/or wilting. These may be combined and considered a general plant decline.

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Root swellings or galls occur from the feeding activity of root-knot nematodes (Meloidogyne hapla) or dagger nematodes (Xiphinema spp.). Dagger nematode form galls on the tips of feeder roots. Root-knot nematodes tend to form galls on smaller roots and, with magnification, the swollen females can also be seen. Excessive root branching may also be observed on root systems with root knot. Small root lesions that become necrotic can be observed with an infestation of Root-lesion nematodes (Pratylenchus spp.). The severity of the symptoms will depend on the soil type and ability of the rose to tolerate various sized populations of root-lesion nematodes. Biology and epidemiology. Nematodes that affect plant growth are classified by feeding behavior. Depending on the nematode, this will dictate what type of samples is needed for diagnosis: roots, soil, or both. Root-knot nematodes (Meloidogyne spp.) are sedentary endoparasites, which tunnel into the roots, establishing permanent feeding sites from which they do not move. The feeding site causes the formation of giant cells, hyperplasia of cortical and vascular parenchyma, and retardation of meristematic activity in root tips (Fig. 12). Root-lesion nematodes (Pratylenchus spp.) are migratory endoparasites that tunnel inside roots, feed inside roots, and freely move back into soil and on to new roots (Fig. 13).

Fig. 12 Adult Meloidogyne sp. and egg mass (Photo courtesy of Tatiana Gomez Rodriguez)

Fig. 13 Adult Pratylenchus sp. and Xiphinema sp. (Photo courtesy of Tatiana Gomez Rodriguez)

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Dagger nematodes (Xiphinema spp.) are migratory ectoparasites feed from outside roots, moving from cell to cell and piercing them to feed without entering root tissue. The galls that are produced at this feeding site are caused by a hyperplastic response of cortical cells. The cells increase two to three times in size and meristematic activity is retarded. Although barely studied in rose, these nematodes can be vectors for some of the viruses in the rose mosaic complex such as Arabis mosaic virus (ArMV) (Fig. 13). Soil types that are more sandy generally result in more nematode injury at lower population levels than less porous soils. Host ranges are wide for many of these nematodes making crop rotation impractical. Management • Cultural practices – Avoidance in floral production is accomplished by eliminating soil from the production system with soilless media and/or hydroponics. If soil must be used then steam treatment is necessary to eradicated nematodes from the soil. Although difficult, soil in planting beds can also be steam treated. Soil solarization is useful to reduce populations in the top foot of soil in order to get plants establish in infested soil. Thermal therapy after infestation of plants is also difficult but possible. Pretreat plants at 38 C for 24 h followed by immersing plants into hot water at 48 C for 35 min. Damage to rose roots with this treatment is possible. Use of resistant rootstocks is complicated by diverse reactions to different nematodes from or in different locations. • Chemical practices – Preplant fumigation can be effective in the field, but available (registered) nematicides are becoming limited each year.

References Aegerter BJ, Nunez JJ, Davis RM (2002) Detection and management of downy mildew in rose rootstock. Plant Dis 86:1363–1368 Bagsic I, Linde M, Debener T (2015) Genetic diversity and pathogenicity of Sphaceloma rosarum (teleomorph Elsinoë rosarum) causing spot anthracnose on roses. Plant Pathol. doi:10.1111/ ppa.12478 Bowen KL, Roark RS (2001) Management of black spot of rose with winter fungicide treatment. Plant Dis 85:393–398 Chase AR (2015) Rose diseases and their control. Chase Agricultural Consulting. http://www.chaseagri culturalconsultingllc.com/resources/pdfs/articlesPdf/48ROSEDISEASESANDTHEIRCONTROL.pdf Farr DF, Rossman AY (2015) Fungal databases, systematic mycology and microbiology laboratory, ARS, USDA. http://nt.ars-grin.gov/fungaldatabases/ Golino DA, Sim ST, Cunningham M, Rowhani A (2011) Evidence of root graft transmission of two rose mosaic viruses, Prunus necrotic ringspot virus and Apple mosaic virus in rose rootstocks. Phytopathology 101:S62 Hagan AK, Akridge JR (2005) Chemical control of cercospora leaf spot on Fuchsia Meidiland ® shrub rose. Alabama Cooperative Extension PP-587 Hammett KRW (1971) Symptom differences between rose wilt virus and Verticillium wilt of roses. Plant Dis Rep 55:916–920

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Hausbeck MK, Moorman GW (1996) Managing Botrytis in greenhouse-grown flower crops. Plant Dis 80:1212–1219 Horst RK, Cloyd RA (2007) Compendium of rose diseases and pests, 2nd edn. APS Press, St. Paul Intrama S (1968) Coniothyrium rose canker in Oregon. PhD thesis, Oregon State University. p 95 Lockhart B, Zlesak D, Fetzer J (2008) Identification and partial characterization of six new viruses of cultivated roses in the USA. In: XII international symposium on virus diseases of ornamental plants. Acta Hortic: Ag Exp Station Technical Bulletin #8. Oregon State College, Corvallis, Oregon 901:139–147 Milbrath JA (1946) Control of black mold fungus Chalaropsis thielavioides Peyr. on Manetti rose, vol 8, Oregon State University technical bulletin Mohan SK, Bijman VP (2010) Bacterial cane blight of rose caused by Pseudomonas syringae. Acta Hortic 870:109–113 Philley G, Hagen AK, Chase AR (2001) Chapter 76. Rose diseases. In: Jones RK, Benson DM (eds) Diseases of woody ornamentals and trees in nurseries. American Phytopathological Society Press, St. Paul Pie K, De Leeuw GTN (1991) Histopathology of the initial stages of the interaction between rose flowers and Botrytis cinerea. Eur J Plant Pathol 97:335–344 Pscheidt JW, Ocamb CM (eds) (2016) Pacific Northwest plant disease management handbook. Oregon State University, Corvallis Rivera PA, Engel EA (2010) Presence of rose spring dwarf-associated virus in Chile: partial genome sequence and detection in roses and their colonizing aphids. Virus Genes 41(2):295–297 Salem N, Golino DA, Falk BW, Rowhani A (2008) Identification and partial characterization of a new Luteovirus associated with rose spring dwarf disease. Plant Dis 92:508–512 Schnabel G, Agudelo P, Henderson GW, Rollins PA (2012) Aboveground root collar excavation of peach trees for Armillaria root rot management. Plant Dis 96:681–686 Suthaparan A, Stensvand A, Torre S, Herrero ML, Pettersen RI, Gadoury DM, Gislerød HR (2010) Continuous lighting reduces conidial production and germinability in the rose powdery mildew pathosystem. Plant Dis 94:339–344

Diseases of Snapdragon Stephen N. Wegulo and A. R. Chase

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-Like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Anthracnose (Colletotrichum antirrhini, C. destructivum) . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Botrytis Blight (Botrytis cinerea) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Cercospora Blight (Cercospora antirrhini) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Downy Mildew (Peronospora antirrhini) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Phyllosticta Blight (Phyllosticta antirrhini) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 Phytophthora Stem Rot and Wilt (Phytophthora cactorum, P. cryptogea, P. parasitica) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7 Powdery Mildew (Golovinomyces orontii (Formerly Erysiphe cichoracearum); Oidium spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.8 Pythium Root Rot (Pythium spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.9 Rhizoctonia Basal Stem Rot (Rhizoctonia solani) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.10 Rust (Puccinia antirrhini) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.11 Verticillium Wilt (Verticillium albo-atrum, V. dahliae) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.12 Additional Fungal Diseases and Pathogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Bacterial Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Seedling Blight (Pseudomonas antirrhini) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Additional Bacterial Diseases and Pathogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Viral Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Impatiens Necrotic Spot Virus (INSV) and Tomato Spotted Wilt Virus (TSWV) . . . . 4.2 Additional Virus Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2 3 3 4 5 6 8 10 11 12 13 14 15 16 16 16 17 18 18 19

S.N. Wegulo (*) Department of Plant Pathology, University of Nebraska-Lincoln, Lincoln, NE, USA e-mail: [email protected] A.R. Chase (*) Chase Agricultural Consulting, Cottonwood, AZ, USA e-mail: [email protected] # Springer International Publishing AG 2016 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florist's Crops Diseases, Handbook of Plant Disease Management, DOI 10.1007/978-3-319-32374-9_24-1

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5 Nematode Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 Root-Knot Nematode (Meloidogyne spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2 Additional Nematode Pathogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 Additional Parasites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 Nonparasitic Disorders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1 Tip Blight . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2 Phytotoxicity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.3 Additional Nonparasitic Disorders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract

Snapdragons (Antirrhinum majus) are grown as bedding and container plants and as cut flowers. Diseases can be a limiting factor in the production of snapdragons. Fungal diseases commonly encountered in snapdragons include downy mildew, powdery mildew, Botrytis blight, and various leaf spots and root, crown, and stem rots. The main bacterial disease is a seedling blight caused by Pseudomonas spp. Impatiens necrotic spot virus (INSV) and tomato spotted wilt virus (TSWV) are the two major virus diseases affecting snapdragons. Root-knot and lesion nematodes can also attack snapdragons. Yield loss can be reduced and profitability increased by concerted efforts to manage these diseases. Management tactics include host resistance and cultural, chemical, biological, and physical control. Integration of as many of these tactics as practical will maximize the effectiveness of disease management. This chapter presents information on the occurrence, symptoms, biology, epidemiology, and management of individual diseases of snapdragon. Details are provided for diseases on which research has been done and published. Some diseases have only been observed and research has not been done or published. Such diseases are only briefly mentioned. Keywords

Anthracnose • Blight • Leaf spot • Root and crown rot • Wilt • Mildew • INSV • TSWV

1

Introduction

Snapdragons (Antirrhinum majus) are popular plants that have been grown for centuries. They originated in the Mediterranean region (Maree and Wyk 2010; Rogers 1992), but are now grown in many parts of the world as cut flowers and garden, bedding, and potted plants. Cultivar groups differ principally in plant size. Dwarf types are used in gardens, whereas intermediate and taller types are grown for cut flowers. They display a range of beautiful flower colors from white to various shades of orange, yellow, peach, pink, red, and purple (Creel and Raymond Kessler 2007; Maree and Wyk 2010). In the USA, snapdragons were first included in the Agricultural Census of Horticultural Specialty Crops in 1959. At that time, they ranked seventh as the most valuable cut flower and made up 3.2 % ($4.5 million) of

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the total wholesale value of flowers produced in the USA ($142.5 million), mostly in midwestern and northeastern states (Rogers 1992). In 2006, the wholesale value of snapdragon as a cut flower in the USA for operations with sales worth $100,000 or more was $12.2 million or 3 % of the total value ($411.3 million) of cut flowers (Jerardo 2007). Diseases can significantly reduce marketability of snapdragons as plants or cut flowers. The most common diseases include Botrytis blight, powdery mildew, downy mildew, leaf spots, and root and crown rots. Viruses and nematodes also can attack snapdragons. The diseases found in a given location or geographical area will vary depending on environmental conditions. Losses due to diseases can be mitigated by integrating available management tactics which include cultural practices, physical control, chemical control, biological control, and host resistance.

2

Fungal and Fungus-Like Diseases

2.1

Anthracnose (Colletotrichum antirrhini, C. destructivum)

Geographic occurrence and impact. Snapdragon anthracnose has been reported in the USA (Stewart 1900; Horst 2013) and in Japan (Tomioka and Nishikawa 2011). It occurs most commonly on snapdragons grown in the field or landscape, but can also occur on greenhouse-grown snapdragons. Symptoms/signs. The disease can occur on snapdragons at any growth stage. Leaf spots are circular and slightly sunken. They are initially yellowish green with indefinite borders, but soon become whitish with definite borders. Spots on stems are circular to elliptical and initially whitish with a narrow brown or reddish-brown border. Spots may enlarge and girdle the stem base causing death of lateral shoots. Spots on older, woody stems are sunken. Under high humidity, numerous acervuli form in diseased tissues on the leaves and stems, causing them to become smoky brown (Forsberg 1975; Nelson and Strider 1985; Pirone 1978). Biology and epidemiology. Conidia are produced in acervuli in a sticky mass. They are dispersed by splashing water and may be carried by wind over long distances. Free water is necessary for conidial germination and infection. Disease development is favored by moisture and high humidity. Therefore, the risk for epidemics is higher in outdoor than in greenhouse snapdragons especially during wet growing seasons (Nelson and Strider 1985). C. antirrhini is a synonym of C. gloeosporioides which has a wide host range (Farr et al. 1989). Management. Adjust heating and ventilation to lower humidity and prevent condensation in the greenhouse. Avoid overhead watering. Apply fungicides labeled for anthracnose control on ornamental crops. Some of the most effective products contain a strobilurin (like pyraclostrobin or trifloxystrobin), a triazole (like triadimefon) or chlorothalonil. Combinations like pyraclostrobin and boscalid are

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especially effective on anthracnose diseases on ornamentals. Where available and known, plant resistant cultivars.

2.2

Botrytis Blight (Botrytis cinerea)

Geographic occurrence and impact. Botrytis blight on snapdragon has been reported in the USA (Horst 2013). It is most common on greenhouse-grown snapdragons, but occurs in field production of cut flowers and in the landscape. Symptoms/signs. Often the first symptom observed is wilting of flower spikes, a result of girdling of the stem in the region of the lowest flowers. The pathogen also causes a soft decay of flowers, stems, and seedlings which is accompanied by wilting (Fig. 1). On mature plants, most infections start in the flowers and progress downward. A gray mold consisting of mycelia and spores of B. cinerea covers diseased plant parts. Sclerotia (compact masses of mycelia) may form inside or on the surface of infected plants (Agrios 2005; Forsberg 1975; Nelson and Strider 1985; Pirone 1978). Biology and epidemiology. Botrytis blight is favored by high relative humidity and cool temperatures. The optimum temperature range for growth and sporulation of B. cinerea is 18–23  C/64–73  F. Free water on the plant surface is required for infection to occur. During favorable conditions, B. cinerea sporulates profusely, producing a mass of gray mycelium and spores. The fungus survives as mycelium or sclerotia in soil and plant debris. Spores are disseminated by air currents and will germinate and infect healthy plants if they land on wounds or senescent flowers. Insects such as moths, thrips, and fruit flies also can spread spores from infected to healthy plants (Forsberg 1975; Holz et al. 2004; Nelson and Strider 1985; Pirone 1978). Management. Scouting for detection of diseased plants coupled with sanitation and good cultural practices can be effective in reducing damage and losses caused by Fig. 1 Wilting (center) of snapdragon caused by Botrytis cinerea [A.R. Chase]

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B. cinerea. Removal and destruction of infected plants and infested plant debris can help to reduce the amount of inoculum. Good aeration keeps the plant surface dry, which prevents spores from germinating. In the greenhouse, humidity can be reduced by ventilation and heating. Use of resistant cultivars where available is an effective and inexpensive means of managing Botrytis blight. Application of chemical and biological fungicides was shown to be effective in controlling Botrytis blight on lisianthus (Wegulo and Vilchez 2006a) and is likely to be effective on snapdragon as well. Rotation is critical since resistance to many previously effective active ingredients has been common for Botrytis in ornamental crops (see Botrytis blight in chapter “▶ Diseases of Lisianthus”).

2.3

Cercospora Blight (Cercospora antirrhini)

Geographic occurrence and impact. Cercospora blight has been reported on snapdragon in Guatemala (Muller and Chupp 1950), the USA (Bolick 1959), and Argentina (Nelson and Strider 1985). The disease is capable of causing significant economic losses. Symptoms/signs. Symptoms of Cercospora blight on snapdragon were described by Jackson (1960). On leaves, lesions are subcircular and discrete to confluent with dull-white or gray centers surrounded by light-brown, slightly raised margins. They measure 0.5–7 mm in diameter. Under high humidity, rapid expansion of lesions may occur, resulting in irregular, poorly defined gray or tan necrotic areas measuring up to 15 mm in diameter with occasional faint concentric zones. The fungus sporulates profusely in the centers of lesions. Heavily infected leaves become chlorotic and drop. On stems, lesions occur mostly at the base near the soil line. They are depressed and dull white to gray with brown margins. Initially they are discrete, subcircular, oval, or elliptical. Eventually they become confluent, measuring up to 4 cm long. Cortical tissue on the stem becomes necrotic and small longitudinal cracks develop in it. Sporulation is sparse in stem lesions. Infected plants with leaf and stem lesions are chlorotic and stunted. Biology and epidemiology. Conidia are disseminated by air currents, wind, or splashing water. Free water is necessary for spore germination and infection. Incubation period in greenhouse- and field-grown snapdragons inoculated at 8 weeks of age with a suspension of conidia and mycelial fragments was 21–30 days at an incubation temperature range of 16–32  C/61–90  F. Disease development was favored by warm temperatures (30  C/86  F) and wetness. The pathogen survived in dry leaf and stem tissues for at least 14 months and in leaf debris or in the soil for at least 3 months (Porter and Aycock 1967). Because seed transmission is common in Cercospora spp. plant host systems, it is possible that C. antirrhini can be seed transmitted. Farr et al. (1989) list Antirrhinum as the only host of C. antirrhini. Management. Remove and destroy infected plants or plant parts. In the field or landscape, burying plant debris by plowing will reduce inoculum. Rotate with non-host crops. Use pathogen-free seed. Avoid overhead irrigation. Space plants to allow adequate air circulation. In the greenhouse, keep humidity low through

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ventilation and aeration. Integrate these cultural tactics with spraying fungicides labeled for leaf spot control on ornamental crops. Thiophanate-methyl remains an excellent rotational choice in addition to strobilurins, triazoles, chlorothalonil, and fludioxonil.

2.4

Downy Mildew (Peronospora antirrhini)

Geographic occurrence and impact. Downy mildew on snapdragon has been reported in Ireland (Murphy 1937), England (Green 1937), Australia (Anon 1941), Italy (Garibaldi and Rapetti 1981), and the USA (Harris 1939; Kirby 1945; Yarwood 1947). The disease can be very destructive both in greenhouse- and field-grown snapdragons. Total loss can result from the disease (Yarwood 1947). Fig. 2 Pale green appearance on the upper surface of a downy mildew-infected snapdragon leaf [S. N. Wegulo]

Fig. 3 The lower surface of a downy mildew-infected snapdragon leaf showing a gray to white fungal growth [S. N. Wegulo]

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Symptoms/signs. Downy mildew affects primarily seedlings, but snapdragons can be infected at all growth stages. On seedlings, a downward curling of leaves and a reduction in the size of the plant and leaves occur. Infected leaves appear pale green on the upper surface and have a downy gray, lavender to white fungal growth on the lower surface (Figs. 2 and 3). This fungal growth can also occur on stems and upper leaf surfaces of young succulent plants. Systemically infected plants can be severely stunted, wilt, and eventually die. Seedling death progresses from the top down to the soil surface. On larger plants, symptoms of systemic infection include stunting (Fig. 4), pale green leaves (Fig. 5), and lack of flowering. Systemic infection can result in rosetting of growing points. Commonly, the shoots die and infected plants produce many secondary shoots from the base. Biology and epidemiology. Snapdragon downy mildew is favored by high relative humidity and cool, wet conditions. Optimal temperatures for disease

Fig. 4 Stunting and wilting of a snapdragon plant (frontcenter) due to systemic infection by downy mildew [S. N. Wegulo]

Fig. 5 A snapdragon plant infected with downy mildew. Note the pale green color on the upper surfaces of leaves [S. N. Wegulo]

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development range from 5  C to 21  C/41 to 70  F. Local infection is characterized by pale round areas on leaves and is rarely destructive. Systemic infection can be very destructive especially on seedlings. The incubation period can be as short as 4 days. Environmental conditions during the incubation period are most critical in determining how much infection occurs in a snapdragon planting or seedling tray (Yarwood 1947). P. antirrhini sporulates in cool, humid conditions. During sporulation, examination of the underside of infected leaves reveals a grayish downy growth consisting of sporangiophores and sporangia (spores). Sporangia are spread by wind, air currents, or splashing water. When they land on healthy plants, they germinate by means of a germ tube and cause new infections. In Michigan, Byrne et al. (2005) found that in field-grown snapdragons, dew periods of 6 h or longer were associated with large releases of sporangia, whereas consecutive days with short leaf wetness periods were associated with low sporangial concentrations in the atmosphere. Thick-walled resting spores (oospores) form abundantly in the roots, stems, and petioles of systemically infected plants and are the means by which the pathogen survives during unfavorable conditions. The host range of P. antirrhini is limited to Antirrhinum (Farr et al. 1989). Management. Keep relative humidity in the greenhouse below 85 % by balancing heat and ventilation. Keeping leaves dry using fans can be effective in preventing infections, but can also increase disease spread because spores can be disseminated by air currents. Avoid overhead irrigation. Thoroughly inspect all seedlings or plug trays for downy mildew before transplanting. Remove and destroy infected plants taking care not to spread the spores. After handling infected plants, thoroughly wash hands with soap before handling healthy plants. Some snapdragon cultivars have been shown in tests to have good resistance to snapdragon downy mildew (Byrne et al. 2004). Fungicides have been shown to be effective in controlling snapdragon downy mildew. Wegulo and Vilchez (2006b) found fenamidone, mancozeb, fosetyl-Al, and dimethomorph to have very good to excellent control of snapdragon downy mildew. To achieve effective control, apply fungicides preventively. The best management strategy for downy mildew is to integrate as many management tactics as practically and economically possible.

2.5

Phyllosticta Blight (Phyllosticta antirrhini)

Geographic occurrence and impact. Phyllosticta blight, also known as Phyllosticta leaf spot, has been reported in the USA (Guba and Anderson 1919). It is damaging primarily on snapdragons grown outdoors, but can also occur in the greenhouse (Nelson and Strider 1985). Symptoms/signs. The pathogen can attack snapdragons at any growth stage. On leaves, initial symptoms are chlorotic spots that form green to black lesions. They enlarge and form yellow to brown concentric rings. The spots are visible on both sides of the leaf and become cream colored and slightly sunken in the center (Fig. 6). In wet weather, the spots continue to enlarge until the entire leaf collapses. Infection of young leaves causes them to become distorted and curled, and they may shrivel

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Fig. 6 Phyllosticta leaf spot on snapdragon [A.R. Chase]

and die but remain clinging to the stem. Lesions on petioles are brown and elongated and may girdle the petiole causing the leaf to droop and die. From the petioles, lesions spread to the stem and cause infections at the leaf axil. Pinpoint size black pycnidia form in diseased tissues. On stems, lesions are dark green to brown with no definite margins. They may elongate up to 3 cm and girdle the stem. They turn dark brown, brittle, and dry and pycnidia eventually form in them. Damping-off occurs as a result of rapid wilting and dying of all aboveground parts of the plant (Forsberg 1975; Guba and Anderson 1919; Nelson and Strider 1985). Biology and epidemiology. The pathogen overwinters as pycnidia on host debris. Moisture is required for conidial germination and infection. Conidia released from pycnidia are dispersed by splashing water. Spore germination was best at 25  C/77  F (Guba and Anderson 1919), indicating that moderate to warm temperatures favor disease development. Disease development continues until the plant dies under wet conditions but is arrested or considerably slowed under dry conditions. In greenhouse infection experiments, lesions appeared on unwounded and wounded leaves of snapdragon plants 9 and 5 days after spray inoculation with conidia, respectively, indicating that wounding is not necessary for infection. Pycnidia began to appear on the fourth or fifth day after lesion appearance (Guba

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and Anderson 1919). In addition to Antirrhinum, Penstemon is also a host of P. antirrhini (Farr et al. 1989). Management. Remove and destroy infected plants as soon as they are noticed. Remove and destroy plant debris at the end of the growing season. Avoid excessive and overhead watering. Space plants to allow good air circulation. Apply fungicides labeled for control of leaf spots on ornamental crops.

2.6

Phytophthora Stem Rot and Wilt (Phytophthora cactorum, P. cryptogea, P. parasitica)

Geographic occurrence and impact. Phytophthora stem rot and wilt has been reported in the USA (Gill 1960; Harris 1934). Gill (1960) cited reports of the disease in other countries including Rhodesia (now Zimbabwe), Madagascar, Mauritius, England, South Africa, India, and Australia. Losses of 50 % or more in greenhousegrown snapdragons were reported by Gill (1960), indicating the destructive nature of the disease. Symptoms/signs. Symptoms of the disease as observed by Harris (1934) include wilting caused by girdling of the stem at or slightly above the soil line. Lesions initially appear as water-soaked areas on healthy white stem tissue. As the lesions enlarge, their older portions become yellow, brown, and eventually almost black. They enlarge, extending up and down, until the stem is girdled (Fig. 7). The outer portion of the stem may slough off exposing the hard woody xylem tissue. When initially infected, plants may wilt slightly during the day and recover at night. Severely affected plants wilt permanently within 2–3 days and later become dry and brown. Under humid and wet conditions, lesions may extend up the main stem onto the side branches. Biology and epidemiology. Phytophthora spp. have a wide host range. They survive as oospores, chlamydospores, or mycelium in infected plant tissues or in soil. Long-term survival is by means of oospores and chlamydospores. Under Fig. 7 Phytophthora stem rot and wilt on snapdragon [A.R. Chase]

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favorable conditions, oospores and chlamydospores germinate by means of zoospores, whereas mycelium grows and forms sporangia which release zoospores. The zoospores swim in water and infect roots or are splash dispersed onto above-ground plant parts where they cause infections. Development of Phytophthora stem rot and wilt of snapdragon is favored by moisture and moderate to warm temperatures (Gill 1960). Management. Harris (1934) observed differences in susceptibility of three snapdragon cultivars to Phytophthora stem rot and wilt, with disease incidence ranging from 20 % in Cheviot Maid and 30 % in Jenny Schneider to 75 % in Roman Gold. Plant snapdragon cultivars are known to have good resistance to Phytophthora stem rot and wilt. Because the disease is favored by wetness, ensure good soil drainage. Avoid overhead watering. Keep plant surfaces dry by maintaining sufficient air circulation. Apply fungicides labeled for Phytophthora control on ornamental crops. Hausbeck and Glaspie (2009a, b, c) found the fungicides mandipropamid, fluopicolide, fenamidone, mefenoxam, and mono- and dipotassium salts of phosphorous acid to be effective in controlling Phytophthora root rot of snapdragons when applied as drenches.

2.7

Powdery Mildew (Golovinomyces orontii (Formerly Erysiphe cichoracearum); Oidium spp.)

Geographic occurrence and impact. Powdery mildew on snapdragon has been reported in England (Moore 1947) and the USA (Guba 1936). Symptoms/signs. A powdery, white fungal growth appears on both surfaces of leaves, but mostly on the upper surface. Plants can be infected at any growth stage. Infections usually begin on the lower leaves, but can become severe on the upper leaves, stems, and flowers. Severely infected leaves may wilt and die. Biology and epidemiology. Powdery mildew fungi are obligate parasites. Fungal growth occurs on the plant surface and is nourished by haustoria (special absorption structures that form in epidermal cells following infection). The fungal growth consists of mycelia and conidia which are dispersed by wind or air currents. When the conidia land on healthy plants, they germinate and cause new infections. Cleistothecia form in older infected plant tissues and produce ascospores. Disease development is favored by shade and high relative humidity but not free water. These fungi typically have a very narrow host range and survive as mycelium in infected plants or as cleistothecia in plant debris. Management. Space plants to allow good air circulation. Avoid planting snapdragons in shaded areas. In the greenhouse, reduce relative humidity by heating and venting. Apply fungicides labeled for powdery mildew control on ornamental crops. In trials conducted by Raabe et al. (1970), the systemic fungicide benomyl achieved complete control of powdery mildew on snapdragons when applied as a spray or a drench. Fungicides are most effective when applied preventively and especially include triazoles and strobilurins.

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Fig. 8 Wilting caused by Pythium root rot [A.R. Chase]

2.8

Pythium Root Rot (Pythium spp.)

Geographic occurrence and impact. Pythium root rot on snapdragons has been reported in the USA (Hanan et al. 1962). Symptoms/signs. Snapdragons can be attacked at any growth stage. Infection of roots in the root hair zone and colonization of cortical tissues causes a brown, watersoaked rot. Basal stem tissues may also rot; however, stem tissues above the soil line are rarely colonized. Pre- or postemergence damping-off of seedlings may occur. Infected seedlings that survive and are transplanted and young plants infected after transplanting are stunted, chlorotic and wilt when exposed to sunlight. Infected plants have a reduced root system. Wilt collapse is a common symptom observed in mature plants (Fig. 8) especially at flowering when water demand cannot be met by the reduced root system. Older plants tend to be more resistant to symptom development than younger plants. In experiments conducted by Mellano et al. (1970), 15-day-old or younger seedlings infected with Pythium ultimum died within 6 days due to rapid and unrestricted colonization of host tissue. In contrast, 25-day-old plants were tolerant to infection and host colonization. Wilting and stunting intensified by high temperature was observed in infected tolerant plants. Biology and epidemiology. Pythium spp. have a wide host range. They live in the soil and produce a white mycelium that grows rapidly and gives rise to sporangia. A sporangium germinates directly by giving rise to a germ tube or by producing a short hypha bearing a balloon-like vesicle at its end. Within the vesicle, 100 or more zoospores are produced. When released from the vesicle, the zoospores move around in a swarm for a few minutes then form cysts which germinate by giving rise to a germ tube. The germ tube infects host tissue by direct penetration. The mycelium can also produce a female oogonium and a male antheridium. The antheridium produces a fertilization tube which enters the oogonium. Nuclei of the antheridium move through the tube and fuse with nuclei of the oogonium to form a zygote which becomes an oospore by producing a thick wall around itself. The oospore is resistant

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to adverse environmental conditions and is the survival and resting stage of Pythium spp. When environmental conditions become favorable, oospores germinate in a manner similar to that of sporangia by directly forming a germ tube or by first forming zoospores and cysts. Rootlets can be infected at any stage of plant growth. Root rot progresses upward to the crown and stem. Disease development is favored by wet and waterlogged soils (Agrios 2005). Zoospores are disseminated by flowing water or splash dispersal. In the greenhouse, the primary means of introducing Pythium propagules are contaminated plant material and soiled hands, tools, and hose ends (Daughtrey et al. 1995). Management. Cultural methods of control include good water drainage. If the planting site is poorly drained, plant snapdragons in raised beds. Composted bark can be incorporated into the soil in the raised bed to improve aeration. Drainage tiles can be used to facilitate drainage and direct water away from the planting site. Do not reuse pots or trays from a previous crop for propagation. If pots or trays are reused, they should sanitized or sterilized first. Fungicides registered for Pythium control such as sodium, potassium and ammonium phosphites, etridiazole + thiophanatemethyl, mefenoxam, and etridiazole can be used to control root rot in snapdragons (Pscheidt and Ocamb 2016). Del Castillo Múnera and Hausbeck (2015) found that the efficacy of fungicides in controlling Pythium root rot on snapdragons depended on the species Pythium. Mefenoxam and fenamidone reduced root rot caused by P. aphanidermatum and P. ultimum; whereas fluopicolide and etridiazole reduced root rot caused by P. ultimum; and fenamidone, potassium phosphite, and the biological control agent Trichoderma harzianum effectively controlled root rot caused by P. irregulare. In the same study, differences in intensity of root rot caused by Pythium spp. were observed among snapdragon cultivars, although none of the cultivars was highly resistant.

2.9

Rhizoctonia Basal Stem Rot (Rhizoctonia solani)

Geographic occurrence and impact. Rhizoctonia basal stem rot on snapdragon has been reported in the USA (Baker and Sciaroni 1952) and Taiwan (Yang and Leu 1981). Symptoms/signs. Snapdragons can be infected at any growth stage. They are most susceptible at the seedling stage and immediately following transplanting (Nelson and Strider 1985). Symptoms include pre- and postemergence dampingoff. Infected seedlings that survive have red-brown lesions on the basal stem at the soil line. Wilting, collapse, and death occur if lesions girdle the stem. Affected tissues show a coarse brown mycelium of R. solani in wet and humid environments. Biology and epidemiology. R. solani has a wide host range. It survives as mycelium or sclerotia in infected plants or plant parts, soil, or plant debris. It is spread by water, contaminated tools, or infected plants or propagative plant material. Basal stem rot on snapdragon is favored by high temperatures and moderately moist soils (Baker and Sciaroni 1952).

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Management. Treat soil with heat (steam or solarization), fumigants, or fungicides labeled for control of Rhizoctonia in ornamental crops. Ensure good soil drainage and avoid excessive watering. Plant snapdragons on raised beds if the area is poorly drained. Use sterilized soil, trays, and tools to raise seedlings before transplanting. Transplant disease-free seedlings. In the greenhouse, keep relative humidity low by venting and heating. Space plants to allow good air circulation. Some of the most effective fungicides include fludioxonil, thiophanate-methyl, and strobilurins (like azoxystrobin and pyraclostrobin).

2.10

Rust (Puccinia antirrhini)

Geographic occurrence and impact. Snapdragon rust was first reported in the USA by Blasdale (1903) who had found it at Berkeley, CA, in 1895 (McClellan 1953). Peltier (1919) noted that the disease caused much loss in Illinois in the 4 years following its discovery in the state in 1913. McClellan (1953) echoed the seriousness of the disease by stating that “Of the numerous diseases of snapdragons, rust probably causes the most concern.” Snapdragon rust occurs throughout the world wherever snapdragons are grown (Gawthrop and Brooks 1979; Nelson and Strider 1985). Symptoms/signs. Seedlings, cuttings, and mature plants can be attacked by rust in the greenhouse as well as in the field. It is most severe on cuttings and on plants just before flowering (Peltier 1919). All above-ground parts of the plant except the florets can be infected. Initial symptoms on leaves are small, chlorotic swellings. These swellings increase in size until the epidermis ruptures, exposing red-brown uredinial pustules. Pustules are limited to the lower leaf surface. On the upper surface of the leaf above each pustule, a circular yellow area is apparent. The fungus continues to grow under favorable environmental conditions and in 4 or 5 days Fig. 9 Rust pustules on the undersides of snapdragon leaves [A.R. Chase]

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forms one or two concentric rings of smaller secondary pustules around the original pustule (Nelson 1962) (Fig. 9). In the presence of moisture and high relative humidity, the pustules are invaded by secondary microorganisms, mainly Fusarium spp., which cause irregularly shaped necrotic spots around the pustule (Dimock and Baker 1951). On stems, initial small pustules elongate, causing swollen cankers which serve as entry points for secondary invaders. These invaders cause lesions that girdle the stem, resulting in wilting and plant death. Severely affected plants have a scorched or brown appearance. Flowers open prematurely and are smaller than normal. Biology and epidemiology. The host range of P. antirrhini is limited to Antirrhinum (Farr et al. 1989). Like other rust fungi, it is an obligate parasite. Urediniospores are spread by wind over long distances and by wind-blown rain or splashing water over short distances (Nelson and Strider 1985). They land on healthy plants and initiate new infections. Optimal conditions for infection are the presence of free water and a temperature range of 10–13  C/50–55  F. Disease development is optimal under a temperature range of 21–24  C/70–75  F and is slowed at higher temperatures (Dimock and Baker 1951). Under optimal conditions for disease development, new uredinia are formed within 8–16 days following inoculation (Peltier 1919), and the cycle of spore production is repeated. The urediniospore is the repeating spore of P. antirrhini. Teliospores form later in the growing season, but do not play an important role in the disease cycle. Under semiarid conditions, injury is primarily from desiccation through the rust pustules. In the presence of moisture and high relative humidity, severe injury is caused by secondary invaders, primarily Fusarium spp. (Dimock and Baker 1951). There are no known alternate hosts of P. antirrhini in the USA. The fungus survives as mycelium or urediniospores on snapdragon plants in the greenhouse or field. Management. Cuttings are extremely susceptible (Peltier 1919). Therefore, only cuttings from rust-free snapdragons should be used. Otherwise start plants from seed. Allow good drainage and good air circulation. Keep temperatures in the greenhouse above 24  C/75  C for several days and not below 16  C/61  F at night (Pscheidt and Ocamb 2016). Avoid overhead watering. Integrate the above cultural management tactics with application of fungicides registered for rust control on ornamental crops. Fungicides shown to provide good results on snapdragon rust include strobilurins (especially azoxystrobin) and triazoles which can show eradicant benefits at times. Mancozeb is also excellent but only as a true preventive application.

2.11

Verticillium Wilt (Verticillium albo-atrum, V. dahliae)

Geographic occurrence and impact. Verticillium wilt on snapdragons has been reported in the USA (Baker and Sciaroni 1952). Symptoms/signs. The pathogens infect roots and colonize the xylem vessels, inhibiting water movement. Initially the lower leaves may become yellow, wilt, and drop. Later wilting and collapse of single branches may occur, causing the plant to appear as if wilting on one side (Nelson and Strider 1985; UC IPM 2014). The entire

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plant finally wilts, collapses, and dies. During periods when cool days or nights alternate with warm days, wilting and plant death can occur rapidly. Wilting often occurs after the onset of blossom development. Vascular tissues are discolored continuously from the base of the stem to the top of the plant. Black microsclerotia may form in diseased tissue. Biology and epidemiology. Verticillium spp. are soilborne and have a wide host range. They survive over long periods of time as microsclerotia in the soil or plant debris. Introduction into non-infested fields or greenhouses is through microslerotiacontaminated soil or tools and mycelium in infected plant material such as cuttings. Disease development is favored by moisture and cool temperatures. Dutta (1981) showed that the severity of Verticillium wilt on snapdragon was higher in alkaline compared to acidic soil. Organic (chitin and green manure) and inorganic (ammonium sulfate, calcium nitrate, and combined NPK) soil amendments reduced disease severity, and this was attributed to boosting antagonistic microorganisms in the soil and direct nutritional effects on snapdragon plants (Dutta and Isaac 1979a, b; Isaac 1956). Excessive nitrogen fertilization increased disease severity (Isaac 1957). Management. Treat soil with heat (steam or solarization) or fumigants before planting or transplanting snapdragons. In the greenhouse, sanitize trays, tools, and benches. Remove and destroy infected plants and infested plant debris. Use diseasefree plants or cuttings. Avoid planting snapdragons in poorly drained soils.

2.12

Additional Fungal Diseases and Pathogens

The following fungal diseases and pathogens have been reported on snapdragon (Horst 2013): southern blight (stem rot, Sclerotium rolfsii), stem and crown canker (Myrothecium roridum), collar rot (Rhizoctonia solani), petal rot (Bipolaris setariae), charcoal rot (Macrophomina phaseoli), root rot (Phymatotrichum omnivorum, Thielaviopsis basicola), and stem rot and wilt (Fusarium sp., Sclerotinia sclerotiorum, S. minor). In addition, seed blight (Alternaria alternata) has been reported (Harman et al. 1973).

3

Bacterial Diseases

3.1

Seedling Blight (Pseudomonas antirrhini)

Geographic impact and occurrence. Seedling blight on snapdragon has been reported in Japan (Takimoto 1920), Australia (Valder 1963), and the United Kingdom (Simpson et al. 1971). Chase (2001, 2004) listed Pseudomonas as a causal agent of Pseudomonas leaf spot on certain cut flower cultivars of snapdragon in the USA (Fig. 10), but did not specify the species. Symptoms/signs. The disease is actually a leaf spot, but when severe it is better described as a seedling blight (Simpson et al. 1971). Leaf lesions are initially small, pale green, and less than 0.5 mm in diameter. They enlarge and become brown with a

Diseases of Snapdragon

17

Fig. 10 Seedling blight caused by Pseudomonas spp. [A.R. Chase]

green halo. When fully developed, they are discrete, almost circular, and measure 4–5 mm in diameter. They appear sunken and papery brown with a defined dark brown margin. Water-soaked dark green zones may be seen around the lesions. As disease development progresses, the lesions coalesce, forming irregular blotches followed by leaf collapse. On dark-leaved cultivars, lesions are less discrete and the tissue surrounding the lesions may appear chlorotic (Simpson et al. 1971). Biology and epidemiology. Simpson et al. (1971) observed that seedling blight appeared on snapdragon seedlings usually after the first transplanting and before reaching a height of 15–18 cm. Beyond this growth stage, the disease did not attack newly emerged foliage. Disease development during the susceptible growth stages is favored by moisture such as that from mist irrigation during propagation. The pathogen is spread by splashing water from rain or irrigation. Pathogenicity tests on 6 to 8-cm-tall seedlings showed that the incubation period was 10–12 days. Injury to leaf surfaces by an abrasive agent resulted in severe symptoms (Simpson et al. 1971). Seed transmission studies confirmed that P. antirrhini can be transmitted through seed. The host range of this pathogen is limited to members of the family Scrophulariaceae, of which snapdragon is the most susceptible. Management. Use pathogen-free seed and propagative plant material. Avoid excessive moisture on the foliage of seedlings. Remove and destroy infected seedlings. Plant resistant cultivars where known and available. Preventively apply bactericides labeled for disease control on ornamental crops. Bactericides can only prevent or reduce infections; they cannot eliminate established infections.

3.2

Additional Bacterial Diseases and Pathogens

Crown gall (Agrobacterium tumefaciens) has been reported on snapdragon (Horst 2013).

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Fig. 11 Necrosis on snapdragon caused by impatiens necrotic spot virus (INSV) [A.R. Chase]

4

Viral Diseases

4.1

Impatiens Necrotic Spot Virus (INSV) and Tomato Spotted Wilt Virus (TSWV)

Geographic impact and occurrence. Impatiens necrotic spot virus has been reported on snapdragon in the USA (Daughtrey et al. 1997; Horst 2013). Significant losses can result from infection by these viruses because symptomatic plants are not marketable. Symptoms/signs. Symptoms of INSV on snapdragons can be variable and include brown to white spots on leaves, stunting, stem necrosis which causes tissues to turn black (Fig. 11), and plant death. Because symptoms are variable and resemble those caused by other diseases, they are not reliable for diagnosis. Confirmation is done by performing laboratory tests such as enzyme-linked immunosorbent assay (ELISA) or polymerase chain reaction (PCR). Biology and epidemiology. INSV is one of two viruses that infect a wide range of plant species including ornamentals, vegetables, and field crops. The other virus is tomato spotted wilt virus (TSWV). Both belong to the genus Tospovirus in the family Bunyaviridae and are transmitted by the western flower thrip (Daughtrey et al. 1997). Spread of these viruses is through movement of infected plants or viruliferous thrips which feed on and transmit the viruses to healthy plants. Many plant hosts can be infected but show no symptoms. These plant hosts which include weeds serve as reservoirs for the viruses and their vectors. Management. Effective control can be achieved by virus exclusion and prevention of virus spread. Virus exclusion involves use of virus-indexed propagation material which can be purchased from certain suppliers. INSV and TSWV can also be excluded by preventing thrips from entering the greenhouse or propagation and storage areas. This can be achieved through the use of fine mesh screens. Weeds and

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19

other thrips and virus plant hosts growing in the vicinity of greenhouses should be controlled with herbicides or other means. Workers should take care not to introduce thrips into the greenhouse or propagation areas by changing clothes before entry and by not wearing clothes with colors such as yellow, green, or blue which attract thrips. Prevention of virus spread is achieved by controlling resident thrip populations with insecticides or biocontrol agents and by roguing and disposing of infected plants. Because virus-infected plants are more susceptible to the feeding and oviposition of thrips (Bautista et al. 1995), such plants should be disposed of as quickly as they are noticed.

4.2

Additional Virus Diseases

Cucumber mosaic virus, an unidentified mosaic virus, and an unidentified ring spot virus have been reported on snapdragon (Horst 2013).

5

Nematode Diseases

5.1

Root-Knot Nematode (Meloidogyne spp.)

Geographic occurrence and impact. Root-knot nematode has been reported on snapdragon in the USA (Horst 2013; Pirone 1978). Symptoms/signs. Root galling and stunting. Biology and epidemiology. Root-knot nematodes have a wide host range. They feed and reproduce within plant roots, inducing small to large galls known as rootknots. The adult females are sedentary in plant roots and have a round to pear-shaped body. Adult males are vermiform, leave the root, and become free-living in the soil. Adult females lay eggs in gelatinous sacs that are deposited on the surface of galled roots or may remain within the galls. The egg develops into a vermiform first-stage juvenile (J1) which molts into a vermiform second-stage juvenile (J2) that emerges from the egg into the soil. If the J2 (the only infective stage) encounters a susceptible host, it enters the root and becomes sedentary. It feeds on the root cells surrounding its head and grows, becoming sausage shaped. The saliva it secretes causes the root cells to enlarge and form galls. The J2 molts into a third-stage juvenile (J3) which molts into a fourth-stage juvenile (J4). Both the J3 and J4 are sedentary within the root. The J4 undergoes a final molt into a mature male which leaves the root or a mature female which remains sedentary in the root and lays eggs. Root-knot nematodes are spread by contaminated equipment, infected plant material, or water. Nematodes attack ornamental plants in the landscape or field; they do not seem to be present in greenhouse- or container-grown crops (Creswell 2009; Ismail 1980; McSorley 1994; McSorley and Fredeik 1994). Management. Sanitize equipment after use. Contaminated soil can be disinfested by fumigation, drenching with nematicides, or heat treatment with steam or

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solarization. Refer to chapter “▶ Soil/Media Disinfestation for Management of Florists’ Crops Diseases” for specific strategies.

5.2

Additional Nematode Pathogens

The lesion nematodes Pratylenchus spp. have been reported on snapdragon (Baker and Sciaroni 1952; Horst 2013).

6

Additional Parasites

Dodder (Cuscuta sp.) has been reported on snapdragon (Horst 2013).

7

Nonparasitic Disorders

7.1

Tip Blight

A tip blight of unknown cause has been described on snapdragons (Forsberg 1975; Horst 2013). Leaves gradually wilt starting at the plant tip and the wilt progresses to affect petioles and stems. Sunken, water-soaked lesions may form on the stem and

Fig. 12 Phytotoxicity (second and third leaves from left) caused by an experimental fungicide on snapdragon. The first, fourth, and fifth leaves from left are from plots sprayed with commercial fungicides. The sixth leaf is from a non-sprayed check plot. [S. N. Wegulo]

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girdle it, causing the plant tip to wilt and die. This disorder usually occurs in winter when it is cloudy and disappears in spring when brighter weather returns.

7.2

Phytotoxicity

Certain pesticides may cause injury to snapdragons. Figure 12 shows injury to snapdragon leaves following application of an experimental fungicide.

7.3

Additional Nonparasitic Disorders

Fasciation has been reported on snapdragon. It is probably a genetic abnormality (Horst 2013).

References Agrios GN (2005) Plant pathology, 5th edn. Elsevier Academic Press, New York Anonymous (1941) Downy mildew of snapdragons. Agric Gaz N S W 52:538–539 Baker KF, Sciaroni RH (1952) Diseases of major floricultural crops in California. California State Florists’ Association, Los Angeles Bautista RC, Mau RFL, Cho JJ, Custer DM (1995) Potential of tomato spotted wilt tospovirus plant hosts in Hawaii as virus reservoirs for transmission by Frankliniella occidentalis (Thysanoptera: Thripidae). Phytopathology 58:953–958 Blasdale WC (1903) On a rust of the cultivated snapdragon. J Mycol 9:81–82 Bolick JH (1959) Cercospora antirrhini found in Florida. Plant Dis Rep 43:511 Byrne JM, Sconyers LE, Hausbeck MK (2004) Evaluation of snapdragon cultivars for resistance to downy mildew, 2000 and 2001. B&C Tests 19:O010 Byrne JM, Hausbeck MK, Sconyers LE (2005) Influence of environment on atmospheric concentrations of Peronospora antirrhini sporangia in field-grown snapdragon. Plant Dis 89 (10):1060–1066 Chase AR (2001) Controlling bacterial diseases of ornamentals. Southeast Floriculture, July/ Aug:20–21 Chase AR (2004) Bacterial diseases: are we losing the battle? GPN Mag, April 2004:32–35 Creel R, Raymond Kessler J (2007) Greenhouse production of bedding plant snapdragons. Alabama Cooperative Extension System. Publication No. ANR-1312 Creswell T (2009) Nematode management in bedding plants in the landscape. NCSU. Ornamental Disease Note 31 Daughtrey ML, Wick RL, Peterson JL (1995) Compendium of flowering potted plant diseases. APS Press, St. Paul Daughtrey ML, Jones RK, Moyer JW, Daub ME, Baker JR (1997) Tospoviruses strike the greenhouse industry. Plant Dis 81(11):1220–1230 Del Castillo Múnera J, Hausbeck MK (2015) Integrating host resistance and plant protectants to manage Pythium root rot on geranium and snapdragon. HortScience 50(9):1319–1326 Dimock AW, Baker KF (1951) Effect of climate on disease development, injuriousness, and fungicidal control, as exemplified by snapdragon rust. Phytopathology 41:536–552

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Dutta BK (1981) Effect of the chemical and physical condition of the soil on Verticillium wilt of Antirrhinum. Plant and Soil 63(2):217–225 Dutta BK, Isaac I (1979a) Effects of inorganic amendments (N, P and K) to soil on the rhizosphere microflora of Antirrhinum plants infected with Verticillium dahliae Kleb. Plant and Soil 52 (4):561–569 Dutta BK, Isaac I (1979b) Effects of organic amendments to soil on the rhizosphere microflora of Antirrhinum infected with Verticillium dahliae Kleb. Plant and Soil 53(1):99–103 Farr DF, Bills GF, Chamuris GP, Rossman AY (1989) Fungi on plants and plant products in the United States. Univeristy of Illinois: APS Press, St. Paul Forsberg JL (1975) Diseases of ornamental plants. Special publication no. 3 revised, University of Illinois: Urbana-Champaign Garibaldi A, Rapetti S (1981) Grave epidemia di peronospora su antirrino [Antirrhinum majus L.] (Downy mildew attacks on Antirrhinum majus L.). Colture Protette 11:35–38 Gawthrop F, Brooks A (1979) The menace of Antirrhinum rust. Garden J Royal Hortic Soc 104:68–70 Gill DL (1960) A stem and branch rot of snapdragon. Plant Dis Rep 44(12):946–947 Green DE (1937) Downy mildew on Antirrhinum majus: a disease new to Great Britain. Gard Chron 102:27–28 Guba EF (1936) Plant disease notes from Massachusetts. Plant Dis Rep 20:302–303 Guba EF, Anderson PJ (1919) Phyllosticta leaf spot and damping off of snapdragons. Phytopathology 9:315–325 Hanan JJ, Langhans RW, Dimock AW (1962) Soil aeration and the Pythium root rot disease of snapdragon. Bull N Y State Flower Grow 195:1–6 Harman GE, Heit CE, Pfleger FL, Braverman SW (1973) Snapdragon seed blight – a serious problem caused by seedborne fungi. Plant Dis Rep 57(7):592–595 Harris MR (1934) A Phytophthora disease of snapdragons. Phytopathology 24:412–417 Harris MR (1939) Downy mildew on snapdragon in California. Plant Dis Rep 23:16 Hausbeck MK, Glaspie SL (2009a) Residual control of Phytophthora root rot of snapdragons, 2007. Plant Disease Management Reports 3:OT014 Hausbeck MK, Glaspie SL (2009b) Control of Phytophthora root rot of dwarf snapdragons with fungicide drenches, 2008. Plant Disease Management Reports 3:OT015 Hausbeck MK, Glaspie SL (2009c) Control of Phytophthora root rot of cut flower snapdragons with fungicide drenches, 2008. Plant Disease Management Reports 3:OT014 Holz G, Coertze S, Williamson B (2004) The ecology of Botrytis on plant surfaces. In: Elad Y, Williamson B, Tudzynski P, Nelson N (eds) Botrytis: biology, pathology and control. Kluwer, London, pp 9–27 Horst RK (2013) Field manual of diseases on garden and greenhouse flowers. Springer, New York Isaac I (1956) Some factors affecting Verticillium wilt of Antirrhinum. Ann Appl Biol 44:105–112 Isaac I (1957) The effects of nitrogen supply upon the Verticillium wilt of Antirrhinum. Ann Appl Biol 45:512–515 Ismail W (1980) Susceptibility of some ornamental plants to the attack of Meloidogyne incognita. Indian J Hortic 37:326–328 Jackson CR (1960) Cercospora blight of snapdragon. Phytopathology 50:190–192 Jerardo A (2007) Floriculture and nursery crops yearbook/FLO-2007. Economic Research Service, United States Department of Agriculture Kirby RS (1945) Downy mildew on snapdragon in Pennsylvania. Plant Dis Rep 29:371 Maree J, Wyk B-E (2010) Cut flowers of the world: a complete reference for growers and florists. Timber Press, Portland McClellan WD (1953) Rust and other disorders of snapdragon. In: Yearbook of agriculture 1953. United States Department of Agriculture, Washington, DC, pp 568–572 McSorley R (1994) Susceptibility of common bedding plants to root-knot nematodes. Proc Fla State Hortic Soc 107:430–432

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McSorley R, Fredeik JJ (1994) Response of some common annual bedding plants to three species of Meloidogyne. J Nematol 26(4S):773–777 Mellano HM, Munnecke DE, Endo RM (1970) Relationship of seedling age to development of Pythium ultimum on roots of Antirrhinum majus. Phytopathology 60:935–942 Moore WC (1947) British fungi. Trans Br Mycol Soc 31:86–91 Muller AS, Chupp C (1950) Cercospora in Guatemala. Ceiba 1:171–178 Murphy PA (1937) Irish Free State: a new outbreak of Peronospora antirrhini in the country. Int Bull Plant Prot 11:176 Nelson P (1962) Diseases. In: Langhans RW (ed) Snapdragons, a manual of the culture, insects and diseases and economics of snapdragons. New York State Flower Growers Association, Ithaca, pp 70–80 Nelson PE, Strider DL (1985) Snapdragons. In: Strider DL (ed) Diseases of floral crops, vol 2. Praeger Publishers, New York, pp 489–510 Peltier GL (1919) Snapdragon rust. Ill Agric Exp Sta Bull 221:535–548 Pirone PP (1978) Diseases and pests of ornamental plants. Wiley, New York Porter DM, Aycock R (1967) Snapdragon leaf spot caused by Cercospora antirrhini. NC Agric Exp Sta Tech Bull No. 179 Pscheidt JW, Ocamb CM (2016) Pacific northwest plant disease management handbook. Oregon State University. http://pnwhandbooks.org/plantdisease/ Raabe RD, Hurlimann JH, Sciaroni RH (1970) Powdery mildew control with benomyl for greenhouse-grown snapdragons. Calif Agric 24(1):8 Rogers MN (1992) Snapdragons. In: Larson RA (ed) Introduction to floriculture, 2nd edn. Academic, New York, pp 93–112 Simpson CJ, Elis Jones G, Taylor JD (1971) A seedling blight of Antirrhinum caused by Pseudomonas antirrhini. Plant Pathol 20:127–130 Stewart FC (1900) An anthracnose and stem rot of the cultivated snapdragon. NY (Geneva) Agric Exp Sta Bull 179:105–110 Takimoto S (1920) On a bacterial leaf-spot of Antirrhinum majus L. Bot Mag (Tokyo) 34:253–257 Tomioka K, Nishikawa J (2011) Anthracnose of snapdragon caused by Colletotrichum destructivum. J Gen Plant Pathol 77:60–63 UC IPM (2014) Pest management guidelines: floriculture and ornamental nurseries, UC ANR Publication 3392. http://www.ipm.ucdavis.edu/PMG/r280111211.html Valder PG (1963) New plant diseases. Plant Disease Survey, 33rd Report, Biol Brch NSW Dep Agric, p 35 Wegulo SN, Vilchez M (2006a) Evaluation of fungicides for control of Botrytis blight of lisianthus, 2004. F&N Tests 61:OT030 Wegulo SN, Vilchez M (2006b) Evaluation of fungicides for control of downy mildew of snapdragon, 2003. F&N Tests 61:OT027 Yang HC, Leu LS (1981) The occurrence of snapdragon wilt disease in Taiwan. Plant Prot Bull 23:55–57 Yarwood CE (1947) Snapdragon downy mildew. Hilgardia 17:241–250

Diseases of Stock Steven T. Koike

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-Like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Cottony Rot (Sclerotinia sclerotiorum) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Downy Mildew (Peronospora parasitica = P. matthiolae or Hyaloperonospora parasitica) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Fusarium Wilt (Fusarium oxysporum f. sp. mathioli) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Rhizoctonia Foot Rot (Rhizoctonia solani) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Verticillium Wilt (Verticillium dahliae and Verticillium zaregamsianum) . . . . . . . . . . . . 2.6 Additional Fungal and Fungus-Like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Bacterial and Phytoplasma Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Bacterial Blight (Xanthomonas campestris pv. incanae) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Blossom Blight (Pseudomonas syringae pv. maculicola) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Viral Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Turnip Mosaic Virus (TuMV, Genus: Potyvirus) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2 2 2 3 6 8 10 11 11 11 15 17 17 18

Abstract

The ornamental plant stock (Matthiola incana R. Br.) is a popular and versatile flowering species that can be grown as a garden or bedding plant as well as a field grown cut flower. Profitability of stock production depends on the management of a number of diseases. As part of the Brassicaceae, stock is subject to a number of pathogens that infect vegetable crucifers as well. Major pathogens include Sclerotinia, downy mildew, two wilt pathogens (Fusarium, Verticillium), Xanthomonas, and Turnip mosaic virus.

S.T. Koike (*) University of California Cooperative Extension, Monterey County, Salinas, CA, USA e-mail: [email protected] # Springer International Publishing AG 2016 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florist's Crops Diseases, Handbook of Plant Disease Management, DOI 10.1007/978-3-319-32374-9_26-1

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Keywords

Fusarium • Sclerotinia • Verticillium • Xanthomonas

1

Introduction

Stock (Matthiola incana R. Br.) is also known as annual stock, column stock, garden stock, or tensweek stock. A closely related ornamental species is M. longipetala (Vent.) DC which is named night-scented stock or evening stock. Likely originating from the Mediterranean area, stock is a popular floral commodity that is strongly scented and is grown as a garden or bedding plant and also as a cutflower (Armitage and Laushman 2003). Depending on the species and region, stock can be grown as annuals, biennials, or perennials. As a member of the Brassicaceae, stock is susceptible to many of the diseases that affect commercial cole crops (broccoli, cabbage, cauliflower, etc.) vegetables (Koike et al. 2007).

2

Fungal and Fungus-Like Diseases

2.1

Cottony Rot (Sclerotinia sclerotiorum)

Geographic occurrence and impact. Cottony rot (also known as white mold and Sclerotinia crown rot) has been reported on stock in Australia, Greece, Korea, New Zealand, Scotland, South Africa, and the United States (California, Michigan, and Pennsylvania). The disease can occasionally be important on stock. Symptoms/signs. Symptoms vary depending on the type of fungal inoculum that comes in contact with stock. The soilborne survival structure, the sclerotium, can germinate directly and the resulting mycelium can infect stock crowns at the soil line. Crown infections first result in a gray to brown lesion on crown tissues. Lesions expand, crowns become completely rotted, and the plant will suddenly collapse (Fig. 1). If conditions are moist enough, the diseased crown will support the growth of white mycelium and large (5–10 mm/0.2–0.4 in. in length), black, irregularly shaped, hard sclerotia (Fig. 2). A second type of inoculum produced by S. sclerotiorum, the ascospore, is airborne, lands on aboveground stock tissues, and causes a blight on stems, leaves, or flowers. To infect the host, the ascospore must land on tissue that is damaged or senescent. Once germinated, the ascospore produces mycelium that will colonize the compromised tissue and also spreads to adjacent healthy tissue, resulting in a brown, soft decay. White mycelium and black sclerotia also develop from these foliar infections. Biology and epidemiology. Cottony rot is caused by Sclerotinia sclerotiorum. There is one report of another species, S. minor, causing a crown rot on stock in Australia. S. sclerotiorum survives and overwinters as sclerotia in soil. Sclerotia can germinate and form infective mycelium or alternatively will produce small, tan, mushroom-like structures called apothecia (Fig. 3). Apothecia emerge from the soil

Diseases of Stock

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Fig. 1 Stock plants infected with Sclerotinia species can suddenly wilt and collapse (Photo courtesy of Steven Koike)

and release airborne ascospores that are carried by wind and infect susceptible stock leaves, stems, and flowers. When infected stock plants are plowed back into the soil, sclerotia that had formed on infected parts are returned to the soil and enable the pathogen to persist. S. sclerotiorum has an extremely broad host range, so sclerotia and ascospores produced from other crops, such as gazania, gaillardia, cauliflower, lettuce, pepper, and many others, can provide inoculum that can infect stock. Management • Cultural practices – Avoid planting stock in fields having a history of cottony rot. Avoid planting stock adjacent to vegetable or other flower crops that are highly susceptible to Sclerotinia and which exhibit symptoms of the disease. • Fungicides – Apply protectant fungicides to stock foliage prior to infection.

2.2

Downy Mildew (Peronospora parasitica = P. matthiolae or Hyaloperonospora parasitica)

Geographic occurrence and impact. This disease occurs worldwide on stock. Severe outbreaks of downy mildew can cause significant damage to the stock foliage and reduce flower quality and marketability. Symptoms/signs. The first symptoms are irregular, light green to slightly yellow, diffuse patches on the top surfaces of leaves. As disease develops, the yellow color intensifies and large portions of the leaf can be affected (Fig. 4). Irregular dark

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Fig. 2 The hard, black sclerotia (arrows) of Sclerotinia sclerotiorum can form on the outside of stock stems (Photo courtesy of Steven Koike)

Fig. 3 Small, tan apothecia of Sclerotinia sclerotiorum will release spores that can infect stock foliage (Photo courtesy of Steven Koike)

specks can be seen within the yellow patches. Diseased leaves can be twisted and, in advanced stages, the entire leaf can turn brown and die. White tufts of the pathogen are seen on the undersides of the lesions (Fig. 5). With a hand lens the tufts can be seen to consist of the branching, treelike structures that bear the downy mildew spores. If conditions are favorable, the sporulation can also occur on the top side of

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Fig. 4 Stock plants infected with downy mildew will first develop yellow patches on leaves (Photo courtesy of Steven Koike)

Fig. 5 The fine, white growth of downy mildew can be seen on the surfaces of stock leaves (Photo courtesy of Steven Koike)

leaves. For some stock cultivars, extensive sporulation can develop well before any symptoms are seen on the top of the leaves (Koike 2000). Downy mildew has been known to kill stock seedlings if conditions allow seedling cotyledons to be infected early and severely. Biology and epidemiology. Downy mildew is caused by the oomycete Peronospora parasitica. Researchers also refer to the pathogen as Peronospora matthiolae or Hyaloperonospora parasitica (Constantinescu and Fatehi 2002; Jafar 1963). Oomycete organisms are no longer considered true fungi but are more closely related to algae. This obligate pathogen forms mycelia and haustoria inside host tissues, and spore-bearing sporangiophores emerge through the leaf stomata. Some P. parasitica isolates and races form the sexual oospore stage, though this has not been observed on stock (Sherriff and Lucas 1989). There is considerable host specialization within the P. parasitica group of pathogens, and strains infecting stock apparently only infect this host (Sherriff and Lucas 1990). Cool (10 to 15  C/50 to 59  F), moist or high humidity conditions favor downy mildew sporulation. Spores,

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Fig. 6 Fusarium wilt can cause considerable damage to stock plantings (Photo courtesy of O’Neill)

which are short-lived, are dispersed via winds and splashing water. Germination of sporangia and infection usually require the presence of free moisture on the leaf surface. Lesion expansion is most rapid at 20  C/68  F. Management • Cultural practices – Irrigating early in the day, to enhance drying of the foliage, may reduce disease severity. • Fungicides – Applying protectant foliar fungicides prior to infection is the best means of controlling downy mildew. Fungicides such as metalaxyl can be used as seed treatments for controlling early downy mildew on seedlings. • Resistance – Differences in susceptibility may exist among different stock cultivars but resistant cultivars do not appear to be available.

2.3

Fusarium Wilt (Fusarium oxysporum f. sp. mathioli)

Geographic occurrence and impact. Fusarium wilt has been reported on stock in Germany (Gerlach 1975), Japan (Saito et al. 2008), South Africa, the United Kingdom, and the United States (Arizona, California) (Baker 1948). The disease has caused considerable damage in the United Kingdom (O’Neill et al. 2003, 2005; O’Neill and Mason 2014, Fig. 6). Symptoms/signs. In some cases, if young seedlings are infected early, these plants will rapidly wilt and die. On larger plants, the lower leaves are affected first,

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Fig. 7 In advanced cases Fusarium wilt will cause all the leaves to dry up (Photo courtesy of O’Neill)

developing a clearing of the veinal tissue, followed by a general yellowing and then wilting. The yellowing and wilting symptoms may occur on only one side of the plant. Plants will be stunted and wilt during the warmer times of the day. In advanced cases, the leaves can dry up and the plant can die (Fig. 7). Examination of the stem and taproot vascular tissues will reveal a brown discoloration, though the root is not decayed (Fig. 8). Infected stock seed plants may exhibit seed pods that appear flattened and tan in color. On stock, Fusarium wilt symptoms closely resemble those of Verticillium wilt. Biology and epidemiology. Like most Fusarium wilt pathogens, the stock pathogen (F. oxysporum f. sp. mathioli) is host specific to its original host and does not infect related species such as cabbage, kale, and radish. The report from Japan names the pathogen as F. oxysporum f. sp. conglutinans (Saito et al. 2008). This stock pathogen is typical of all F. oxysporum wilt isolates and forms four- to six-celled, fusiform, curved macroconidia, one- to two-celled oval- to kidney-shaped microconidia, and resilient chlamydospores. The pathogen can be seedborne and can survive via chlamydospores for long periods in the soil. The disease is most severe if the crop is grown during the warm summer and in regions where temperatures are high. The fungus is favored by temperatures around 25  C/77  F.

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Fig. 8 Fusarium wilt causes a brown discoloration in stem vascular tissue (Photo courtesy of O’Neill)

Management • Cultural practices – Avoid planting stock in fields known to be infested with the pathogen. Since the pathogen may survive in soil for 2 or more years, practice crop rotation with nonstock plants. Stock grown during the cooler winter or spring seasons and in cool coastal locations may escape the disease or experience less severe impacts from Fusarium wilt. • Soil disinfestation – Preplant soil treatments such as fumigation and steaming can help reduce soil inoculum.

2.4

Rhizoctonia Foot Rot (Rhizoctonia solani)

Geographic occurrence and impact. While this pathogen is broadly distributed throughout the world, Rhizoctonia foot rot of stock has been reported only from Australia, Greece, the United Kingdom, the United States (California, Florida), and Zimbabwe. The disease has also been called Rhizoctonia root rot or wirestem, and the pathogen also is implicated in damping-off disease of emerging stock seedlings. Symptoms/signs. The damping-off phase of the disease pertains only to young stock seedlings. For direct seeded crops, the fungus can attack seed or newly germinated seedlings and kill them prior to emergence (preemergence dampingoff), or can attack roots and lower stem (hypocotyl) tissues shortly after the plant has emerged above ground (postemergence damping-off). For recently emerged plants, the stems in contact with soil become water soaked and later brown in color; these stems are often girdled and the plants fall over and die.

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Fig. 9 Rhizoctonia solani infects the stock stem and causes the “wirestem” symptom (Photo courtesy of O’Neill)

The wirestem phase pertains to older seedlings and consists of browning and cracking of the stem epidermis that is in contact with soil (Dimock 1941, Fig. 9). These stem infections develop into lesions, with the outer tissue decaying away. When the outer stem layers deteriorate, only the fibrous inner xylem remains intact as a wiry strand, hence the name wirestem. Affected plants develop yellow foliage and later wilt. Seedling stems may break at the soil line, resulting in the plant falling over. Plants that survive will likely remain stunted and behind in development. Signs of the pathogen consist of coarse mycelium that can be observed, with a hand lens, in the lesion. The mycelium sometimes causes soil particles to adhere to and dangle from diseased stems. The pathogen is also known to cause a root rot on stock. Biology and epidemiology. Rhizoctonia foot rot or wirestem is caused by the soil inhabitant Rhizoctonia solani. In culture, R. solani forms coarse, brown, relatively thick hyphae that are characterized by right-angle branching. Hyphae are constricted at branch points, and a cross wall with a dolipore septum is formed right after each branch. Older cultures form small, brown, loosely aggregated clumps of mycelia that function as sclerotia. The fungus is multinucleate and has a sexual phase, Thanatephorus cucumeris (Sneh et al. 1991), though this stage has not been reported on stock. This species is extremely diverse and can be divided into anastomosis groups (AG) based on the process of hyphal fusion between compatible isolates.

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AG-2-1 is associated with Rhizoctonia diseases on Brassicaceae hosts (Keijer et al. 1997). While AG-2-1 isolates are pathogenic on plants in this family, these isolates may infect plants outside of the Brassicaceae. Likewise, non-AG-2-1 isolates may infect Brassicaceae hosts. Therefore, these AG pathogens are not absolutely restricted to certain plant groups. R. solani survives saprophytically in soil as mycelia or sclerotia; these fungal structures are the inocula for infecting stock seedlings and older plants. The pathogen is favored by warm soil conditions (25 to 30  C/77 to 86  F) but is capable of causing problems at much lower temperatures as well. Management • Cultural practices – When placing transplants in the field, avoid planting them too deeply in the soil as the hypocotyl stem tissue is the most susceptible part of the plant. Practice crop rotation so that nonhosts are included in the rotation (Henis et al. 1978). Do not plant stock in fields having undecomposed crop residues, since R. solani may be actively colonizing such residues. • Fungicides – When direct seeding in the field, use seed that has been treated with a fungicide. If available, direct-spray fungicides to the base of young stock plants in the field. • Sanitation – Practice thorough sanitation at nurseries to prevent contamination by R. solani. Clean and sanitize transplant trays and benches. Ensure that rooting media are not contaminated by infested soil or diseased plant residues.

2.5

Verticillium Wilt (Verticillium dahliae and Verticillium zaregamsianum)

Geographic occurrence and impact. Verticillium wilt has been reported on stock in Japan, New Zealand, and the United States (California, New York). Symptoms/signs. Lower leaves are affected first, becoming yellow and then wilted. In some cases, the leaf interveinal tissue turns yellow first and the veins remain green longer. Plants will be stunted and wilt during the warmer times of the day. The yellowing and wilting of leaves advances up the stem until the stock plant becomes completely affected and can die (Raabe and Wilhelm 1958). Examination of the stem and taproot vascular tissues can reveal a light tan to brown discoloration (Fig. 10). On stock, Verticillium wilt symptoms closely resemble those of Fusarium wilt. Biology and epidemiology. Verticillium wilt of stock appears to be caused by two Verticillium species (Inderbitzin et al. 2011). Most cases of stock Verticillium wilt are caused by V. dahliae. However, Verticillium wilt of stock in Japan is caused by V. zaregamsianum. This etiology is supported by inoculation experiments. For example, stock is susceptible to a tomato isolate of Verticillium which is the V. dahliae species; however, even though stock is a member of the Brassicaceae, stock is not susceptible to a V. longisporum isolate that infects Brussels sprouts and other brassica crops. In culture these stock pathogens form the typical Verticillium structures consisting of hyaline, verticillate conidiophores bearing three to four

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Fig. 10 Internal stem tissues infected with Verticillium will turn light tan to brown (Photo courtesy of Steven Koike)

phialides at each node. Conidia formed from phialides are hyaline, oval, and singlecelled. Older cultures form darkly pigmented, multicelled microsclerotia. The pathogen can survive as resilient microsclerotia for long periods in the soil. Management • Cultural practices – Avoid planting stock in fields known to be infested with the pathogen. Since the pathogen may survive in soil for many years, practice crop rotation with nonhost plants. • Soil disinfestation – Preplant soil treatments such as fumigation and steaming can help reduce soil inoculum.

2.6

Additional Fungal and Fungus-Like Diseases

Alternaria leaf spot (Alternaria japonica) (Davis et al. 1949; Simmons 2007). Club root (Plasmodiophora brassicae) (Samson and Walker 1982). Damping-off (Pythium species) (Fig. 11). Gray mold (Botrytis cinerea) (Baker et al. 1954). Phytophthora root rot (Phytophthora species) (Fig. 12). Powdery mildew (Erysiphe cruciferarum) (Braun and Cook 2012). White rust (Albugo candida) (Choi et al. 2007; Garcia-Blazquez et al. 2006).

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Bacterial and Phytoplasma Diseases

3.1

Bacterial Blight (Xanthomonas campestris pv. incanae)

Geographic occurrence and impact. Bacterial blight is a significant disease of stock that can cause significant losses in yield and quality (Fig. 13). The disease is known to occur wherever stock is grown because it is seedborne (Minardi et al. 1988; Rahimian and Okhovatian 1989).

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Fig. 11 Pythium infects stock seedlings early and results stunted growth and/or plant death (Photo courtesy of O’Neill)

Fig. 12 Phytophthora infections result in rotted roots and a reduced root system (Photo courtesy of O’Neill)

Symptoms/signs. Young seedlings can develop the disease and exhibit yellowing of the stem and leaves, followed by complete wilting of the foliage and plant death. On older plants, initial symptoms consist of yellowing and wilting of the lower leaves. Where lower leaves are attached, the stem develops water-soaked lesions that later turn dark brown to black and are sunken (Fig. 14). Lesions enlarge and can girdle the stem, resulting in complete wilting of the plant. These lesions also weaken the stem and are the points where the stem cracks and causes the plant to fall over (Fig. 15). Examination of the vascular tissue shows a darkened discoloration. Biology and epidemiology. The pathogen, Xanthomonas campestris pv. incanae, is apparently host specific to stock and did not infect cabbage, cauliflower, or kale in

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Fig. 13 Bacterial blight can cause significant losses in stock plantings (Photo courtesy of Steven Koike)

inoculation experiments. In culture the pathogen is typical of most X. campestris pathovars and is characterized by slow growing, mucoid, yellow colonies. The pathogen is seedborne (Kendrick 1938) and can also persist in the soil in association with diseased stock plant residues. Disease is initiated by planting infested seed or by planting stock into soil containing plant residues of diseased stock. Once disease has developed, the bacteria are splashed to other plants via rain and overhead sprinkler irrigation. Cool, wet weather favors infection and disease development. Management • Cultural practices – Stock seed can be treated with hot water (50 to 55  C/122 to 131  F for 10 min) to disinfest seed; however, such treatments can result in loss of seed germination and need to be implemented carefully. Use drip or furrow irrigation instead of overhead sprinklers. Seed crops should be produced in regions where rain does not occur during seed production. Avoid planting stock in back-to-back plantings; a crop rotation of nonstock crops for 2–3 years likely results in eradication of the pathogen from soil. For stock grown in nurseries and containers, remove and discard symptomatic plants. • Resistance – Resistant M. incana cultivars are not yet available, though resistance has been found in M. aspera, M. longipetala, and M. tricuspidata (Ecker et al. 1995).

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Fig. 14 Stock plants infected with bacterial blight will first develop dark lesions on stems (Photo courtesy of Steven Koike)

Fig. 15 Bacterial blight infections weaken the stems of stock, causing them to crack and break (Photo courtesy of Steven Koike)

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Fig. 16 Brown, collapsed petals caused by Pseudomonas syringae pv. maculicola (Photo courtesy of Cother)

3.2

Blossom Blight (Pseudomonas syringae pv. maculicola)

Geographic occurrence and impact. Blossom blight has been reported only from Australia (Cother and Noble 2009). However, the blossom blight pathogen is widely distributed worldwide on cole crop and other crucifer vegetables (Cintas et al. 2001; Koike et al. 2007; Peters et al. 2004; Shackleton 1996). Symptoms/signs. Disease first develops on flowers located on the lower portion of the spikes. Infected petals turn brown and later collapse (Fig. 16). Collapsed petals adhere to adjacent flowers. As disease develops, infections progress up the spike and in severe cases affect all the flowers (Cother and Noble 2009). Lesions can also develop where the flower base is attached to the spike. Foliar symptoms consist of small, angular, brown leaf spots that can be seen from both upper and lower leaf surfaces (Cother and Noble 2009, Fig. 17). This symptom has been referred to as “pepper leaf spot” on vegetable crucifers (Koike et al. 2007). Leaf spots may be surrounded by yellow borders. A slight discoloration can be observed in stem vascular tissue (Fig. 18). For stock plants producing seed, seed pods may develop lesions (Fig. 19). Biology and epidemiology. The cause of blossom blight is Pseudomonas syringae pv. maculicola (Cother and Noble 2009). The pathogen is an aerobic, Gram-negative bacterium that on standard microbiological media forms colonies that are cream to light yellow in color and smooth (Cintas et al. 2001). When cultured on King’s medium B, strains that infect stock do not produce the diffusible pigment that fluoresces blue under ultraviolet light; this stock pathogen, therefore, is a nonfluorescing type of P. syringae. Consistent with P. syringae pv. maculicola isolates from crucifer vegetable crops, this stock pathogen can infect cauliflower and tomato (Cother and Noble 2009; Wiebe and Campbell 1993). The pathogen is seedborne in vegetable hosts and appears also to be seedborne in stock. More details on disease epidemiology are not available.

16 Fig. 17 The blossom blight pathogen can also cause small, brown leaf spots that may be surrounded by yellow borders (Photo courtesy of Cother)

Fig. 18 A brown discoloration in stem tissue can be caused by Pseudomonas syringae pv. maculicola (Photo courtesy Cother)

Fig. 19 Stock seedpods infected with Pseudomonas syringae pv. maculicola can develop dark brown lesions (Photo courtesy of Cother)

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Management. Disease control options are based on standard integrated disease control principles since the exact details are not available on how the disease develops. Resistant M. incana cultivars are not available. • Cultural practices – Stock seed can presumably be infested with the pathogen and can be treated with hot water, though such treatments can result in loss of seed germination and need to be implemented carefully. Use drip or furrow irrigation instead of overhead sprinklers. Seed crops should be produced in regions where rain does not occur during seed production. Avoid planting stock in back-to-back plantings. This pathogen likely does not persist for long periods in soil; however, a crop rotation using nonbrassica and nonstock crops for 2 years is advisable. For stock grown in nurseries and containers, remove and discard symptomatic plants.

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Viral Diseases

4.1

Turnip Mosaic Virus (TuMV, Genus: Potyvirus)

Geographic occurrence and impact. Stock is reported to be susceptible to a dozen or more viruses wherever the flower is grown. Some of the more familiar virus pathogens are Alfalfa Mosaic Virus, Beet Curly Top Virus, Cauliflower Mosaic Virus, Cucumber Mosaic Virus, Tomato Spotted Wilt Virus, and Turnip Mosaic Virus. It is not rare to have a stock plant infected with more than one virus (Alioto et al. 1994; Yoon et al. 1998). It appears that Turnip Mosaic Virus (TuMV) is perhaps the most commonly reported virus pathogen of stock and occurs worldwide (Bahar et al. 1985; Rosciglione and Cannizzaro 1983). Symptoms/signs. As with most viruses, symptoms caused by TuMV vary depending on the strain of the virus, stock cultivar, age of plant when infected, and environmental conditions. Leaves will exhibit a range of symptoms consisting of mottling, mosaic, formation of dark green patches or “green islands,” and clearing of veins. Leaves may also be curled, twisted, or otherwise distorted. The dark-colored (pink, red, purple) flowers that contain anthocyanin will show breaking, in which the petal coloration is broken up by streaks and sectors of nonpigmented tissue (Severin and Tompkins 1948, 1950; Tompkins 1939, Fig. 20). Cultivars with white and yellow colored flowers do not show flower breaking. Infected plants may be distorted. Biology and epidemiology. The virus survives between stock crops in living plants such as cruciferous weeds and volunteer stock. Aphids feed on these reservoir hosts, move to stock plants, and inject the virus during feeding. The virus is borne on the stylet of aphids and is transmitted in a nonpropagative (nonpersistent) manner. Management • Cultural practices – Old and infected stock plantings should be disked and buried in the soil. Remove cruciferous weeds and volunteer stock from around fields.

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Fig. 20 If infected with Turnip mosaic virus, darkcolored stock flowers will “break” and develop white streaks and sectors (Photo courtesy of Steven Koike)

• Vector management – Control the aphids by using insecticides and other IPM methods. • Resistance – Some stock lines and cultivars are resistant to TuMV (Johnson and Barnhart 1956; San Juan and Pound 1963).

References Alioto D, Stavolone L, Aloj B (1994) Serious alterations induced by Cauliflower mosaic virus (CaMV) and Turnip mosaic virus (TuMV) on Matthiola incana. Inf Fitopat 44:43–46 Armitage AM, Laushman JM (2003) Specialty cut flowers, 2nd edn. Timber Press, Portland, 586 pp Bahar M, Danesh D, Dehghan M (1985) Turnip mosaic virus in stock plant. Iranian J Plant Pathol 21(11–12):33–39 Baker KF (1948) Fusarium wilt of garden stock (Matthiola incana). Phytopathology 38:399–403 Baker KF, Matkin OA, Davis LH (1954) Interaction of salinity injury, leaf age, fungicide application, climate, and Botrytis cinerea in a disease complex of column stock. Phytopathology 44:39–42 Braun U, Cook RTA (2012) Taxonomic manual of the Erysiphales (Powdery mildews). CBS-KNAW Fungal Biodiversity Centre, Utrecht, 707 pp Choi Y-J, Shin HD, Hong S-B, Thines M (2007) Morphological and molecular discrimination among Albugo candida materials infecting Capsella bursa-pastoris world-wide. Fung Divers 27:11–34 Cintas NA, Bull CT, Koike ST, Bouzar H (2001) A new bacterial leaf spot disease of broccolini, caused by Pseudomonas syringae pv. maculicola, in California. Plant Dis 85:1207 Constantinescu O, Fatehi J (2002) Peronospora-like fungi (Chromista, Peronosporales) parasitic on Brassicaceae and related hosts. Nova Hedwigia 74:291–338 Cother EJ, Noble DH (2009) Identification of blossom blight in stock (Matthiola incana) caused by Pseudomonas syringae pv. maculicola. Australas Plant Pathol 38:242–246 Davis LH, Sciaroni RH, Pritchard F (1949) Alternaria leafspot of garden stock in California. Plant Dis Rep 33:432–433 Dimock AW (1941) The Rhizoctonia foot-rot of annual stocks (Matthiola incana). Phytopathology 31:87–91 Ecker R, Zutra D, Barzilay A, Osherenko E, Rav-David D (1995) Sources of resistance to bacterial blight of stock (Matthiola incana R. Br.). Genet Res Crop Evol 42:371–372

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Garcia-Blazquez G, Constantinescu O, Telleria MT, Martin MP (2006) Preliminary checklist of Albuginales and Peronosporales (Chromista) reported from the Iberian Peninsula and Balearic Islands. Mycotaxon 98:185–188 Gerlach W (1975) The first case of Fusarium wilt on garden stock (under glass) in Germany. Nachrichtenblatt des Deutschen Pflanzenschutzdienstes 27:17–20 Henis Y, Ghaffar A, Baker R (1978) Integrated control of Rhizoctonia solani damping-off of radish: effect of successive plantings, PCNB, and Trichoderma harzianum on pathogen and disease. Phytopathology 68:900–907 Inderbitzin P, Bostock RM, Davis RM, Usami T, Platt HW, Subbarao KV (2011) Phylogenetics and taxonomy of the fungal vascular wilt pathogen Verticillium, with the descriptions of five new species. PLoS One 6:1–22 Jafar H (1963) Studies on downy mildew (Peronospora matthiolae (Roume-guere) Gaumann) on stocks (Matthiola incana R.Br.). N Z J Agric Res 6:70–82 Johnson BL, Barnhart D (1956) Transfer of mosaic resistance to commercial varieties of Matthiola incana. Proc Am Soc Hort Sci 67:522–533 Keijer J, Korsman MG, Dullemans AM, Houterman PM, de Bree J, van Silfhout CH (1997) In vitro analysis of host plant specificity in Rhizoctonia solani. Plant Pathol 46:659–669 Kendrick JB (1938) A seed-borne bacterial disease of garden stocks, Matthiola incana. Phytopathology 28:12 Koike ST (2000) Downy mildew of stock, caused by Peronospora parasitica, in California. Plant Dis 84:103 Koike ST, Gladders P, Paulus AO (2007) Vegetable diseases: a color Handbook. Manson Publishing Ltd, London, 448 pp Minardi P, Mazzucchi U, Parrini C (1988) Epidemics of bacterial blight of stock (Matthiola incana R. Br.) caused by Xanthomonas campestris pv. incanae in Tuscany. Inf Fitopat 38:43–46 O’Neill TM, Mason L (2014) Contrasting effects on Fusarium wilt of column stock following use of bark as a soil amendment. Acta Hortic 1044:139–143 O’Neill TM, Shepherd A, Inman AJ, Lane CR (2003) Wilt of stock (Matthiola incana) caused by Fusarium oxysporum in the United Kingdom. New Dis Reports 8:10 O’Neill TM, Green KR, Ratcliffe T (2005) Evaluation of soil steaming and a formaldehyde drench for control of Fusarium wilt in column stock. Acta Hortic 698:129–133 Peters BJ, Asha GJ, Cother EJ, Hailstones DL, Nobleb DH, Urwin NAR (2004) Pseudomonas syringae pv. maculicola in Australia: pathogenic, phenotypic and genetic diversity. Plant Pathol 53:73–79 Raabe RD, Wilhelm S (1958) Verticillium wilt of garden stock (Matthiola incana). Phytopathology 48:610–613 Rahimian H, Okhovatian H (1989) Bacterial blight of stock in Mazandaran. Iranian J Plant Pathol 25(11–12):29–37 Rosciglione B, Cannizzaro G (1983) Matthiola incana R. Br., natural host of Turnip mosaic virus in Sicily. Tecnica Agricola 35:251–257 Saito K, Domon K, Honma T, Kawasaki T, Ogata M, Hori Y (2008) Soil reduction disinfection of stock wilt caused by Fusarium oxysporum f. sp. conglutinans race 3. Annu Rep Soc Plant Protect N Jpn 59:71–73 Sampson PJ, Walker J (1982) An Annotated List of Plant Diseases in Tasmania. Department of Agriculture, Tasmania. 121 pp San Juan MO, Pound GS (1963) Resistance in Matthiola incana to the Turnip mosaic virus. Phytopathology 53:1276–1279 Severin HHP, Tompkins CM (1948) Aphid transmission of mild mosaic virus of annual stock. Hilgardia 18:539–547 Severin HHP, Tompkins CM (1950) Aphid transmission of severe mosaic virus of annual stock. Hilgardia 20:93–108 Shackleton DA (1996) A bacterial leaf spot of cauliflower in New Zealand caused by Pseudomonas syringae pv. maculicola. N Z J Sci 9:872–877

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Sherriff C, Lucas JA (1989) Heterothallism and homothallism in Peronospora parasitica. Mycol Res 92:311–316 Sherriff C, Lucas JA (1990) The host range of isolates of downy mildew, Peronospora parasitica, from Brassica crop species. Plant Pathol 39:77–91 Simmons EG (2007) Alternaria: an identification manual. Centraalbureau voor Schimmelcultures, Utrecht, 775 pp Sneh B, Burpee L, Ogoshi A (1991) Identification of Rhizoctonia Species. APS Press, St. Paul, 133 pp Tompkins CM (1939) Two mosaic diseases of annual stock. J Agric Res 58:63–77 Wiebe WL, Campbell RN (1993) Characterization of Pseudomonas syringae pv. maculicola and comparison with Pseudomonas syringae pv. tomato. Plant Dis 77:414–419 Yoon J-Y, Choi H-S, Ryu H-Y, Harm Y-I, Choi J-K (1998) Colour breaking syndrome of Matthiola incana caused by double infection of Cucumber mosaic virus and Turnip mosaic virus. Korean J Plant Pathol 14:220–222

Diseases of Sunflower Thomas J. Gulya, Febina Mathew, Robert Harveson, Samuel Markell, and Charles Block

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Alternaria Leaf Spot and Blight (Alternaria spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Botrytis Blight or Gray Mold (Botrytis cinerea Persoon) . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Charcoal Rot (Macrophomina phaseolina (Tassi) Goidànich) . . . . . . . . . . . . . . . . . . . . . . 2.4 Damping-off (Pythium spp., Phytophthora spp., and Rhizoctonia solani Kühn) . 2.5 Downy Mildew (Plasmopara halstedii (Farlow) Berlese and de Toni) . . . . . . . . . . . . . 2.6 Fusarium Stalk Rot and Wilt (Fusarium spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7 Leaf Smut (Entyloma spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.8 Phoma Black Stem (Phoma macdonaldii Boerema) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.9 Phomopsis Stem Canker (Diaporthe helianthi and Other Diaporthe spp.) . . . . . . . . . 2.10 Powdery Mildews (Erysiphe and Other Genera) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.11 Rhizopus Head Rot (Rhizopus spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.12 Rust – Puccinia helianthi Schwein. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Thomas J. Gulya was retired. T.J. Gulya (*) USDA-Agricultural Research Service, Sunflower and Plant Biology Research Unit, Fargo, ND, USA e-mail: [email protected] F. Mathew Department of Agronomy, Horticulture, and Plant Science, South Dakota State University, Brookings, SD, USA R. Harveson Department of Plant Pathology, University of Nebraska, Panhandle Research and Extension Center, Scottsbluff, NE, USA S. Markell Department of Plant Pathology, North Dakota State University, Fargo, ND, USA C. Block Seed Science Center, Iowa State University, Ames, IA, USA # Springer International Publishing Switzerland 2016 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florists' Crops Diseases, Handbook of Plant Disease Management, DOI 10.1007/978-3-319-32374-9_27-1

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Rusts – Other Minor Ones (Puccinia spp., Coleosporium helianthi Arthur) . . . . . . . Sclerotinia Stalk Rot and Head Rot (Sclerotinia sclerotiorum (Libert) de Bary and Sclerotinia minor Jagger) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.15 Septoria Leaf Spot and Blight (Septoria spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.16 Sclerotium Blight (Sclerotium rolfsii Saccardo) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.17 Verticillium Leaf Mottle/Wilt (Verticillium dahliae Klebahn) . . . . . . . . . . . . . . . . . . . . . . . 2.18 White Rust (Albugo tragopogonis (DeCandolle) Gray = Pustula helianthicola Rost and Thines) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Bacterial and Phytoplasma Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Apical Chlorosis and Bacterial Leaf Spot (Pseudomonas syringae pv. tagetis and Pseudomonas syringae pv. helianthi) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Bacterial Stalk Rot/Head Rot (Erwinia = Pectobacterium spp.) . . . . . . . . . . . . . . . . . . . . . 3.3 Aster Yellows (Phytoplasma spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Viral Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Sunflower Mosaic Virus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Nematodes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 Root-Knot Nematodes (Meloidogyne Species) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2 Root-Lesion Nematodes (Pratylenchus Species) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3 Other Nematodes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.13 2.14

21 23 27 28 29 31 33 33 35 37 38 38 40 40 42 43 43

Abstract

Sunflower (Helianthus annuus L.) has become a popular cut flower in the United States and globally. Easy to grow, bright and cheerful large flowers, with ray flowers from white to yellow to orange to magenta, it is often a centerpiece in mixed bouquets. Sunflower and the rest of the Helianthus genus are native to North America, and, as such, there is a native population of disease organisms and insect pests that can be a production challenge. Fortunately, ornamental sunflower and oilseed sunflower are the same species, and thus genetic advances made with oilseed germplasm can be readily transferred to ornamentals, if the need arises. The major diseases posing threats to ornamental sunflower are the same as those threatening oilseed sunflower hybrids, namely downy mildew, rust, Phomopsis stem canker, Sclerotinia wilt, Verticillium wilt, and charcoal rot. However, greenhouse production presents a different environment, and thus, there can be diseases affecting cut sunflowers that are seldom seen under field production. General management strategies for diseases of florists’ crops may be found in the introductory chapters on integrated disease management. Keywords

Alternaria • Botrytis • Fusarium • Phomopsis • Plasmopara halstedii • Puccinia helianthi • Phoma • Sclerotinia sclerotiorum • Verticillium • White rust

1

Introduction

Sunflower is a unique crop in that it is grown for ornamental use, as an oilseed crop, and for human consumption. All three uses dictate a different germplasm base, but all are Helianthus annuus. Additionally, wild H. annuus is native to the United

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States, and it occurs in almost all of the 48 contiguous states. Another unique feature of sunflower is that it is one of the few crops whose progenitors are native to the United States and was domesticated by Native Americans (Heiser 2008). There are 53 wild Helianthus species, of which all but one are found in the United States (Heiser et al. 1969; Schilling 2006). Among the 53 Helianthus species, 14 are annual species and 39 are perennial species. Many of these species are adapted to unique environmental niches and thus potential sources of genes, which can be transferred to cultivated sunflower through conventional breeding methods. The popularity of ornamental sunflowers is relatively recent, and this has spurred university research in many areas of the country, addressing the challenges of many different environments. Oilseed sunflower, in comparison, has been a large and essential global oilseed crop for many decades, and there are numerous sunflower disease guides in many languages besides English (Chattopadhyay et al. 2015; Gulya et al. 1997) including French (CETIOM 2015), Portuguese (Leite 1997), Serbo-Croatian (Maric et al. 1987), Spanish (Alonso et al. 1988), and Russian (Artokhin and Ignatova 2013), whose information is just as relevant to cut flower production as for oilseed sunflower. In addition, the American Phytopathology Society has recently issued the Compendium of Sunflower Diseases (Harveson et al. 2016), which will have similar information to this chapter, but with extensive images which should aid with identification. In the United States, extension agencies of several universities have addressed the need for information on the production of cut and potted sunflowers, but few have detailed information on disease management (Garfinkel and Panter 2015; Gill et al. 2003; Schoellhorn et al. 2003; Whipker et al. 1998). The vast majority of public sunflower research in the United States is centered in Fargo, North Dakota, where both North Dakota State University and the USDA-ARS Sunflower and Plant Biology Research Unit are located. North Dakota State University and the USDA-ARS in Fargo have excellent research information available online, at www.ag.ndsu.ext/crops/sunflower and www.ars.usda.gov/main/ site_main.htm?modecode=30-60-05-15, respectively. Lastly, there are two other organizations that foster sunflower research, namely the National Sunflower Association (NSA) and the International Sunflower Association (ISA), whose websites are www.sunflowernsa.com and www.isasunflower.org, respectively. The NSA website is publically accessible and contains archives of magazine articles, papers and posters presented at over 30 past “research forums,” and a photograph gallery, one part of which deals with diseases and deformities.

2

Fungal and Fungus-like Diseases

2.1

Alternaria Leaf Spot and Blight (Alternaria spp.)

Geographic occurrence and impact. Alternaria leaf blight affects sunflower throughout the world and is a major defoliating pathogen in warm, humid climates. The fungus may also cause linear spots on the stems and water-soaked, sunken lesions on the back of sunflower head.

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Fig. 1 (a) Alternaria leaf blight lesions. (b) Alternaria stem lesions

Symptoms/signs. Many Alternaria species can cause leaf spots on sunflower, but symptoms are similar for all, making field identification impractical. The primary symptom is dark brown lesions on leaves, and also on stems, petioles, and bracts. Initially the leaf spots are small, dark, and angular (Fig. 1), but with time, they coalesce into large, necrotic areas resulting in defoliation. Defoliation starts with the lower leaves where the microclimate is most favorable. Stem lesions typically are narrow (1–3 mm) black streaks up to 3 cm long. Biology and epidemiology. Although Alternaria helianthi (Hansford) Tubaki and Nishihara, now reclassified as Alternariaster helianthi (Hansford) Simmons, is the major causal agent, there are eight other species reported on sunflower, including A. zinniae Pape, A. helianthicola Rao and Rajagopalan, A. helianthiinficiens Simmons, Walcz and Roberts, A. leucanthemi Nelen (syn. Teretispora leucanthemi (Nelen) Simmons), A. protenta Simmons, A. roseogrisea Roberts, A. tenuissima (Nees) Wiltshire, and A. alternata (Fries) Keissler. Most of these Alternaria spp. are specific for sunflower (except the opportunistic saprophyte A. alternata). However, A. zinniae has a broad host range that includes several Asteraceae genera, including weeds (Bidens, Cardus, Eupatorium, and Xanthium) and ornamentals (Aster, Calendula, Chrysanthemum, Dahlia, Tagetes, Tithonia, and Zinnia). Because symptoms caused by the Alternaria pathogens look very similar, field identification based on symptoms is difficult and may be inaccurate. Conidial morphology (Simmons 2008) combined with genetic analysis is the only sure means to delineate species.

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As A. helianthi is the primary causal agent and most widespread, this section addresses just this species (Allen et al. 1983a, b, c). Alternaria helianthi overwinters on infected plant residue, but wild or volunteer sunflowers may also serve as reservoirs. All species including A. helianthi may also be seedborne. The conidia are windborne and spread by splashing water onto the lower leaves. They germinate best at temperatures >26  C and require a minimum of 4 h of leaf wetness for sporulation. Disease progress is also heavily dependent on the duration of leaf wetness following initial infection, as the generation of new spores can occur within 2 days. Young seedlings are more susceptible than older plants, but senescing lower leaves on mature plants frequently are defoliated by Alternaria spp. Management • Cultural practices – including removal of wild and volunteer sunflowers, removal or incorporation in the soil of previous sunflower residues, and minimizing extended leaf wetness will all reduce disease potential. • Fungicides – Seed treatments (captan, thiram, mancozeb) may offer some control (Jeffrey et al. 1985), but most growers rely upon multiple applications of fungicides containing chlorothalonil, iprodione, procymidone, or vinclozolin. More recent fungicide tests with ornamental sunflowers demonstrated that seed treatments with difenoconazole, prochloraz, pyrifenox or triadimenol were effective (Wu and Wu 2003). Consult government fungicide guides for specific products and rates. • Resistance – Disease resistance has been found in oilseed sunflower and some attempts have been made to incorporate that into hybrids, but there have been no reports of resistance in ornamental cultivars.

2.2

Botrytis Blight or Gray Mold (Botrytis cinerea Persoon)

Geographic occurrence and impact. The fungus causing gray mold, Botrytis cinerea (syn. Botryotinia fuckeliana (de Bary) Whetzel), is found worldwide and impacts many floral, fruit, and vegetable crops. On sunflower, the pathogen causes mostly a head rot. However, it can also cause spots on leaves and petals and is most often observed in areas with cool summers accompanied by frequent rains. It is rarely seen outdoors in most areas of the United States but could be a problem in greenhouses where sunflower leaves remains wet for extended periods of time. Symptoms/signs. Gray mold first appears as sunken brown spots on the back surface of the head. During periods of high humidity or rainfall, these lesions will be covered with conidia, giving the infected tissue a gray color (Fig. 2). The gray color is an easy, macroscopic way to distinguish Botrytis head rot from other head rots. If heads are infected early and wet conditions persist, Botrytis may form small, black sclerotia within the sunflower head. Botrytis has also been observed to cause petiole infections in Egypt, India, and Pakistan, which can progress to the stem, resulting in lesions up to 10 cm long. Biology and epidemiology. Botrytis cinerea is a ubiquitous fungus and a facultative pathogen with a wide host range encompassing most ornamentals, soft fruit, and monocots.

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Fig. 2 Botrytis head rot

B. cinerea overwinters in the soil as sclerotia or as mycelium on infected plant debris and is also seedborne (Coley-Smith et al. 1980; Elad et al. 2007). Sclerotia germinate to produce conidia, which can colonize dead organic matter as well as living host tissue. Conidia are windblown and water splashed and require a wound or senescent tissue to gain entry into a plant. Optimal conditions for infection are 15–25  C and 90 % relative humidity. Under optimal conditions, Botrytis may produce a new crop of conidia every 5–7 days. Botrytis produces copious amounts of spores, which develop rapidly in decaying vegetation and on senescing flowers. Because of the wide host range, the inocula for sunflower infection may come from many other floral crops or weeds. Management. For control of Botrytis on greenhouse or field grown ornamental sunflowers, fungicide applications coupled with cultural practices are useful. • Cultural practices – Since spores readily form on infected plant tissue, elimination of infected plants is essential. Avoidance of overhead watering, especially prior to bloom, is important, or at least irrigate early in the day to promote drying. Wider spacing between plants will minimize the RH within the canopy, but this may not be practical under the intensive plant populations as seen in commercial field operations. The fungus can develop at low temperatures, so cut flowers with no apparent infection may develop gray mold even under refrigeration. As free water is necessary for infection, minimizing conditions that lead to condensation in cold storage is critical (Gibson et al. 2014). • Resistance – Oilseed germplasm with resistance to Botrytis has been identified, but to date these genes have not been incorporated into either oilseed hybrids or ornamental germplasm (Kanyion and Friedt 1993).

2.3

Charcoal Rot (Macrophomina phaseolina (Tassi) Goidànich)

Geographic occurrence and impact. Charcoal rot is a serious root-infecting disease that is found worldwide but is most severe in hot, dry climates. It is both soil- and

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Fig. 3 (a) Charcoal rot, showing pith compression and microsclerotia. (b) Charcoal rot on sunflower, showing pith with microsclerotia

seedborne (Raut 1983). The fungus has little effect on plants until they become stressed; the stress may be due to heat, lack of soil moisture, or seed filling. The disease may kill plants outright, but it also results in reduced head size. Symptoms/signs. Damping-off may occur if M. phaseolina-infected seeds germinate in soil at or near 35  C. Usually, symptoms appear on older plants after flowering, with a silvery-gray lesion at the base of the plant and eventually encircling the stem. Upon splitting open the stem longitudinally, one will notice pith disintegration, and often a characteristic compression of the pith into layers. Examination with a 5 to 10X hand lens will reveal tiny microsclerotia, which appear as pepper grains scattered within the pith and on the inside of the rind. Severe infections, especially on stressed plants, will cause premature death (Fig. 3a, b). Biology and epidemiology. Charcoal rot is incited by Macrophomina phaseolina. This fungus has had 17 binomials as synonyms, of which Rhizoctonia bataticola and Sclerotium bataticola are most commonly found in older literature. The fungus is characterized by producing black microsclerotia, 60–200 μm in diameter. Occasionally an isolate will be found which produces pycnidia (and pycnidiospores) in culture, but this is very rare in infected sunflower tissue. The fungus has a wide host range, but there is no evidence of isolates having host specificity. Macrophomina phaseolina has an extremely broad host range, encompassing 500 species of both monocots and dicots (Dhingra and Sinclair 1978). Thus, nearly all ornamentals are hosts, as well as field crops, vegetables, and weeds. The fungus does not produce any conidia, but persists by forming small, irregular, black microsclerotia (60–200 μm in diameter) which are barely visible with the unaided eye. Microsclerotia in infected plant residue are released into soil as the host tissues decompose, and they can persist in the soil for at least 3 years. The fungus is unusual in its affinity or tolerance of high temperatures and grows best at soil temperatures > 35  C. Other conditions which favor disease development include low soil moisture, light sandy soils, as well as stress of the host plants due to herbicides or nematodes

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(Singh et al. 2012). Charcoal rot is frequently associated with Fusarium stalk rot. Additionally, the sunflower stem weevil has been shown to vector Macrophomina phaseolina in sunflowers (Yang and Owen 1982). Management • Cultural practices – Minimizing plant stress by lowering plant densities, and providing adequate soil moisture, will reduce the onset of the disease. However, since sunflowers grown for cut flowers are harvested in the late bud stage, charcoal rot should seldom be seen. Crop rotations will have little effect on the fungus due to its wide host range and the longevity of the microsclerotia. • Fungicides – Seed treatments with fungicides will minimize the introduction of M. phaseolina (Raut 1983) and protect against early infection.

2.4

Damping-off (Pythium spp., Phytophthora spp., and Rhizoctonia solani Kühn)

Geographic occurrence and impact. Damping-off of oilseed sunflower is rarely noted in the literature, but is more likely to be observed with ornamental sunflowers due to their intensive management practices. Damping-off, as the name implies, leads to seedling death, whether pre- or postemergence, and thus can be quite devastating. Pathogens inciting damping-off are global in occurrence, although exact species may differ from region to region. Symptoms/signs. Damping-off may be the result of seeds or seedlings rotting prior to emergence, or more frequently, the seedlings collapse as they emerge as the roots and/or stem at the soil line becomes necrotic and the plant collapses. When older seedlings are affected, the plant becomes stunted and dies later. Pythium spp. generally attack the root tips, causing a dark brown to black rot, while various Phytophthora spp. may attack the fine roots including the tap root, crown, and all belowground parts (Banihashemi 1975). Rhizoctonia solani commonly causes a brown to black lesion encircling the stem at the soil line and may occur both on seedlings and older plants (Srinivasan and Visalakchi 2010). Biology and epidemiology. All three pathogen groups have broad host ranges. Pythium species reported on sunflower include P. aphanidermatum (Edson) Fitzpatrick, P. irregulare Buisman, P. debaryanum Hesse, P. rostratum Butler, and P. splendens Hans Braun (Hendrix and Campbell 1973). Phytophthora species reported on sunflower include P. cryptogea Pethybridge and Lafferty, and P. drechsleri Tucker. Rhizoctonia solani isolates from sunflower belong to anastomosis groups AG-3 and AG-4. Pythium spp., Phytophthora spp., and R. solani are all soilborne and found worldwide, and Rhizoctonia has been shown to be seedborne in sunflower (Lakshmidevi et al. 2010). They can survive for years in soil or infected crop residues with overwintering structures (sclerotia for Rhizoctonia and oospores for Pythium and Phytophthora). Sclerotia of R. solani germinate to form mycelium which attacks seedling roots. The oospores produced by Pythium spp. and

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Phytophthora spp. germinate to form motile zoospores that are chemotactically attracted to root exudates. Thus, water-logged soils are necessary for infection by Pythium spp. and Phytophthora spp., with longer periods of saturation leading to more damping-off. Management • Cultural practices – Crop rotation is generally of little use due to the broad host ranges and the longevity of overwintering fungal structures. At this time, disease management is aimed at prevention. Using disease-free planting media and avoiding overwatering are the easiest means of minimizing damping-off. • Sanitation – Discard pots or flats with plants showing damping-off. • Soil disinfestation – For field production in warmer climates, solarization has been reported to provide control. • Fungicides and biocontrols – Fungicidal seed treatments, initially registered for control of downy mildew (metalaxyl and mefenoxam) will generally give control of Pythium and Phytophthora, while broad-spectrum strobilurin fungicides will help manage Rhizoctonia. Various biocontrol formulations of Bacillus, Gliocladium, and Trichoderma spp. are available that are applied as seed inoculations, incorporated into potting soil, or used as soil drenches. Fungicide soil drenches are also used in nonorganic production. Consult government publications for recommended products. • Resistance – Although differences in susceptibility have been noted among oilseed sunflowers to these three pathogens, resistance has not been incorporated into either oilseed or ornamental sunflower.

2.5

Downy Mildew (Plasmopara halstedii (Farlow) Berlese and de Toni)

Geographical occurrence and impact. The pathogen causing downy mildew is found worldwide, with the exception of Australia. Downy mildew is one of the major diseases affecting oilseed sunflower, and can be devastating on ornamentals as well. The pathogen primarily causes systemic infection, which renders the plant unsalable thus affecting florists’ trade. Symptoms/signs. The most commonly seen symptoms are those caused by root infection of seedlings leading to a systemic infection (Friskop et al. 2009). Affected plants have chlorosis on the upper surface of leaves (either the entire leaf or the interveinal areas), with a white covering of spores on the underside of the leaves. Affected plants are severely stunted, and if they do not die within a few weeks, they will produce a head which is horizontal rather than vertical. The pathogen, by means of airborne spores, can also incite leaf spots typical of other downy mildews, with chlorotic spots on the upper surface, with a white coating of spores on the underside of each lesion. These “local lesions” do not lead to systemic infection and are of less impact as the affected leaves can be removed (Fig. 4a, b).

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Fig. 4 (a) Downy mildew infected plants showing stunting and leaf chlorosis. (b) Downy mildew infected leaves showing chlorosis on upper surface and white sporulation on bottom surface

Biology and epidemiology. Downy mildew is caused by the obligate pathogen Plasmopara halstedii (Farlow) Berlese & de Toni (syn. Peronospora halstedii Farlow). The host range of the causal pathogen includes sunflower and all wild Helianthus species, plus a number of related Asteraceae weeds and native plants, including Ambrosia, Bidens, Iva, Eupatorium, Silphium, Solidago, and Xanthium. P. halstedii is also reported to cause downy mildew on some ornamental genera such as Centaurea, Coreopsis, Dimorphotheca, Erigeron, Gerbera, Rudbeckia, Senecio, and Verbena; however, cross-pathogenicity studies to prove isolates from these can infect sunflower, and vice versa, have not been done. Plasmopara halstedii exists as physiological races on sunflower, and there is evidence that there are biotypes infective on specific genera, even though morphologically the pathogen isolates are indistinguishable. Plasmopara halstedii overwinters as resistant oospores, which germinate in water-logged soils to produce motile zoospores. The spores swim towards sunflower roots and infect young seedlings to produce a systemic infection. Infection of older plants will produce a club root without leading to systemic infection. Once the fungus becomes systemic and produces spores on the undersurface of leaves, these spores can be blown some distance and can initiate foliar infection (i.e., local lesions). As the plants die, the fungus produces oospores which can remain viable for years in the soil. The fungus and disease development is fostered by cool (4 h). Thus, long periods of dew or fog producing noninterrupted periods of free water on the plant is more conducive to infection than short periods of rain, regardless of the amount. Temperatures > 25  C and lack of free water are detrimental to ascospore germination/infection (Masirevic and Gulya 1992). The ascospores can be dispersed short distances (100 m) by wind currents, as well as by splashing water. The fungus quickly produces abundant white mycelia in all infected plant parts, and then sclerotia, which can be found externally on plant roots and the lower stem, inside the lower stem, and on the head. S. minor sclerotia, in contrast, germinate only to form mycelium, which initiates root infection. Thus, S. minor does not produce head rot since it produces no airborne ascospores. Sclerotinia sclerotiorum has an extremely broad host range, encompassing almost 400 plants in 278 genera of herbaceous plants with the Asteraceae, Cruciferae, Leguminosae, and Solanceaceae among the most important families. Important crop hosts include lettuce, beans, soybeans, canola, and sunflowers, plus many Asteraceae ornamentals. Sclerotinia minor has a slightly less broad host range of

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94 species in 66 genera. Important hosts include lettuce, Brassica crops, peanuts, and sunflower. Management. Diseases caused by either of the Sclerotinia species are very difficult to control, and the best management is aimed at prevention. • Cultural practices – Since infected plants can spread Sclerotina via root-to-root contact, lower plant densities will minimize wilt incidence (Nelson et al. 1989). To minimize head infection, anything which decreases foliage, increases air movement, or decreases head/leaf wetness will help, including lower nitrogen fertilization, lower plant densities, and furrow/drip versus overhead irrigation. Many different biocontrol/cultural practices have been shown to be partially effective at decreasing soil populations of sclerotia, and thus minimizing both root and head infections. Deep plowing, with a moldboard plow to completely invert the soil profile to >15 cm, will bury sclerotia into an aerobic environment where microbes should degrade the sclerotia (Mueller et al. 2002). However, there is conflicting data on this practice, and most experimental trials suggest that shallow burial leads to faster degradation of sclerotia (Ćosić et al. 2012; Subbarao et al. 1996, P. 48). Planting dense cover crops of cereal grains will produce the microenvironment conducive to apothecial formation, and if this crop is tilled under just prior to a susceptible crop, the net effect is to sap the sclerotia of energy/biomass sufficient to prevent production of more apothecia (Mueller et al. 2002). Rotation is of minimal use, since there are so many susceptible crops, but if several years of a nonhost monocot (for example, grasses or cereals) are planted, this also would hasten lowering the sclerotial soil population (Mueller et al. 2002). Once either stem rot or head rot are observed, there are no curative measures to save those plants. The best option at that point is to physically remove affected plants and dispose of them away from the field, thus minimizing soil contamination with more sclerotia. • Biological control – Many commercial biocontrol products are available, based on fungi such as Coniothyrium, Gliocladium, and Trichoderma, and bacteria such as Bacillus, and these applied as soil drenches immediately following a Sclerotinia infestation will hasten sclerotial degradation and shorten the interval between planting another susceptible crop. • Soil disinfestation – Preventative measures include using pasteurized soil, solarization, soil fumigation, or, where practical, flooding of fields for several weeks, all of which are aimed at decreasing the soil population of sclerotia. Biofumigation by planting Brassica cover crops and tilling them under will release isothiocyanates, which are toxic to a range of fungi (Griffiths et al. 2011). • Resistance – In certain sunflower hybrids, some progress has been made in developing partial (incomplete) resistance or tolerance to S. sclerotiorum. There are no hybrids available with complete resistance to either of the two Sclerotinia spp.

Diseases of Sunflower

2.15

27

Septoria Leaf Spot and Blight (Septoria spp.)

Geographic occurrence and impact. Septoria leaf spot is noted on sunflower throughout the world on every continent. The disease is most destructive in areas with heavy rainfall but is seldom observed in areas with drier climates. Under optimal disease conditions, premature leaf senesce and defoliation will result. Symptoms/signs. Septoria blight is characterized by angular to circular necrotic leaf spots reaching 10–15 mm in diameter. Lesions are tan to gray with dark brown margins, and may be surrounded by a yellow halo. Symptoms may be observed on cotyledons and leaves of young seedlings, but it is more common to see severe symptoms on older plants. One characteristic feature which distinguishes Alternaria and Septoria leaf spots is the presence of pycnidia in the lesions produced by Septoria spp., which appear as small, black dots, barely visible with the naked eye. In addition, Septoria lesions are generally much larger than Alternaria lesions. Lesions coalesce in time, turning the leaf necrotic and defoliating the plant from the bottom leaves up (Fig. 14a–b). Biology and epidemiology. Two species of Septoria are known to infect sunflower, Septoria helianthi Ellis and Kellerman found worldwide (Holliday and Punithalingam 1970) and Septoria helianthina Petrov and Arsenijevic, initially described in Yugoslavia (Petrov and Arsenijevic 1996). Both Septoria spp. are thought to be restricted to sunflower and wild Helianthus spp. Septoria spp. overwinter as pycnidia on infected plant residue. Spores ooze from the pycnidia when wet and are spread by splashing rain and wind. Initial infections occur on lower leaves, where the microclimate is more conducive, and the upper leaves become affected later. Rapid disease development is fostered by abundant rain and moderately high temperatures. The development of disease is arrested with hot, dry weather but can resume with the return of favorable conditions. Management • Cultural practices – Any cultural practice such as crop rotation and tillage of crop residue will lessen the carryover of inocula.

Fig. 14 (a) Septoria leaf blight. (b) Closeup (10X) of lesion showing the fungal pycnidia, visible as tiny black dots

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• Fungicides – Broad spectrum fungicides commonly used to control foliar diseases on other ornamental crops should be effective against these Septoria spp. • Resistance – Genetic resistance has been noted in oilseed germplasm (Block 2005; Carson 1985), but this has not been incorporated into commercial hybrids nor into ornamental varieties.

2.16

Sclerotium Blight (Sclerotium rolfsii Saccardo)

Geographic occurrence and impact. Sclerotium blight or wilt is found in Africa, Asia, Australia, Europe, and both Americas in areas that have tropical or semitropical climates (Punja 1985). In the United States, it is most frequently seen in Florida and the Gulf coast. Thus, ornamental sunflowers grown indoors in hot, humid conditions have the potential for significant losses due to this disease. Symptoms/signs. The main symptom of Sclerotium blight or “Southern blight” is a girdling lesion at the soil surface, generally appearing on plants as they reach the bud stage. This tan to dark brown lesion may have alternating light and dark bands, corresponding to the diurnal growth of the fungus. Under moist conditions, a white mycelial mat may form over the lesion and around the base of the plant. Sclerotia will form on the exterior of the lesion. The sclerotia are uniformly round, 0.5–2 mm, and brown, in contrast to the black sclerotia formed by Sclerotinia spp. The fungus will rot the pith but the rind will be intact, and thus, there seldom is lodging (Fig. 15). Biology and epidemiology. Sclerotium rolfsii is the asexual stage of this pathogen; the sexual stage Athelia rolfsii (Curzi) Tu and Kimbrough is seldom observed in Fig. 15 Sclerotium blight

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nature but can be produced in culture. The mycelium produced by this fungus is abundant, white, and cottony, and very similar to that produced by Sclerotinia spp. The sclerotia are the best means to distinguish S. rolfsii from the two Sclerotinia spp. Sclerotinia sclerotiorum produces sclerotia which are black, irregularly shaped but generally long and thin, and can reach up to 5 cm long. S. minor produces small round sclerotia (0.5–2 mm) black in color, while S. rolfsii sclerotia are similarly small (0.5-2 mm) but tan to dark brown. Soilborne sclerotia germinate to form a mycelium which infects stems at or slightly below the soil line (Aycock 1966). Sclerotial germination is favored by moist, but not saturated soil at temperatures from 25  C to 35  C. From initially infected plants, mycelia can move plant-to-plant via root contact or even grow on the soil surface under wet conditions. Sclerotia can survive for several years in soil, but do not withstand alternating freeze/thaw cycles, so this fungus is thus not seen in colder climates. Like the Sclerotinia spp., this pathogen has a wide host range of nearly 500 dicot plants, but it also infects some shrubs and trees. Management • Cultural practices – Crop rotation with nonhosts (for example, cereals and grasses) and deep burial of sclerotia (>15 cm) will aid disease control (Hagan 2004). Early season planting may have less disease due to cooler soil temperatures. • Soil disinfestation – Solarization or fumigation is more effective against sclerotia than soil-applied fungicides. • Fungicides – Seed or soil-applied fungicides for control of the seedling phase of the disease can be effective.

2.17

Verticillium Leaf Mottle/Wilt (Verticillium dahliae Klebahn)

Geographic occurrence and impact. The fungus causing Verticillium wilt is found globally, and with its wide host range and longevity in the soil, it can be a serious problem not just for sunflower but for any other floral crops grown in the same field/ soil. Symptoms/signs. External symptoms of Verticillium infection are more accurately described as a leaf mottle (Sackston et al. 1957). Leaf symptoms generally appear on plants as they reach bud stage, with symptoms appearing on lower leaves and progressing up the plant. Interveinal areas become chlorotic and then quickly die to produce dark brown-black areas surrounded by a yellow margin. These symptoms are often mistaken for the initial leaf symptoms of Phomopsis stem canker. However, with Phomopsis stem canker, the necrotic areas are centered on the veins, while with Verticillium wilt, they are always between the veins. The entire leaf surface will be affected with this mottle pattern, and eventually the first affected leaves will senesce and die. Wilting is not always observed, despite the inaccurate common name. Severely affected plants may die prior to maturity, producing small heads. Another way to confirm Verticillium infection is to cut the lower stem, either longitudinally or

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Fig. 16 (a) Verticillium leaf mottle/wilt. (b) Stem sections showing vascular browning caused by Verticillium

in cross section. The fungus will cause the vascular elements to be discolored brown. In severely affected plants, when the stem is cut open, the surface of the pith will be uniformly black, due to the presence of tiny microsclerotia (Fig. 16). Biology and epidemiology. Older literature incorrectly identified the causal fungus as V. albo-atrum, “microsclerotial form” but Verticillium dahliae is the correct causal agent (Sackston et al. 1957). Another fungus, Phialophora asteris (Dowson) Burge and Isaac (syn. Cephalosporium asteris Dowson), produces symptoms in sunflower very similar to Verticillium, and its morphology in culture is also similar, making identification difficult (Hawksworth and Gibson 1976). Verticillium dahliae produces conidia, but the microsclerotia are the propagules that overwinter and are responsible for infection. Verticillium dahliae is very slow growing in culture, and isolations made from diseased plant tissue are often mixed with faster growing contaminating fungi, further complicating pathogen identification. Verticillium dahliae has a host range of over 400 species of herbaceous and woody plants, including trees, vegetables, flowers, and weedy species. Susceptible flower crops include aster, chrysanthemum, cineraria, dahlia, geranium, gerbera, marigold, rose, snapdragon, statice, and stock, among others. Among field crops, potatoes and mint are severely affected. Verticillium dahliae overwinters in soil or plant residue as microsclerotia, and these remain viable for many years. Microsclerotia are stimulated to germinate by root exudates to form a mycelium which penetrates the roots, enters the vascular system, and eventually becomes systemic throughout the plant. Moist, but not water-

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logged soil, and temperatures between 21  C and 27  C are optimal for sclerotial germination and root infection. The fungus produces a toxin that is translocated to the leaves, which induces the foliar symptoms. Thus, isolation from symptomatic leaves may yield no Verticillium as the symptoms are the result of the toxin produced some distance away. Wilting, when it occurs, is the result of fungal mycelia and polysaccharides plugging the vascular system. If the mycelium reaches the head, the seeds will become infected. The fungus produces copious amounts of black microsclerotia internally, which gives the pith a charred, blackened appearance. The microsclerotia are smaller than those produced by Macrophomina phaseolina (charcoal rot) and cannot be seen with the naked eye or even a 10X lens. Management. As with other soilborne fungal pathogens forming sclerotia, management of Verticillium is challenging (Berlander and Powelson 2000). The sclerotia, despite their small size, are difficult to kill with fungicides; they exist in large numbers in the soil, and the fungus has a large host range. Since Verticillium is a serious problem on some high cash value crops like potatoes, there are commercial laboratories that assay soil samples to determine the presence and quantity of Verticillium microsclerotia. • Soil disinfestation – Thus, prevention is the first aim of a management program, and this starts with clean seed and pathogen-free soil, via pasteurization, solarization, or fumigation. Flooding the soil for 20 days significantly reduces the number of microsclerotia, as does soil acidification with aluminum sulfate. As mentioned for Sclerotinia wilt, the practice of biofumigation with Brassica cover crops has been shown to decrease the number of microsclerotia and the severity of the disease. • Cultural practices – While extra irrigation might be considered to help plants infected with a wilt pathogen, research has shown that frequent or excessive irrigation actually increases the severity of Verticillium. • Resistance – Genetic resistance is available (Hoes et al. 1973) and widely deployed in oilseed sunflower germplasm. The fungus exists as several races (Gulya 2007) and vegetative compatibility groups (VCGs) (García-Carnero et al. 2014), and this needs to be known so that breeders can select the appropriate single, dominant resistance gene.

2.18

White Rust (Albugo tragopogonis (DeCandolle) Gray = Pustula helianthicola Rost and Thines)

Geographic occurrence and impact. Albugo or white rust is a disease historically found primarily in the Southern Hemisphere (Argentina, Australia, South Africa) (Kajornchaiyakul and Brown 1976; Delhey and Kiehr-Delhey 1985) but more recently has been noted on both ornamental and oilseed sunflowers in Europe (Crepel et al. 2006; Thines et al. 2006), China (Chen et al. 2006), and, very infrequently, in the United States (Gulya et al. 2002b). The disease requires very specific environmental conditions, and the pathogen appears to be very host specific,

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Fig. 17 White rust on sunflower – leaves

and thus the disease is unlikely to appear year-in and year-out. While the disease is usually not devastating to the oilseed crop, the white blisters on the foliage detract from the value of ornamental sunflower. Symptoms/signs. The disease is usually manifested by raised, small to fairly large (1–2 mm or 5–10 mm) pustules or blisters on the upper leaf surface, with white sporulation on the underside of the leaf (Kajornchaiyakul and Brown 1976). With age, the Albugo pustules become necrotic and the infected tissue may fall out, leaving a “shot hole” appearance. The causal fungus can also cause similar white blisters on floral bracts, and may also initiate faintly black, water-soaked lesions on petioles and the stem. Stem lesions are usually internodal and may be colonized by secondary fungi, leading to lodging in severe cases (Krüger et al. 1999; Van Wyk et al. 1995) (Fig. 17). Biology and epidemiology. The fungus causing this disease is not a true rust and is actually more closely related to the downy mildews. The causal agent of white rust was classified as Albugo tragopogonis for over a century but was recently renamed Pustula helianthicola (Rost and Thines 2012). The fungus overwinters as oospores on infested plant debris and may also be seedborne (Viljoen et al. 1999). Oospores, the primary inoculum, are spread by rain and splashing water. They germinate to form motile zoospores that enter through leaf stomata. The white asexual sporangia produced on the underside of leaf pustules can be windblown to cause secondary infections on leaves, stems, petioles, and bracts. Oospores typically develop only in stem and petiole lesions, which cause the subtle, dark lesions. The disease is dependent on free water, from rain, dew, or irrigation. Optimum infection takes place at night between 10  C and 15  C, while optimal disease development is at temperatures between 20  C and 25  C. Thus, the disease is favored in environments with cool nights and warm days (Kajornchaiyakul and Brown 1976). Management • Cultural practices – Overhead irrigation at night will foster conditions conducive to foliar infection, and should be avoided.

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• Sanitation – Removal of plant debris from previous sunflower crops, either in the same or nearby fields, will diminish the possibility of infection. Leaves stripped prior to shipping cut sunflowers, if infected, should be disposed of outside the field, and preferably composted or removed entirely. • Fungicides – Seed-applied fungicides which control downy mildews (metalaxyl, mefenoxam, and strobilurins such as azoxystrobin) will provide early season control. Thereafter, foliar applications of these fungicides should provide protection against leaf infection (Lava et al. 2015). • Resistance – In oilseed sunflower germplasm, resistance to white rust has been noted and is controlled by multiple genes, each governing resistance in different plant parts. No effort has been made to incorporate any type of white rust resistance into ornamental sunflowers, although it has been noted to occur in one older commercial cultivar (“Abendsonne”). Many annual and most perennial Helianthus species appear to be resistant (Lava et al. 2015).

3

Bacterial and Phytoplasma Diseases

3.1

Apical Chlorosis and Bacterial Leaf Spot (Pseudomonas syringae pv. tagetis and Pseudomonas syringae pv. helianthi)

Geographic occurrence and impact. Apical chlorosis was first identified on marigold in Australia (Trimboli et al. 1978) and later on sunflowers in the United States in the early 1980s (Gulya et al. 1982). The causal bacterium is found worldwide on many Asteraceae flowers and weedy species. Bacterial leaf spot is caused by a very closely related bacterium, and it also is found worldwide (Piening 1976). Less is known about its host range, and it may be restricted solely to Helianthus. Apical chlorosis is more likely to be observed and cause alarm to growers due to its spectacular appearance, but since its symptoms are confined to the foliage, which is stripped off for cut flower production, its impact is lessened. Symptoms/signs. Apical chlorosis first manifests itself as a general chlorosis of the youngest leaf, either a portion or the entire leaf (Gulya et al. 1982). The affected area may be a pale yellow, or in extreme cases the leaf will be almost white. This symptom was initially confused with nitrogen deficiency, but the chlorosis is much more pronounced. The chlorosis will affect subsequent leaves as they develop, but the chlorotic leaves will not turn necrotic. When temperatures warm up, the newly emerging leaves will be normal green in color, but lower, chlorotic leaves will remain so (Fig. 18). Symptoms of bacterial leaf spot vary, depending upon the environment and sunflower cultivar. Lesions begin as small necrotic spots of varying size and shape and may be surrounded by chlorotic haloes. Under severe conditions the individual lesions coalesce to form large necrotic areas, giving the leaf a shothole appearance. Lesions may also form on petioles and stems, often leading to defoliation. Biology and epidemiology. Pseudomonas syringae pv. tagetis (Hellmers) Young, Dye, and Wilkie is the causal bacterium for apical chlorosis, while

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Fig. 18 Apical chlorosis

Pseudomonas syringae pv. helianthi (Kawamura) Young, Dye, and Wilkie causes bacterial leaf spot (Piening 1976). Recent molecular studies suggest that both bacteria may be identical, differing only in a toxin production gene present in P. syringae pv. tagetis. Pseudomonas syringae pv. tagetis has a wide host range within the Asteraceae, affecting ornamental flowers such as Tagetes and Zinnia, as well as many weedy genera such as Ambrosia, Cirsium, and Taraxacum (Rhodehamel 1985). Less research has been done with P. syringae pv. helianthi, but its host range appears to be restricted to Helianthus spp. Both bacteria can easily be identified as P. syringae pathovars based on the fluorescence under UV light of colonies grown on King’s B medium. Both bacteria survive in the soil and are seedborne. Apical chlorosis also persists on infected weedy hosts. Both are spread by splashing water and are typically seen early in the growing season during cool weather with wet soils. Apical chlorosis is typically seen in low areas of a field. Individual plants may be affected or there may be several affected plants in a row. Pseudomonas syringae pv. tagetis produces a toxin that interferes with chloroplast formation in developing leaves. In severe cases,

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the leaf may completely lack chlorophyll, but other pigments are not affected. Chlorotic leaves remain so throughout the life of the plant, but the appearance of new chlorotic leaves will cease as soils dry out and air temperatures warm up. Bacterial leaf spot is quite common, and its appearance is not restricted to cool early season and wet soil (Arsenijevic et al. 1994). Infection generally starts on lower leaves where the microclimate is more favorable for both fungal and bacterial infection. Management. Since there are no curative bactericides, there is no management option once symptoms are observed. • Cultural practices – In field situations, limiting irrigation to prevent water-logged soils and water splashing, especially during cool weather, will likely preclude the appearance of apical chlorosis. • Resistance – Differences in susceptibility to both bacteria have been noted in oilseed germplasm, but neither disease has been severe enough to warrant transferring that resistance into either oilseed hybrids or ornamental cultivars. • Sanitation – As the symptoms of apical chlorosis will cease when temperatures warm up, the damaged leaves can be stripped off ornamental sunflowers, and similarly with bacterial leaf spot.

3.2

Bacterial Stalk Rot/Head Rot (Erwinia = Pectobacterium spp.)

Geographic occurrence and impact. The bacteria causing stem and head rot are ubiquitous in soils and are present in every continent. Stem infections, though uncommon, would render ornamental sunflowers unsaleable. Head rot, although potentially serious on oilseed sunflower, does not occur in the bud stage, so it is unlikely to be observed on cut sunflowers but could be present in potted plants or homegrown sunflowers. Symptoms/signs. Bacterial stem rot and head rot can usually be recognized by smell as well as visual symptoms. The exterior of affected stems may have no discernible lesions or may have a blackened area, often centered around a petiole (Gudmestad et al. 1984). Upon splitting the stem open longitudinally, the pith will appear very water soaked, and if the infection has progressed long enough, there will be a strong odor of rotting potatoes. In time, due to gas produced by the bacteria, the stem will split open longitudinally and bacterial slime may ooze out, which often attracts flies and other insects. After this phase the stalks will either lodge or then may dry up and turn completely black, and the odor will be gone (Fig. 19). The bacterial head rot phase is characterized by symptoms that initially are identical with those caused by many fungal incited head rots. Small, depressed, watery, soft rotted areas develop on the back of the heads that enlarge and turn from tan to brown. Again there is a distinct odor of rotting potatoes, and if one grasps such a head, the soft mushy condition will be evident. Head infections may also produce copious amounts of slime and exudate, which may smell either of rotting potatoes, or in some cases, of alcohol, due to the presence of other secondary organisms such as

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Fig. 19 Bacterial stalk rot

yeasts. These exudates, combined with the bacterially produced gas, often look like foam on the head and stem. The smell and sugars in the exudates attract many insects, including flies, bees, and various beetles. Biology and epidemiology. Pectobacterium carotovorum (Jones) Waldee and Pectobaterium atrosepticum (van Hall) Lehmann and Neumann are both responsible, alone or in combination, for soft rots of sunflower stems and heads. These “softrotting bacteria” can affect many plants, causing rots of flowers, soft fruit, and vegetables. A species from Mexico, Pectobacterium cacticidum, whose primary hosts are cacti, was recently isolated from sunflower (Valenzuela-Soto et al. 2015). Both Pectobacterium spp. are soft-rotting bacteria that are found in soils and decaying plant debris (Charkowski 2015). They are very weak pathogens and require wounds to infect healthy tissue. These wounds can be mechanical damage, or caused by insects, birds, or hail. Free water and high temperatures speed the development of soft rot once the bacteria have colonized a wound. If insect larvae tunnel into the stems, they will not only vector the bacteria but will hasten its spread

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within a plant. Once the bacteria become established internally within the stem or head, the influence of air temperature and rainfall is negligible. Management. Bacterial diseases, especially caused by ubiquitous organisms, are hard to control, and prevention is the best approach. Controlling insects and birds to minimize wounds through which the bacteria gain entrance is a sound first step (Chattopadhyay et al. 2015). Ground irrigation rather than overhead will minimize the free water necessary for infection. Once an infected plant is noted, it should be physically removed and disposed of outside the greenhouse or field, which will eliminate the inoculum that feeding insects might spread.

3.3

Aster Yellows (Phytoplasma spp.)

Geographic occurrence and impact. Aster yellows has been previously confused with virus diseases, then classified as caused by a mycoplasma (now known as phytoplasmas) (Bertacinni and Duduk 2009). The causal agent causes a “yellows” disease on over 200 crops and is the most widely seen “yellows” on nursery crops. The complex of phytoplasmas affecting sunflower has been reported from North and South America, Europe, and Northern Africa. Symptoms/signs. Sunflower plants affected by “yellows” exhibit a range of symptoms including leaf chlorosis (of the entire leaf or in sectors), proliferation of secondary shoots, and various abnormalities of the head. There may be wedgeshaped sectors of the head showing hypertrophy, and bracts and ray petals may appear in the center of the head (Fig. 20). Biology and epidemiology. Aster yellows is caused by one or more closely related phytoplasmas. These organisms are smaller than bacteria, larger than viruses, and a few can, with difficulty, be grown in vitro. The phytoplasma causing aster yellows in sunflower and other crop is referred to as “Candidatus Phytoplasma asteris” (Harrison and Helmick 2008). Another phytoplasma causing similar symptoms has been studied in Argentina and tentatively termed “Candidatus Phytoplasma Fig. 20 Aster yellows

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Sunflower Phyllody” (Guzmán et al. 2014), and a third one was characterized in Iran (Salehi et al. 2015). Phytoplasmas are vectored by several genera of leafhoppers. The phytoplasma survives in perennial Asteraceae weed species and leafhopper vectors. Aster yellows multiplies within the body of the leafhopper and is transmitted through the insects saliva when feeding. The Aster yellows phytoplasma is believed to have a host range of more than 150 species, including many flower crops, vegetables (carrot, lettuce, potato, tomato), and even grains and grasses. Cool, wet weather seems to favor both the leafhopper vector and the disease. Management • Vector management – No chemicals are directly effective against phytoplasmas. Controlling the insect vectors, however, whether done by insecticides, biological control, or cultural practices (i.e., exclusion netting on greenhouse vents) will all but eliminate the disease. Removing weeds and volunteer sunflowers from the field and perimeter will reduce the inocula and vectors. • Cultural practices – Some of the more susceptible floral crops include Petunia, Tagetes, Viola, Chrysanthemum, Rudbeckia, and Salvia, and thus it would be wise not to plant these in the vicinity of sunflowers, and to monitor all susceptible crops closely for symptoms and leafhoppers. • Resistance – Aster yellows was once severe enough in Canada on oilseed sunflower to warrant research, and resistance was found within oilseed germplasm (Putt and Sackston 1970). Thus, it would be possible to transfer that level of resistance into ornamental sunflowers.

4

Viral Diseases

4.1

Sunflower Mosaic Virus

Geographic occurrence and impact. Sunflower mosaic virus is the only virus documented to occur naturally on sunflower in the United States, and currently is known nowhere else. Its range within the United States at the moment is restricted to southern Texas. As such it is classified as a quarantine pathogen by many importing countries, and while its impact upon ornamental sunflowers may not be great, the threat of restricted international movement of plants or seeds would be an economic hardship. Several other viruses have been confirmed on sunflowers in other countries, but their occurrence has usually been sporadic and limited to the specific countries. Thus, sunflower chlorotic mottle virus (SuCoMoV) has been found in Argentina (Giolitti et al. 2010), Pelargonium zonate spot virus (PZSV) also in Argentina (Giolitti et al. 2014), sunflower necrosis caused by tobacco streak virus (TSV) in India and Australia (Sharman et al. 2009; Prasada Rao et al. 2000), Bidens mottle virus (BiMV) in Florida (Wisler 1984) and Taiwan (Liao et al. 2009), plus

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Fig. 21 Sunflower mosaic virus

several other instances of virus diseases of undetermined origin (Loebenstein and Thottappilly 2003). Symptoms/signs. Sunflower mosaic is characterized by a mild mosaic pattern on leaves of young plants, becoming more pronounced, and then fading as the plant matures (Gulya et al. 2002a). The virus does not cause stunting, or any change in leaf or head morphology, nor does it cause ringspots (Fig. 21). Biology and epidemiology. The disease is caused by Sunflower mosaic virus (SuMV), a Potyvirus. Virus particles are long, flexuous rods. Their long size, along with characteristic pinwheel inclusion bodies, is diagnostic for Potyviruses. SuCoMoV is also a Potyvirus, while PZSV and TSV are Ilarviruses, with small, isometric virus particles. The Rio Grande river valley of southern Texas, where the disease is endemic, is subtropical, so that both wild sunflower and insect vectors can be found year-round. The virus was vectored experimentally by aphids (Myzus persicae and Capitphorus elaegni), both of which are common greenhouse inhabitants. Sunflower plants are optimally susceptible for 1 month, after which they become progressively less receptive to the virus. The virus was found to be seed transmitted. SuMV has a very narrow host range, consisting of Helianthus, Zinnia, and Sanvitalia. Other sunflower viruses, such as SuCoMoV, TSV, and PZSV, have much broader host ranges. Management. While resistance to SuMV and SuCoMoV has been noted, it has not been transferred to commercial oilseed hybrids or to ornamental cultivars (Gulya et al. 2002a; Lenardon et al. 2005). The most effective management for these sunflower viruses include the use of virus-free seed, minimizing insect vectors (aphids and thrips) through insecticide sprays or cultural methods, and removing

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infected plants. The use of silver reflective mulch in field plantings has been shown to repel aphids, as well as conserving soil moisture.

5

Nematodes

5.1

Root-Knot Nematodes (Meloidogyne Species)

Geographic occurrence and impact. Root-knot nematodes are found worldwide on many hosts but are especially serious on sandy soils and in warm climates. Countries where root-knot nematodes have had the greatest impact on sunflower are Egypt, Mozambique, South Africa, and Zambia in Africa (Bolton et al. 1989b; Lawn et al. 1988); Pakistan and India in Asia (Ravindra et al. 2015); and Greece and Italy in Europe (Tzortzkzkis et al. 2014; Zazzerini et al. 1997). In the United States, rootknot nematodes have been documented on sunflower in Florida and Texas (Rich and Dunn 1982; Isakeit 2011). There are currently 30 Meloidogyne species in the United States, of which five are known to parasitize sunflower experimentally. M. hapla is the most widespread, found in every state except Alaska, while M. incognita, M. arenaria, and M. javanica are most common in southern and coastal states, and M. chitwoodi is only found in the western states of WA, OR, CA, CO, UT, and NM, and in Mexico (Walters and Barker 1994). M. incognita appears to be most serious on sunflower. Lack of confirmation of root-knot nematodes on sunflower in a particular state may be due to the lack of nematologists to confirm their presence. Symptoms/signs. Symptoms of root-knot nematode infection may not be apparent aboveground, or the plant may display stunting, chlorosis, and daytime wilting. Affected plants may also have fewer and smaller leaves and flowers. Root galls are the belowground symptom (Fig. 22). The extent and size of root galls or “knots” depends upon the nematode population density, the species of Meloidogyne, and the host cultivar. Among ornamentals and field crops, sunflower is rated as highly susceptible (Crow 2014; Ferris et al. 1993). In the absence of galls, only a soil analysis in a nematology lab will confirm Meloidogyne as the causal agent, and be able to pinpoint the exact species. Biology and epidemiology. There are five Meloidogyne species documented to parasitize sunflower: M. arenaria (Neal) Chitwood; M. chitwoodi Golden, O’Bannon, Finley and Santo; M. hapla Chitwood; M. incognita (Kofoid and White) Chitwood; and M. javanica (Treub) Chitwood. Root-knot nematodes are sedentary endoparasitic nematodes, meaning they spend most of their life cycle within plant tissue. They hatch from eggs, stimulated by both temperature and exudates from plant roots. The juvenile nematodes migrate through the soil, penetrate the tips of fine roots, and the nematode’s head becomes embedded in the vascular bundle where it begins feeding and becomes immobile. Root cells around the nematode enlarge and form galls, which inhibit root function, leading to the aboveground symptoms. Female nematodes can produce eggs parthenogenetically, and up to several hundred eggs in a gelatinous mass can be found on the root surface. Generation time can vary from 20 to 45 days, allowing for multiple infections in a growing season.

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Fig. 22 Root-knot nematode damage on sunflower hybrid “954”, with smaller galls caused by Meloidogyne hapla (left) and larger galls caused by M. arenaria (right)

Management • Cultural – Depending on the root-knot species present, rotation with nonhosts (arugula, onion, sesame, sorghum-sudangrass) will decrease the nematode population but will not completely eliminate Meloidogyne (Dover et al. 2004; El-Ghonaimy et al. 2015; Kokalis-Burelle et al. 2013). Conversely, planting of “trap crops” of susceptible hosts, such as cowpea, and removing them completely, has also been shown to decrease damage on subsequent sunflower crops. Selecting preceeding crops in a commercial nursery which are least susceptible will minimize damage to sunflower (Creswell 2000). Solarization, in climates where feasible, will reduce nematode populations. • Chemical – While no nematicides are registered for use by home owners, several fumigants are commonly used in high-value annual crops and the nursery trade which are effective against all soilborne nematodes, including chloropicrin, 1,3-dichloropropene, metam sodium, and the nonfumigant ethoprop (Stapleton et al. 2014). • Genetic Resistance – Evaluations of oilseed sunflowers have demonstrated that there is considerable variation in susceptibility to Meloidogyne spp., but to date there have been no public releases of root-knot tolerant or resistant germplasm (Fabiyi and Atolani 2013; Rehman et al. 2006). • Biological – Several nematode-trapping fungi and antagonistic bacteria have been identified and shown to reduce, but not eliminate, populations of Meloidogyne spp. (Karssen et al. 2013; Duncan & Moens 2006). At least two

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mycopesticides, Myrothecium verrucaria (DITERA), and Paecilomyces lilacinus (MELOCON) are registered for use on ornamentals in the United States (Stapleton et al. 2014; Crow 2014).

5.2

Root-Lesion Nematodes (Pratylenchus Species)

Geographic occurrence and impact. Root-lesion nematodes, of the genus Pratylenchus, have a very wide host range and are found throughout the world, ranging from cool temperate to hot tropical climates (Davis and MacGuidwin 2005). They rank third behind root-knot and cyst nematodes as the nematodes of greatest economic impact on all crops worldwide. Their small size (0.4–0.7 mm long) relative to other plant parasitic nematodes enables them to survive in almost any texture soil. At least eight species have been found on sunflower worldwide, with P. penetrans the most widely distributed and important species. It interacts with wiltinciting fungi like Fusarium and Verticillium, thus augmenting the physical damage it causes (Castillo and Vovlas 2007). Symptoms/signs. Aboveground symptoms due to root-lesion nematodes are nonspecific, and like root-knot damage, are manifested by stunting and chlorosis, and wilting during hot days. Nematode damage in fields often occurs in random patches which are associated with higher populations of the nematodes. Root-lesion nematode feeding will cause brown lesions on the exterior of roots, while feeding inside roots will result in cracking and eventual rotting. Biology and epidemiology. Pratylenchus species documented on sunflower include P. alleni Ferris, P. brachyurus (Godfrey) Filipjev and Schuurmans Stekhove, P. crenatus Loof, P. hexincisus Taylor and Jenkins, P. penetrans (deMan) Filipjev, P. scribneri Steiner, P. thornei Sher and Allen, and P. zeae Graham. Pratylenchus spp. are usually migratory endoparasites, and remain vermiform throughout their life (Castillo and Vovlas 2007). Males only occur in some species, and thus reproduction is mostly by parthenogenesis. The life cycle can be completed in 28–56 days, depending upon the species, plant host and environment, so several generations can occur during a growing season. As with all plant-feeding nematodes, there are four juvenile life stages following the egg, and culminating in the adult. All juvenile and adults feed on roots, burrowing into the cortical tissue, and they can enter and leave roots during any life stage. Reports on how far they may travel range from 2 cm up to 1–2 m in the soil to reach other plants. Movement of soil and/or infected plant material provides a much wider dispersion. Pratylenchus overwinter in infected plants, including weedy hosts, in any life stage. Eggs can also dehydrate and become dormant and remain viable for over 2 years. Management. Cultural, biological, and chemical management recommendations for root-lesion nematodes are much the same as for root-knot nematodes, covered previously (Creswell 2000; Stapleton et al. 2014; Thompson et al. 2008). Differences in susceptibility to Pratylenchus in oilseed sunflower have been noted (Bolton and De Waele 1989a), but there have been no reports of resistance in any ornamental sunflower germplasm.

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43

Other Nematodes

Many other nematodes have been identified as causing damage on sunflower. These include dagger (Xiphinema), pin (Paratylenchus), spiral (Helicotylenchus), sting (Belonolaimus), stubby-root (Paratrichodorus), and stunt (Tylenchorhynchus) nematodes, among others (Bridge and Starr 2007). Identification of the causal nematodes always depends upon soil analysis by a nematology lab, as the aboveground symptoms are not nematode specific.

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Diseases of Zinnia Dorota Szopińska

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-Like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Alternaria Blight (Alternaria zinniae Pape) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Botrytis Blight, Gray Mold, Stem Canker (Botrytis cinerea Pers ex Pers) . . . . . . . . . . 2.3 Cercospora Leaf Spot (Cercospora zinniae Ellis & G. Martin) . . . . . . . . . . . . . . . . . . . . . 2.4 Fusarium Root Rot (Fusarium spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Pythium Root Rot (Pythium spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 Phytophthora Root and Crown Rot (Phytophthora spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7 Powdery Mildew [Golovinomyces cichoracearum (DC.) V.P. Heluta (Formerly Erysiphe cichoracearum DC. ex Merat); Euoidium sp.] . . . . . . . . . . . . . . . . . 2.8 Rhizoctonia Root Rot [Rhizoctonia solani Kühn (Teleomorph: Thanatephorus cucumeris (A.B. Frank) Donk)] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.9 Sclerotinia Stem Rot [Sclerotinia sclerotiorum (Lib.) de Bary] . . . . . . . . . . . . . . . . . . . . . 2.10 Sclerotium Stem Blight, Southern Blight [Sclerotium rolfsii Sac. (Teleomorph: Athelia rolfsii (Curzi) C.C. Tu & Kimbr.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Bacterial and Phytoplasma Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Apical Chlorosis [Pseudomonas syringae pv. tagetis (Hellmers) Young, Dye & Wilkie, syn. Pseudomonas tagetis (Hellmers)] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Bacterial Leaf Spot (Xanthomonas campestris pv. zinniae Hopkins and Dawson) . . . 3.3 Ralstonia Wilt (Ralstonia solanacearum (Smith 1896) Yabuuchi et al. 1996, comb. nov.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4 Aster Yellows (“Candidatus Phytoplasma Asteris”) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Viral Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Bromoviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Bunyaviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Caulimoviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4 Closteroviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5 Comoviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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D. Szopińska (*) Department of Phytopathology, Seed Science and Technology, Poznań University of Life Sciences, Poznań, Poland e-mail: [email protected] # Springer International Publishing Switzerland 2016 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florists' Crops Diseases, Handbook of Plant Disease Management, DOI 10.1007/978-3-319-32374-9_28-1

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4.6 Geminiviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7 Potyviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.8 Other Viruses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Nematode Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 Angular Spots on the Leaves (Aphelenchoides spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2 Root Knot (Meloidogyne spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract

Modern garden zinnias are widely cultivated ornamentals produced for cut flowers and flowerbeds. Diseases often reduce the quantity and quality of zinnia flowers, causing their production to be unprofitable. Three major, widespread diseases occurring on zinnias are: Alternaria blight (Alternaria zinniae), powdery mildew (Golovinomyces cichoracearum, formerly Erysiphe cichoracearum), and bacterial leaf spot (Xanthomonas campestris pv. zinniae). Other pathogens, such as Botrytis cinerea, Cercospora zinniae, Fusarium spp., Rhizoctonia solani, aster yellows phytoplasma, and several viruses may also be harmful to zinnias; however, their economic impact on zinnia production is less significant. Various management strategies for zinnia diseases currently studied include chemical, biological, and physical control and integration of these methods. Keywords

Alternaria zinniae • Golovinomyces cichoracearum • Xanthomonas campestris pv. zinniae • Integrated management

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Introduction

The genus Zinnia (family Asteraceae) consists of 19 species of annual or perennial plants originating in North and South America and particularly in Mexico (Spooner et al. 1991; Stimart and Boyle 2007). Among them common zinnia (Zinnia elegans Jacq., syn. Zinnia violacea Cav.) is the most widely cultivated species. Variation in ray floret morphology and color, plant height, and habit has led to development of many cultivars useful for landscape borders or cut flowers. The other two commonly planted species, Zinnia angustifolia Kunth (syn. Zinnia linearis Benth.) and Zinnia haageana Regel, are less conspicuous, characterized by a large number of small inflorescences and often with single whorl of ray florets (Linderman and Ewart 1990; Spooner et al. 1991; Stevens et al. 1993). The common zinnia is highly susceptible to three pathogens: Alternaria zinniae Pape (Alternaria blight), Golovinomyces cichoracearum (DC.) V.P. Heluta (powdery mildew), and Xanthomonas campestris pv. zinniae Hopkins and Dawson (bacterial leaf and flower spot), which may cause severe epiphytotics in zinnia, resulting in plant losses, decrease of ornamental value, or both. Zinnia angustifolia on the other hand is highly resistant to these pathogens and has been used as a source of resistance genes for Zinnia breeding programs (Boyle and Wick 1996; Jones and

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Strider 1979; Spooner et al. 1991). Management of the most important zinnia diseases is presented in this chapter. General management strategies, especially referring to widespread pathogens causing diseases of numerous crops, may be found in the introductory chapters.

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Fungal and Fungus-Like Diseases

2.1

Alternaria Blight (Alternaria zinniae Pape)

Geographic occurrence and impact. The pathogen is widely distributed. The disease has been reported in North and South America (Dimock and Osborn 1942; Gombert et al. 2001; Judd 1976; Palacios et al. 1991), Europe (Beaumont et al. 1958; Christova et al. 1964; Dias and Lucas 1978; Gambogi et al. 1976; Imre 1974; Łacicowa et al. 1979, 1991; Mowsiesjan 1976; Szopińska 2013, 2014a, b; Szopińska et al. 2012), Asia (Rao 1971; Wu and Yang 1992), and New Zealand (Dingley and Brien 1956). Symptoms/signs. The most conspicuous symptom of Alternaria blight is spotting of the foliage. Characteristic reddish-brown spots, sometimes with grayish-white centers, may also appear on stems and blossoms. The spots on the leaves are first circular and vary in size from 2 to 10 mm in diameter but soon enlarge and become irregular. The affected leaves become brown and desiccated. On blossoms and stems, the spots are relatively smaller, rarely more than 2 mm across. On the ray flowers, sporulation on the spots is often profuse and secondary infection may be abundant leading to darkening and withering of petals. On stems, internodal spots frequently show elongation and are usually superficial. Nodal lesions do not remain superficial and the distal parts of the affected stems may be killed by girdling at the nodes. Dark brown to black cankers with sunken centers may also appear near the soil line. Infected roots may turn dark gray, rot, and slough off resulting in wilting and death of the plant. Seedlings may also wilt and collapse (damp-off) (Fig. 1a, b). Biology and epidemiology. Alternaria spp. usually overseason as mycelium or spores in plant debris and seeds. Fungal sporulation and infection are enhanced by moderate temperatures. In the case of A. zinniae, the optimum temperatures for conidial germination in vitro and plant infection are 20–24  C and 22–26  C, respectively. Free moist is required for infection, but the pathogen survives better under dry conditions. The fungus can persist in the seeds for up to 6–7 years, although in plant debris usually not longer than 2 years (Rotem 1994). Seeds are a very important source of inoculum for Alternaria blight. Deeply seated, severe seed infections may lead to preemergence death of seedlings, while superficial seed infections usually cause diseases on plants after emergence (Gambogi et al. 1976). Management • Cultural practices – The seeds and transplants used for propagation should to be free of A. zinniae. Rotation to nonsusceptible crops for 2 years, control of susceptible weeds, plant spacing ensuring good air circulation, and proper time

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Fig. 1 Alternaria blight symptoms: first spots (left), irregular patches (right) (Photos D. Szopińska)

• •

•

•

and type of irrigation are the most important cultural control measures. Overhead irrigation as well as late afternoon and evening irrigation prolong the time during which the plants remain wet, and therefore favor spread of the disease (Gombert 1998). Sanitation – Crop debris from the planting area should to be removed and destroyed after harvest. Physical treatment – To eradicate the pathogen, the seeds can be treated with hot water at 51.5  C for 20 min (Beaumont et al. 1958; Lamboy and Call 2001). A high risk of germination percentage reduction, especially if older seeds are treated, is a reason why this approach is not widely use by the ornamental seed industry (Daughtrey and Benson 2005). Fungicides – The plants should be treated with protective fungicides at regular intervals (7–14 days), especially during warm wet weather or during warm dry weather if overhead irrigation is applied. Fungicides which are known to control Alternaria diseases include mancozeb, iprodione, chlorothalonil, copper, and triflumizole (Hagan 2009). Organic treatment – Szopińska (2013) observed that treating zinnia seeds with 1.0 % and 2.5 % lactic acid for 30 min decreased the percentage of seeds infected with A. zinniae from 56.0 % to 33.0 % and 23.0 %, respectively. Significant reduction of zinnia seed infestation with this pathogen was also observed if the seeds were treated with 6 % hydrogen peroxide (H2O2) for 60 min and 12 % H2O2 for 20 min. These treatments decreased seed infestation from 76 % to 28.0 % and 27.5 %, respectively (Szopińska 2014b).

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• Resistance – All Z. elegans selections are moderately to highly susceptible to Alternaria blight (Gombert et al. 2001), while Z. angustifolia is highly resistant to this disease (Hagan 2009). Therefore, colchicine-induced amphiploids have been developed in breeding zinnia plants resistant to A. zinniae (Terry-Lewandowski and Stimart 1983, 1985; Terry-Lewandowski et al. 1984). A new species of Zinnia marylandica, an artificial hybrid between Z. angustifolia and Z. elegans, was described and illustrated by Spooner et al. in 1991. Plants of this species exhibit high levels of resistance to A. zinniae and G. cichoracearum and moderate to high levels of resistance to X. campestris pv. zinniae (Terry-Lewandowski and Stimart 1983).

2.2

Botrytis Blight, Gray Mold, Stem Canker (Botrytis cinerea Pers ex Pers)

Geographic occurrence and impact. The pathogen is distributed worldwide and affects over 200 plant species in temperate and subtropical regions (Williamson et al. 2007). Botrytis cinerea has been reported on zinnias in Canada, Great Britain, Poland, the USA, and Venezuela (Bolton 1976; Kiecana and Mielniczuk 2010; Moore 1959; Palacios et al. 1991; Ruhl et al. 1987). Symptoms/signs. Large areas of petals, leaves, and stems turn brown. The infected plant parts develop a dusty gray mycelium during humid conditions (Fig. 2a, b). Biology and epidemiology. The pathogen can survive as mycelia and conidia or for extended periods as sclerotia in crop debris (Williamson et al. 2007). Botrytis cinerea produces tremendous quantities of airborne spores, which are moved around by wind or splashing water onto blossoms or young leaves. Cool temperatures (7–15  C) and high humidity (93 %) favor spore germination;

Fig. 2 Botrytis blight symptoms/signs: leaf blight (left) flower and stem blight with profuse sporulation (right) (Photos D. Szopińska)

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however, the disease may occur within wide range of temperatures. The pathogen invades host plant tissues directly or through natural openings or wounds (Daughtrey et al. 2000). Management • Cultural practices – Crucial for controlling B. cinerea is reducing free water on plant surfaces and lowering humidity (Daughtrey and Benson 2005). • Sanitation – Strict sanitation is required to prevent the disease from spreading. The dead or dying tissue from the plants and from the soil surface as well as old blossoms and leaves should to be removed and destroyed. • Fungicides and biocontrols – There is a high risk of resistance arising in B. cinerea if a chemical product is applied repeatedly. Since 1980s, there have been numerous reports about the resistance of B. cinerea to benzimidazoles (e.g., benomyl, carbendazim, thiophanate-methyl) and dicarboximides (e.g., iprodione, procymidone, vinclozolin) (LaMondia and Douglas 1997; Northover and Matteoni 1986; Vali and Moorman 1992; Yourman and Jeffers 1999). This phenomenon has also been recorded to a lesser extent in the case of the anilinopyrimidines (e.g., cyprodinil, pyrimethanil), the hydroxyanilide fenhexamid, and some multisite fungicides and fluazinam (Leroux 2004; Leroux et al. 2010). Therefore, application of multisite fungicides such as chlorothalonil, copper hydroxide, copper sulfate pentahydrate, and mancozeb as well as mixed spray programs have been advised (Daughtrey et al. 2000; Williamson et al. 2007). Fungicide resistance has also led to the intensive evaluation of microbial antagonists for control of gray mold. The greatest potential for control of B. cinerea has been shown by the fungi Trichoderma harzianum, Gliocladium roseum, and Ulocladium oudemansii; the bacteria Bacillus subtilis, Pseudomonas syringae, and Streptomyces griseoviridis; and the yeasts Candida oleophila and Pichia pastoris (Elad and Stewart 2004).

2.3

Cercospora Leaf Spot (Cercospora zinniae Ellis & G. Martin)

Geographic occurrence and impact. The pathogen has been reported on Z. elegans in India, Jamaica, Mauritius, Thailand, the USA, and Venezuela (Felix 1960; Palacios et al. 1991; Pande 1975; Pereira 2008; Phengsintham et al. 2013; Wehlburg 1969). Symptoms/signs. Cercospora leaf spot closely resembles Alternaria leaf spot, and symptoms of both diseases often occur together on the same leaf. Leaf symptoms include relatively large, round, reddish brown or dark purple spots, with light gray centers, which become ashen gray, papery, and brittle. Severely infected leaves become brown and desiccated. The dead tissues often crack and tear (Fig. 3). Biology and epidemiology. The fungus is favored by warm weather and high humidity. Conidia of C. zinniae germinate within 6 h after inoculation of the zinnia leaf surface. Germ tubes penetrate leaf tissue through the stomatal pores. Spots on the leaves appear 5–7 days after inoculation. Development of the first conidia begins

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Fig. 3 Cercospora leaf spot symptoms (Photo courtesy of R. J. McGovern)

on the spots at 9 days after inoculation. The spores are dispersed in the field by wind, insects, splashing water, and humans (Pereira 2008). Management • Cultural practices – Avoiding overhead irrigation and proper plant spacing are important to prevent spread of this pathogen. Crop rotation with hosts not affected by Cercospora zinniae is recommended. • Fungicides – Under field conditions mancozeb (0.2 %) followed by carbendazim (0.1 %) and chlorothalonil (0.2 %) reduced Cercospora leaf spot percent disease index from 53.62 (water spray) to 5.94, 13.36, and 18.26, respectively (Yadahalli et al. 1994). Similar results were recorded by Raghavendra Rao and Chacko (1986) and Madhumeeta and Shyam (1989).

2.4

Fusarium Root Rot (Fusarium spp.)

Geographic occurrence and impact. Several species of Fusarium, e.g., F. avenaceum, F. equiseti, F. culmorum, F. oxysporum, and F. solani have been isolated from diseased zinnia plants in Poland. Fusarium equiseti showed the highest

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pathogenicity towards zinnia seedlings (Kiecana and Mielniczuk 2010; Łacicowa et al. 1979). Symptoms/signs. Disease symptoms include reduction of growth, necrotic streaks on the hypocotyls, necrotic spots with yellow halos limited to the edge of leaves, and root rotting of severely infected plants. Fusarium spp. may also cause pre- and postemergence damping-off. Biology and epidemiology. Fusarium root rot is favored by high air and soil temperatures that exceed 23  C. At lower soil temperatures, disease symptoms may not occur, and even though the plant is diseased, it may appear to be healthy. The pathogens survive in the soil associated with plant debris but also directly in the soil as mycelium, spores, and, except in the case of F. avenaceum, as chlamydospores (especially in cooler regions). They spread by means of water and contaminated equipment. Growing mycelia invade the root system of healthy plants through wounds or at the point of formation of lateral roots. Management. The fungi are so persistent and widespread that control measures, such as crop rotation and soil sterilization, are of limited value. Some biological methods were proposed but chemical control is currently the only practical way to manage Fusarium spp. effectively in zinnia.

2.5

Pythium Root Rot (Pythium spp.)

Geographic occurrence and impact. Pythium spp. belongs to the kingdom Chromista, class Oomycetes. These pathogens have been reported on zinnia in the USA (Lewis et al. 1996; Lumsden and Locke 1989). Symptoms/signs. The pathogens attack roots which become dark and rot. Infected plants wilt very quickly. Pythium spp. may also cause pre- and postemergence damping-off. Biology and epidemiology. The pathogens from genus Pythium are distributed as zoospores with water, and long-term survival is enabled by chlamydospores and sexually produced oospores. Wet soil with poor aeration favors Pythium growth and development. Pythium spp. have wide range of hosts worldwide. Management • Biocontrol – Gliocladium virens controlled damping-off of zinnia caused by P. ultimum and R. solani in a nonsterile soilless mix. Disease control efficacy lasted for at least 2 m when G. virens was introduced with the pathogen inoculum and the mix was planted at intervals with zinnia seeds (Lumsden and Locke 1989). Alginate prills, formulated with the isolate GI-22 of G. virens and various food bases (wheat bran, corn cobs, peanut hulls, soy fiber, castor pomace, and chitin) also significantly reduced the damping-off of zinnia in a soilless mix caused by these pathogens (Lewis et al. 1996).

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Phytophthora Root and Crown Rot (Phytophthora spp.)

Geographic occurrence and impact. Phytophthora spp. also belong to the kingdom Chromista and class Oomycetes. These pathogens infest a wide range of crops, including zinnia (Moore 1959). Symptoms/signs. Symptoms include rot of roots and lower stems and dampingoff of seedlings. Infected plants wilt and die. Biology and epidemiology. Phytophthora spp. are disseminated as zoospores with water through irrigation and rain. Wet soil favor incidence of this disease. Longterm survival is by chlamydospores and sexually produced oospores. Management. Susceptible crops should be planted in pathogen free soil. Crop rotation with resistant or less susceptible crops is advised.

2.7

Powdery Mildew [Golovinomyces cichoracearum (DC.) V.P. Heluta (Formerly Erysiphe cichoracearum DC. ex Merat); Euoidium sp.]

Geographic occurrence and impact. Powdery mildews are common and widespread diseases, usually more abundant in semiarid regions than in areas of high rainfall (Kamp 1985). The disease caused by G. cichoracearum has been reported on zinnia in Egypt, India, Japan, New Zealand, the Republic of South Africa, the USA, and Venezuela (Amano 1986; Baker and Locke 1946; Dingley 1965; Gombert et al. 2001; Hegazi and El-Kot 2010a, b; Husain and Akram 1995; Kamp 1985; Linderman and Ewart 1990; Mir et al. 2012; Palacios et al. 1991; Ruhl et al. 1987; Schmitt 1955; Terry-Lewandowski and Stimart 1983; Tesfagiorgis 2008). A powdery mildew caused by Euoidium sp. has been reported on zinnia in Japan (Hoshi et al. 2013). High incidence and severity of powdery mildew on zinnias in some regions of Asia and North America appears to be a major factor for the declining commercial value of these plants (Mir et al. 2012; Spooner et al. 1991). Symptoms/signs. White fungal growth forms on the upper surface of leaves and may form on flower petals. Leaves die from base of plant upwards. Affected plants lose vigor and cease growth. Biology and epidemiology. The fungi causing powdery mildews are obligate parasites. The mycelium of most powdery mildew fungi grows only on the surface and produces usually short conidiophores. When environmental conditions become unfavorable, the fungus may produce sexual structures called cleistothecia, containing one or a few asci. The spores can be released, germinate, and cause infection even if there is no film of water on the plant surface as long as the relative humidity in the air is moderately high (Agrios 2005). Baker and Locke (1946) revealed that seed transmission of the pathogen is possible; however, primary inocula of G. cichoracearum for zinnia infections are likely to come from adjacent wild or cultivated Asteraceae (Boyle and Wick 1996).

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Management • Cultural practices – For powdery mildew, unlike Alternaria blight and bacterial leaf and flower spots, drip irrigation should to be avoided, because it may produce greater powdery mildew severity than overhead irrigation (Gombert 1998). • Fungicides – Currently, the only practical method for controlling powdery mildew on zinnias in commercial production is by applying synthetic chemicals. Fungicide applications must be started when the first sign of the white powdery colonies on the lower leaves appears and should to be reapplied according to the specifications on the fungicide label. Such fungicides as tebuconazole, chlorothalonil, triforine, propioconazole may be use to control powdery mildew (Hagan 2009). Kamp (1985) demonstrated that the use of a polymer-based antitranspirant significantly reduced powdery mildew on Z. elegans. The author suggested that the epidermal coating may interrupt pathogen development on the leaf surface by repelling the film of free water from the leaves and disorientation of pathogen germ tubes by changes in surface topology. Locke et al. (2006), on the other hand, observed delay of expression of powdery mildew on zinnia grown hydroponically in Hoagland’s solution fortified with silicon. Tesfagiorgis (2008) found that zinnia plants have the ability to accumulate high levels of silicon in the leaves, which may support an active mechanism of defense within the host plant. • Biological control and organic treatment. Hegazi and El-Kot (2010a) sprayed zinnia plants four times at 1 week intervals with 25 and 50 % culture filtrates of Trichoderma harzianum, Epicoccum sp., Streptomyces endus, as well as two plant extracts, miswak (Salvadora persica) and henna (Lawsonia inermis) to control powdery mildew. All applied treatments significantly decreased disease incidence and severity; however, Epicoccum sp. and T. harzianum at 50 % concentration followed by henna and miswak extracts inhibited spread of disease to the highest extent. The authors also observed a significant decrease of powdery mildew incidence and severity as well as improvement of plant growth parameters (i.e., plant height, number of branches per plant, leaf area, fresh and dry weights of shoots and roots, root length) when zinnia plants were treated with cinnamon, clove, and ginger oils at a concentration of 0.1 ppm (Hegazi and El-Kot 2010b). In organic farming use of preparations based on copper, organic, and mineral oils (e.g., neem oil, paraffinic oil) and Bacillus subtilis strain QST 713 are recommended (Hagan 2009). • Resistance – There are only a few Z. elegans cultivars that are resistant to G. cichoracearum (Gombert et al. 2001). Artificial hybrids between Z. angustifolia and Z. elegans, on the other hand, are highly resistant or immune to powdery mildew (Terry-Lewandowski and Stimart 1983, 1985; TerryLewandowski et al. 1984). This disease has rarely been noted on the Crystal and Star series as well as the hybrid Profusion series zinnias (Hagan 2009).

Diseases of Zinnia

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11

Rhizoctonia Root Rot [Rhizoctonia solani Kühn (Teleomorph: Thanatephorus cucumeris (A.B. Frank) Donk)]

Geographic occurrence and impact. Rhizoctonia diseases occur worldwide and cause losses in almost all crops. Rhizoctonia root, crown, and stem rot have been documented in zinnia in Australia, Poland, and the USA (Łacicowa et al. 1979; Lumsden and Locke 1989; Schisler et al. 1994). Symptoms/signs. The first sign of infection in seedlings is damping off. In older plants, the infection begins on the stem as dark-brown lesion below the soil line and then a cottony, white mycelium grows upward in the plant. Root discoloration and rot can also occur. Numerous small black sclerotia may be produced on all infected tissues. Biology and epidemiology. The pathogen overwinters as mycelium or sclerotia in the soil and in some species, including zinnia, may be carried in the seeds. Very young seedlings may be killed before or after emergence. The fungus spreads with rain and irrigation, tools contaminated with soil, and with infected seeds. High temperatures (about 35  C) and moderately wet soil favor infection. Management • Cultural practices – Disease-free seeds should be planted. • Biocontrol – Gliocladium virens controlled damping-off of zinnia caused by Pythium ultimum or R. solani in a nonsterile soilless mix (Lewis et al. 1996; Lumsden and Locke 1989).

2.9

Sclerotinia Stem Rot [Sclerotinia sclerotiorum (Lib.) de Bary]

Geographic occurrence and impact. The pathogen has been reported on zinnias in Argentina, Great Britain, Poland, and the USA (Grabowski and Malvick 2015; Kiecana and Mielniczuk 2010; Kiehr et al. 2010; Łacicowa et al. 1979; Moore 1959). Symptoms/signs. Sclerotinia sclerotiorum causes crown rot, bulb rot, stem rot, wilt, and death of a wide variety of plants. The typical early sign of Sclerotinia disease on zinnia is the appearance on the infected plant of a white fluffy mycelium in which variable sized sclerotia soon develop. The mycelium first develops on the base of stem. When the fungal infection develops and the stem rots, the foliage above the lesion wilts and dies. The sclerotia of S. sclerotiorum may be also formed inside the stem. Moreover, the pathogen is frequently responsible for damping-off of zinnia seedlings (Łacicowa et al. 1979). Biology and epidemiology. The pathogen overwinters as sclerotia in the soil and as sclerotia and mycelium on or within infected tissues of dead plants. The sclerotia germinate in the spring or early summer producing apothecia which discharge a large number of ascospores into the air. The spores reaching available sources of food (e.g., old plant parts) start to germinate and cause infection. Infection may also start

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from the soil when sclerotia produce mycelial strands which invade young plants (Agrios 2005). The pathogen may cause pre- and postemergence damping-off. Management • Cultural practices – Susceptible cultivars should be planted in well-drained soil. Weeds as a potential hosts should to be removed. Because sclerotia may survive in the soil, at least a 3-year rotation with non-susceptible crops is recommended.

2.10

Sclerotium Stem Blight, Southern Blight [Sclerotium rolfsii Sac. (Teleomorph: Athelia rolfsii (Curzi) C.C. Tu & Kimbr.)

Geographic occurrence and impact. Sclerotium rolfsii has been reported on zinnia in India, Japan, and New Zealand (Boesewinkel 1977; Ishii and Abiko 1997; Ramakrishnan 1930; Thakur 1969). Symptoms/signs. The pathogen attacks the stem bases, leaves turn yellow, the plant initially wilts, and then dies. A large number of white turning tan, mustard seed-sized sclerotia are formed on the infected tissue. Biology and epidemiology. Wet and warm weather favor disease progress. Optimal temperatures for fungal growth and disease development are 25–35o C. High relative humidity is required for germination of sclerotia (Mordue 1974). Sclerotia in the soil are the main source of this disease. Management • Cultural practices – Management strategies include a 3- to 4-year rotation with non-susceptible crops, raising the soil pH, and removal of crop debris (Mordue 1974). Additional fungal pathogens of zinnia. The following fungi have also been reported as pathogenic to zinnia: Alternaria carthami S. Chowdhury – in Taiwan (Wu and Chou 1995) Colletotrichum acutatum J. H. Simmonds – in India (Kulshrestha 1976) Colletotrichum falcatum Went. – in Venezuela (Palacios et al. 1991) Nigrospora sphaerica (Sacc.) E.W. Mason – in the USA (Meepagala et al. 2015) Sclerotinia minor Jagger – in Argentina (Kiehr et al. 2010)

3

Bacterial and Phytoplasma Diseases

3.1

Apical Chlorosis [Pseudomonas syringae pv. tagetis (Hellmers) Young, Dye & Wilkie, syn. Pseudomonas tagetis (Hellmers)]

Geographic occurrence and impact. This disease has been reported on zinnia in Australia and the USA (Gulya et al. 1982; Jutte and Durbin 1979; Trimboli et al. 1978).

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Fig. 4 Apical chlorosis symptoms caused by Pseudomonas syringae (Photo courtesy of T. J. Gulya)

Symptoms/signs. Leaf spots, sometimes with chlorotic haloes, and apical chlorosis are typical symptoms of this disease (Fig. 4). Biology and epidemiology. Pseudomonas syringae pv. tagetis is a Gram negative aerobic rod. The pathogen overseasons in infected plant debris in the soil. In sunflower, this pathogen is seed-borne, but it is not known if this type of transmission occurs in zinnia. Weeds as well as cultivated crops within the family Asteraceae may provide natural sources of inoculum. Wet weather favors spread of this disease. Management. A 3-year or longer rotation is recommended to avoid building up large soil populations of the bacterium. Plants should to be watered in a manner that keeps the leaves dry to inhibit spread of this disease.

3.2

Bacterial Leaf Spot (Xanthomonas campestris pv. zinniae Hopkins and Dawson)

Geographic occurrence and impact. The disease has been observed in many countries including Australia, Brazil, Bulgaria, Hungary, India, Italy, Korea, Malawi, Pakistan, Rhodesia, Sierra Leone, and the USA (Akhtar and Khokhar 1988; Bertus and Hayward 1971; Boyle and Wick 1996; Deighton 1957; Dimitrov and Tsoneva 1977; Hopkins and Dowson 1949; Jones and Strider 1979; Myung et al. 2012; Nannizzi 1929; Peregrine and Siddiqi 1972; Rangaswami and Gowda 1963; Robbs 1954; Sahin et al. 2003; Schwarczinger et al. 2008a, b; Sleesman et al. 1973; Strider 1973, 1976, 1979a, b, 1980; Terry-Lewandowski and Stimart 1983). Symptoms/signs. The pathogen can attack all aboveground parts of zinnia plants, causing necrotic lesions of leaves, stems, and flowers. The first symptoms of the disease are small (1–2 mm), dispersed, transparent spots encircled by broad yellowish haloes. Under wet conditions, the lesions slowly enlarge. The spots become angular to irregularly circular and develop a reddish center. The lesions may merge to form irregular dead areas (0.5–1.0 cm long), which often crack as they dry. During very humid weather, small brown spots may appear on the ray flowers. Severely infected flower heads are seriously disfigured and may completely decay (Fig. 5).

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Fig. 5 Bacterial leaf spot symptoms (Photo courtesy of R. J. McGovern)

Biology and epidemiology. The pathogen has been reported to overseason in diseased plant residue in the field and in or on seeds (Strider 1973, 1979a). Infected seeds are known to be the major means of long-distance dispersal and primary inoculum source of the pathogen (Strider 1979b). Xanthomonas campestris pv. zinniae is capable of spreading rapidly through warm, humid, and rainy weather. Infected zinnias develop characteristic spots on the leaves within 2 weeks. Jones and Strider (1979) observed that plants at a 29–31/21–23  C (max/min) temperature regime developed the most severe disease symptoms. Management • Cultural practices – The use of pathogen free seeds and transplants has been recommended for control of bacterial leaf spot (Strider 1979a, 1980). Whenever possible, plants should be irrigated in a manner that keeps the leaves dry to inhibit spread of this disease. • Sanitation – Plants with leaf spots should be discarded and diseased plant debris should be removed from the growing area. Washing hands after handling diseased plants or soil is recommended. Handling of wet foliage should be avoided. • Chemical control – Chemical control of bacterial leaf spot on zinnia is difficult if not impossible to obtain. Copper hydroxide may provide some protection from this disease. To be effective, treatments have to be applied on a weekly schedule, beginning when the spots first appear on the leaves (Hagan 2009). Bactericides are only marginally effective in controlling Xanthomonas spp. Strider in 1980 recorded the efficacy of captan as a bactericide for control bacterial leaf spot on zinnia. • Resistance – Most common zinnia cultivars as well as Z. haageana “Persian Carpet”, Z. angustifolia “Pixie Sunshine”, Z. tenuifolia “Red Spider”, and Z. pumilia “Cut and Come Again” are susceptible to X. campestris pv. zinniae (Hagan 2009; Gombert et al. 2001). However, hybrids of Z. elegans and Z. angustifolia exhibit moderate to high levels of resistance to this pathogen (Terry-Lewandowski and Stimart 1983). The Crystal and Star series zinnias are

Diseases of Zinnia

15

highly resistant or immune to bacterial leaf spots. The Profusion series zinnias are moderately resistant to this disease. Some spotting of the leaves in the lower canopy as well as occasional but unobtrusive leaf death has been seen on Profusion series zinnias in the field (Hagan 2009).

3.3

Ralstonia Wilt (Ralstonia solanacearum (Smith 1896) Yabuuchi et al. 1996, comb. nov.)

Geographic occurrence and impact. Ralstonia solanacearum (formerly Pseudomonas solanacearum) causes a severe and devastating wilt to many economically important crops around the world (Hayward 1991). This disease has been reported on zinnia in Australia, India, and Malaysia (Abdullah 1992; Papdival and Deshpande 1978; Thammakijjawat et al. 2001). Strider et al. (1981) reported that a strain of R. solanacearum isolated from geranium was also pathogenic on zinnia. Ralstonia solanacearum is a pathogen of quarantine importance. Symptoms/signs. Infected young plants die rapidly. On older plants, symptoms appear first on the youngest leaves or one-sided wilting and stunting is observed. Finally plants wilt completely and die. The vascular tissues of stem and roots turn brown (Agrios 2005). Biology and epidemiology. According to host range, R. solanacearum strains have been classified into five races (Buddenhagen and Kelman 1964; Hayward 1991). A zinnia strain of R. solanacearum originating in Australia has been classified as race 1 (Thammakijjawat et al. 2001). This race attacks all the solanaceous and many non-solanaceous crops and appears as a sudden wilt. The bacteria overwinter in plant debris and spread in soil with water and then enter plants through wounds made in roots by insects, nematodes, or by handling. Management. There are no chemicals or biological agents to effectively control this bacterium. Infected plants have to be discarded as soon as possible. Pathogenfree propagating plant material is the main way to avoid problems with Ralstonia. Soil and equipment should to be disinfested.

3.4

Aster Yellows (“Candidatus Phytoplasma Asteris”)

Geographic occurrence and impact. This disease has been reported on zinnia in Canada and India (Rao et al. 2012; Singh et al. 2011; Wang and Hiruki 2001). Symptoms/signs. The pathogen causes general yellowing and dwarfing of the plant, abnormal production of shoots (formation of witches’ brooms), and malformation of organs (greening or sterility of flowers, shortening of the internodes). The disease reduces yield and may lead to more or less rapid dieback, decline, and death of the plant. Biology and epidemiology. Taxonomically, phytoplasmas are member of the class Mollicutes and are currently classified within the provisional genus “Candidatus Phytoplasma” based primarily on 16S rDNA sequence analysis

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(Marcone 2014). The phytoplasmas detected in zinnia have been placed in the subgroup II “Candidatus Phytoplasma asteris” (16SrI-B) (Rao et al. 2012; Wang and Hiruki 2001). Phytoplasmas are prokaryotic organisms that have no cell walls and can rarely be grown on artificial nutrient media. Most plant mollicutes are vectored by leafhoppers (Cicadellidae) but some by planthoppers (Fulgoroidea). Phytoplasmas are generally present in the sap of a small number of sieve tubes. Insects become vectors when feeding on leaves and stems of infected plants. The phytoplasma then multiplies and infects various body organs of insect vectors, a process that can take 1–3 weeks. When infected leafhoppers feed on healthy plants, the phytoplasmas in their salivary glands are injected into the plant phloem and disease symptoms develop in 10–40 day. A leafhopper carrying the aster yellows phytoplasma can continue infecting plants for the rest of its life. Outbreaks of aster yellows may be expected in cool and wet summers, which favor both leafhoppers and phytoplasma development (Agrios 2005). Management. There are currently few effective control strategies for aster yellows. Infected plants should to be promptly removed and destroyed. Insect vectors should to be monitored and controlled systematically, because they are essential for dispersal of the pathogen and long-term survival in nature. Management of aster yellows includes chemical and biological control of leafhoppers, repellence and disorientation of insects in the field, and avoidance of plant production near infected crops or alternate hosts.

4

Viral Diseases

4.1

Bromoviridae

Geographic occurrence and impact. This family contains five genera of viruses: Alfamovirus, Bromovirus, Cucumovirus, Ilarvirus, and Oleavirus. Several members of these genera are distributed worldwide. Four viruses from the Bromoviridae have been reported in zinnia: Alfalfa mosaic virus (AMV, genus Alfamovirus) in Israel, Cucumber mosaic virus (CMV, genus: Cucumovirus) in India, Iran, and the USA, Tomato aspermy virus (TAV, genus Cucumovirus), and Asparagus virus 2 (AV2, genus Ilarvirus) in Japan (Fujisawa et al. 1983; Kameya-Iwaki et al. 1996; Nitzany and Cohen 1960; Price 1935; Raj et al. 1997; Shahmohammadi et al. 2015). According to Brunt et al. (1996–), zinnia is also susceptible to the following viruses from this family: Peanut stunt virus from genus Cucumovirus and Elm mottle virus, Humulus japonicus virus, Plum American line pattern virus, Prune dwarf virus, Prunus necrotic ringspot virus, and Tobacco streak virus from genus Ilarvirus. Symptoms/signs. All viruses detected in zinnia produce similar symptoms: leaf mosaic and distortion, stunting, and flowers malformation (Fig. 6). Biology and epidemiology. Aphids (Hemiptera: Aphididae) and thrips (Thysanoptera: Thripidae) are main vectors of Bromoviridae. Aphis gossypii,

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Fig. 6 CMV symptoms (Photo courtesy of R. J. McGovern)

A. craccivora, A. fabae, and Myzus persicae are responsible for CMV transmission in zinnia (Nitzany and Cohen 1960). All cucumoviruses are transmitted by aphids in noncirculative (nonpersistent) manner. They may be also transmitted in some species by seeds or mechanically by handling of the plants in the field or greenhouse (e.g., CMV). Seed transmission by zinnia seeds has been reported only for Asparagus virus 2 (Fujisawa et al. 1983) Management. Controlling insect vectors, avoiding growing near infected crops, and removing weeds that serve as hosts help to control viral diseases. Vector populations may be reduced by chemical and biological methods and various physical barriers. Use of reflective mulches disorients the vectors and protects the plants.

4.2

Bunyaviridae

Geographic occurrence and impact. Tospoviruses make up one genus of the viruses within this family, including several economically important species with an extremely wide host range. The main member of this genus – Tomato spotted wilt virus (TSWV) – has been detected in zinnia in Bulgaria and Mexico (Bakardjieva et al. 1998; Morales-Díaz et al. 2008). Symptoms/signs. Typical symptoms on zinnia are chlorotic rings and spots on the leaves, wilting of the leaves, and plant death. Biology and epidemiology. Tospoviruses are transmitted by thrips (e.g., Frankliniella occidentalis, F. fusca, F. schultzei, Thrips tabaci, T. palmi, etc.). The virus is acquired from infected plants by thrips larvae after at least 30 min of feeding. The thrips transmits the virus to healthy plants for the rest of its life. Management. Refer to the Bromoviridae management section above and introductory chapter 3, “▶ Insect Management for Disease Control in Florists’ Crops.”

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Caulimoviridae

Geographic occurrence and impact. Dahlia mosaic virus DMV belonging to the genus Caulimovirus has been reported on zinnia in the USA (Kitajima et al. 1969). Symptoms/signs. The symptoms range from none to occasional chlorotic spots to severe mosaic and stunting. Biology and epidemiology. DMV can be transmitted by 16 different aphid (Hemiptera: Aphididae) species, mechanical inoculation, and through the seed (not recorded for zinnia). Management. Refer to the Bromoviridae management section above and introductory chapter 3, “▶ Insect Management for Disease Control in Florists’ Crops.”

4.4

Closteroviridae

Geographic occurrence and impact. Two viruses belonging to genus Crinivirus have been reported in zinnia in Taiwan: Tomato chlorosis virus (ToCV) and Tomato infectious chlorosis virus (TICV) (Tsai et al. 2004). According to Brunt et al. (1996), zinnia is also susceptible to two other viruses from this genus: Beet pseudoyellows virus and Lettuce infectious yellows virus. Symptoms/signs. Pronounced yellowing symptoms on the lower leaves of plants, similar to those caused by nitrogen deficiency; the brittleness of the discolored leaves; and occasional upward leaf rolling were observed on infected zinnia plants. Biology and epidemiology. Criniviruses are confined to the phloem and phloem parenchyma cells. TICV is transmitted exclusively by the whitefly Trialeurodes vaporariorum, whereas ToCV is vectored by T. vaporariorum, and another whitefly, Bemisia tabaci. Management. Refer to the Bromoviridae management section above and introductory chapter 3, “▶ Insect Management for Disease Control in Florists’ Crops.”

4.5

Comoviridae

Geographic occurrence and impact. Two nepoviruses have been reported on zinnia: Tobacco ringspot virus (TRSV) in Japan and the USA and Tomato black ring virus (TBRV) in Kenya (Iizuka 1973; Kaiser et al. 1978; Moore and McGuire 1968). According to Brunt et al. (1996–), zinnia is also susceptible to the following viruses from this family: Cowpea mosaic virus from genus Comovirus and Arabis mosaic virus, Artichoke vein banding virus*, Caraway latent virus*, Cherry leaf roll virus, Dogwood mosaic virus*, Peach enation virus*, Croton yellow vein mosaic virus, and Soybean crinkle leaf virus from genus Nepovirus. (*Species not recorded currently in the ICTV Master Species List). Symptoms/signs. Common symptoms of plant infection by these viruses are mosaic and ring spot, sometimes accompanied by systemic necrosis.

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Biology and epidemiology. Tobacco ringspot virus is transmitted by the nematode Xiphinema americanum and also nonspecifically by insects and mites such as: Aphis gossypii, Myzus persicae, Melanopus sp. Epitrix hirtipennis, Thrips tabaci, and Tetranychus sp. Iizuka (1973) reported 5 % transmission of TRSV by zinnia seeds. This virus causes systemic infection in susceptible cultivars, moving from infected leaves to the tips of stems and into roots. Movement from roots to leaves is uncommon (Moore and McGuire 1968). Kaiser et al. (1978) detected seed transmission of TBRV in 1 seedling infected out of 4 tested. Management. Refer to the Bromoviridae management section above and introductory chapter 3, “▶ Insect Management for Disease Control in Florists’ Crops.”

4.6

Geminiviridae

Geographic occurrence and impact. These viruses are responsible for a significant amount of crop damage worldwide. Four species in genus Begomovirus were detected in zinnia: Ageratum enation virus (AEV) in India, Ageratum yellow vein China virus (AYVCNV) and Alternanthera yellow vein virus (AlYVV) in Vietnam, and Tomato yellow leaf curl virus (TYLCV) in China. Two other species Zinnia leaf curl virus (ZLCV) identified in India and Zinnia mosaic virus recorded in India and Spain are not recorded currently in ICTV Master Species List and their taxonomic position is unsure (Ha et al. 2008; Huertos 1953; Jabri et al. 1985; Kumar et al. 2010; Li et al. 2013, 2014); Maritan et al. 2004; Panday and Tiwari 2012; Verma and Singh 1973). Zinnia plants are also susceptible to the following viruses from this family: Beet curly top virus belonging to genus Becurtovirus and two species from genus Begomovirus: Croton yellow vein mosaic virus and Soybean crinkle leaf virus (Brunt et al. 1996–). Symptoms/signs. Symptoms caused by AEV, AYVCNV, TYLCV, and ZLCV have been described as leaf curling, yellow mosaic, and stunting (Figs. 7 and 8). Zinnia mosaic symptoms have been characterized as a strong mosaic, stunting, decrease of the number of flowers, and color breaking. Biology and epidemiology. Begomoviruses are transmitted circulatively and nonpropagatively by the whitefly Bemisia tabaci. Once the virus is acquired, the whitefly remains viruliferous for life. Management. Refer to the Bromoviridae management section above and introductory chapter 3, “▶ Insect Management for Disease Control in Florists’ Crops.”

4.7

Potyviridae

Geographic occurrence and impact. Potyviruses include many of the viruses causing some of the most severe and economically important plant diseases. Five species belonging to this genus have been identified in zinnia: Bidens mottle virus (BiMoV) and Sunflower mosaic virus (SuMV) in the USA, Sunflower chlorotic

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Fig. 7 Ageratum enation virus symptoms (Photo courtesy of Kumar et al. 2010)

Fig. 8 Tomato yellow leaf curl virus symptoms (Photo courtesy of Li et al. 2014)

mottle virus (SuCMoV) in Brazil, and Vanilla distortion mosaic virus (VDMV) and Zinnia mild mottle virus* in India (Balaji et al. 2014; Gulya et al. 2002; Logan et al. 1984; Maritan et al. 2004; Padma et al. 1972, 1974). According to Brunt et al. (1996–), zinnia is also susceptible to many other viruses from this family: Sweet potato mild mottle virus from genus Ipomovirus and Bean

Diseases of Zinnia

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yellow mosaic virus, Beet mosaic virus, Bidens mosaic virus, Carrot thin leaf virus, Celery mosaic virus, Clover yellow vein virus, Lettuce mosaic virus, Nasturtium mosaic virus*, Patchouli mosaic virus, Pea seed-borne mosaic virus, Pepper veinal mottle virus, Plum pox virus, Primula mosaic virus, Tobacco etch virus, Tropaeolum 2 virus, Turnip mosaic virus, Watermelon mosaic 2 virus, and Wisteria vein mosaic virus belonging to genus Potyvirus. (*Species not recorded currently in ICTV Master Species List). Symptoms/signs. Diseases caused by potyviruses appear primarily as mosaics, mottling, chlorotic rings or color breaking on foliage, flowers, and stems. Many of these viruses may cause severe stunting of young plants, stem malformations, and necroses of various tissues. Biology and epidemiology. Potyviruses are spread by aphids (Hemiptera: Aphididae) in a noncirculative (nonpersistent) manner and several are transmitted through seeds (not recorded for zinnia). These viruses overseason in perennial cultivated and weed hosts. Management. Refer to the Bromoviridae management section above and introductory chapter 3, “▶ Insect Management for Disease Control in Florists’ Crops.”

4.8

Other Viruses

According to the VIDE Database (Brunt et al. 1996–), zinnia is also susceptible to the following viruses: Family Flexiviridae, genus Carlavirus: Artichoke latent S virus*, Butterbur mosaic virus*, Cassia mild mosaic virus*, Pea streak virus*, Poplar mosaic virus; genus Potexvirus: Asparagus 3 virus, Cymbidium mosaic virus, Foxtail mosaic virus, Plantain X virus Family Luteoviridae, genus Luteovirus: Beet mild yellowing virus, Beet western yellows virus, Subterranean clover red leaf virus Family Rhabdoviridae, genus Nucleorhabdovirus: Sonchus yellow net virus Family Secoviridae: Strawberry latent ringspot virus Family Tombusviridae, genus Carmovirus: Cucumber leaf spot virus, Galinsoga mosaic virus; genus Dianthovirus: Carnation ringspot virus; genus Necrovirus: Lisianthus necrosis virus*, Tobacco necrosis virus; genus Tombusvirus: Cymbidium ringspot virus, Pepper Moroccan virus, Tomato bushy stunt virus Family Virgaviridae, genus Tobamovirus: Maracuja mosaic virus, Odontoglossum ringspot virus, Ribgrass mosaic virus Genus Sobemovirus: Lucerne transient streak virus Genus Ourmiavirus: Ourmia melon virus (*Species not recorded currently in ICTV Master Species List).

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5

Nematode Diseases

5.1

Angular Spots on the Leaves (Aphelenchoides spp.)

Geographic occurrence and impact. Foliar nematodes are widespread pathogens of ornamentals grown in greenhouses, nurseries, and in the field. There are three species of economic importance: A. besseyi, A. fragariae, and A. ritzemabosi. Aphelenchoides besseyi prefers warmer climates, while A. fragariae and A. ritzemabosi are often found in temperate zones (Kohl 2011). All three species can cause angular spots on Z. elegans leaves; however, A. ritzemabosi has been the most prevalent. Aphelenchoides spp. have been reported on zinnias in India, New Zealand, Poland, and the USA (Boesewinkel 1977; Crossman and Christie 1936; Gill and Sharma 1979; Gill and Uppal 1979; Gill and Walia 1980; Kohl 2011; Madej et al. 2000). Symptoms/signs. Infected plants appear stunted and dwarfed. Infected leaves become shrunken, occasionally accompanied by discoloration, with blotches and chlorotic patches turning into brown and white-yellow areas limited by the veins. Biology and epidemiology. Water is necessary for the movement and dispersal of foliar nematodes; however, spread from the direct contact of an infected leaf with uninfected plant tissue is possible. Adult and fourth-stage juvenile Aphelenchoides spp. enter leaf tissue through stomata on the leaf undersurface where feeding and reproduction occurs. The nematodes feed by piercing neighboring cells with their stylets. Eggs are laid within healthy, green sections of the leaf tissues. The endoparasitic feeding results in the disintegration of the spongy parenchyma and palisade cells; however, nematodes also feed ectoparasitically on stems, buds, and flowers. Males are necessary for reproduction. Fertilized females are able to lay eggs even after emergence from months of dormancy in an anhydrobiotic state. Each female lays approximately 32 eggs, which hatch in 4 days. The second stage juveniles reach reproductive maturity in 6–7 days. The life cycle of A. ritzemabosi can be completed in 14 days, with 5 days for embryonic development and another 5 days for maturation. Adults and fourth-stage juveniles are able to overwinter in an anhydrobiotic state within desiccated plant tissue (dried leaves, dormant buds, but not in plant roots) and can survive for several months to up to 3 years. Low air temperature and high relative humidity increase nematodes populations in leaves (Kohl 2011). Management. Foliar sprays of quinalphos and methyl parathion at 0.05 % applied five times at 10-day intervals effectively controlled A. ritzamabosi on Z. elegans, reducing both the symptoms of infestation and the final nematode population (Gill and Walia 1980). However, due to environmental concerns and toxicity, these nematicides are no longer available. Many other nematicides were proposed (e.g., chlorfenapyr); however, the most effective chemical controls will be those that are truly systemic within the plants (Kohl 2011). Integration of chemical methods with cultural practices such as sanitation, removal of crop debris from the planting area, management of leaf wetness by reducing overhead irrigation, and increasing air flow through a crop are necessary to manage foliar nematodes (LaMondia 1999).

Diseases of Zinnia

5.2

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Root Knot (Meloidogyne spp.)

Geographic occurrence and impact. Root knot nematodes have been reported on zinnias in Pakistan, Taiwan, and the USA (Jabri et al. 1985; Mc Sorley and Frederick 1994; Tsay et al. 2004). Symptoms. Infected plants are stunted and poorly growing with yellowing leaves. Infected root systems show characteristic knots or galls. Biology and epidemiology. Zinnia elegans exhibited some galling in response to Meloidogyne incognita, but incidence of galling increased when the plants were infected with Zinnia mosaic virus (Jabri et al. 1985). Mc Sorley and Frederick (1994) found that Z. elegans cv. Scarlet was nearly free of galling from M. incognita and M. arenaria but was susceptible to M. javanica (3,400 eggs per plant). Tsay et al. (2004) also observed that some cultivars of Z. elegans and Z. hageana were highly and moderately resistant to M. incognita, while Z. angustifolia was more susceptible. The life cycles of Meloidogyne spp. differ slightly (De Guiran and Ritter 1979). The life cycle of M. incognita can be completed in 37 days. Second-stage juveniles penetrate root tips, occasionally invading roots in the zone of root elongation. Invasion of nematodes results in development of giant cells in the meristematic, cortical, and xylem tissues of the root and galling of roots occurs. Third- and fourthstage juveniles and young females occur after about 6–8 and 15 days, respectively. Adult females are observed after 20 days and egg laying begins after 25 days. Meloidogyne spp. are mostly spread by water and through equipments contaminated with infested soil. Management. Control of root knot nematodes is difficult and options are limited, especially in field production. Equipment should to be frequently cleaned to prevent dissemination of the nematodes from field to field. In greenhouses soil, disinfestation may be obtained by steam heat treatment, solarization, fumigation, or chemical drenches. • Biocontrol – The biological control agent Paecilomyces lilacinus strain 251 has been successfully applied in several crops to control M. incognita (Anastasiadis et al. 2008; Hashem and Abo-Elyousr 2011; Kiewnick and Sikora 2006). • Resistance – There have been several reports about the resistance of various zinnia cultivars against root knot nematodes (Mc Sorley and Frederick 1994; Tsay et al. 2004).

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Raghavendra Rao NN, Chacko CJ (1986) Evaluation of fungicides for the control of foliar diseases of marigold and zinnia. Pestology 14:14–15 Raj SK, Chandra G, Singh BP (1997) Some Indian strains of Cucumber mosaic virus (CMV) lacking satellite RNA. Indian J Exp Biol 35(10):1128–1131 Ramakrishnan TS (1930) A wilt of Zinnia caused by Sclerotium rolfsii. Madras Agric J 16:511–519 Rangaswami G, Gowda SS (1963) On some bacterial diseases of ornamentals and vegetables in Madras State. Indian Phytopathol 16:74–85 Rao VG (1971) An account of the fungus genus Alternaria Nees from India. Mycopath Mycol Appl 43(3–4):361–374 Rao GP, Tiwari AK, Singh M, Chaturvedi Y, Srivastava R, Madhupriya (2012) Characterization of an isolate of ‘Candidatus Phytoplasma asteris’ infecting Zinnia elegans in India. Phytopathogenic Mollicutes 2(1):33–36. doi:10.5958/j.2249-4669.2.1.004 Robbs CF (1954) Bacterias fitopatogenicas do Brasil [Phytopathogenic bacteria from Brazil]. Agronomia, Rio de Journal 13:265–282 (In Portuguese) Rotem J (1994) The genus Alternaria: biology, epidemiology and pathogenicity. APS Press, The American Phtopathological Society, St. Paul, 326 pp Ruhl GE, Latin RX, Pecknold PC, Scott DH (1987) A compilation of plant disease and disorders in Indiana. Proc Indiana Acad Sci 97:121–127 Sahin F, Kotan R, Abbasi PA, Miller SA (2003) Phenotypic and genotypic characterization of Xanthomonas campestris pv. zinniae strains. Eur J Plant Pathol 109:165–172. http://www.oardc. ohio-state.edu/sallymiller/images/Pathogen_Identification-Sahin_2_2003.pdf Schisler DA, Neate SM, Masuhara G (1994) The occurrence and pathogenicity of Rhizoctonia fungi in South Australian plant nurseries. Mycol Res 98(1):77–82 Schmitt JA (1955) The host specialization of Erysiphe cichoracearum from zinnia, phlox and cucurbits. Mycologia 47(5):688–701. doi:10.2307/3755579 Schwarczinger I, Vajna L, Süle S (2008a) First report of bacterial leaf and flower spot of Zinnia elegans caused by Xanthomonas campestris pv. zinniae in Hungary. Plant Pathol 57:367. doi:10.1111/j.1365-3059.2007.01720.x Schwarczinger I, Vajna L, Süle S (2008b) Reappearance of bacterial leaf and flower spot on Zinnia elegans caused by Xanthomonas campestris pv. zinniae in Europe. Acta Phytopathol Entomol Hung 43(1):63–68. doi:10.1556/APhyt.43.2008.1.8 Shahmohammadi N, Dizadji A, Koohi Habibi M, Nateqi M (2015) First report of Cucumber mosaic virus infecting Bougainvillea spectabilis, Coleus blumei, Kalanchoe blossfeldiana and Zinnia elegans in Iran. J Plant Pathol 97(2):394. doi:10.4454/JPP.V97I2.041 Singh M, Chaturvedi Y, Tewari AK, Rao GP, Snehi SK, Raj SK, Khan MS (2011) Diversity among phytoplasmas infecting ornamental plants grown in India. Bull Insectol 64 (Supplement):69–70. http://www.bulletinofinsectology.org/pdfarticles/vol64-2011-S069-S070singh.pdf Sleesman J, White DG, Ellett CW (1973) Bacterial leaf spot of zinnia. A new disease in North America. Plant Dis Rep 57(7):555–557. http://babel.hathitrust.org/cgi/pt?id=uc1. 31175001263642;view=1up;seq=9 Spooner DM, Stimart DP, Boyle TH (1991) Zinnia marylandica (Asteraceae: Heliantheae), a new disease-resistant ornamental hybrid. Brittonia 43(1):7–10. http://pubag.nal.usda.gov/pubag/ downloadPDF.xhtml?id=2321&content=PDF Stevens S, Stevens AB, Gast KLB, O’Mara JA, Tisserat NA, Bauernfeind R (1993) Commercial specialty cut flower production. Zinnia. Cooperative Extension Service, Manhattan. online http://www.ksre.ksu.edu/bookstore/pubs/mf1079.pdf Stimart DP, Boyle TH (2007) Zinnia. Zinnia elegans, Zinnia angustifolia. In: Anderson NO (ed) Flower breeding and genetics: issues, challenges and opportunities for the 21st century. Springer, Dordrecht, pp 337–357 Strider DL (1973) Bacterial leaf and flower spot of zinnia in North Carolina. Plant Dis Rep 57 (12):1020. http://babel.hathitrust.org/cgi/pt?id=uc1.31175001263642;view=1up;seq=486 Strider DL (1976) An epiphytotic of bacterial leaf and flower spot of zinnia. Plant Dis Rep 60 (4):342–344. http://babel.hathitrust.org/cgi/pt?id=uc1.31175001303299;view=1up;seq=370

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Strider DL (1979a) Detection of Xanthomonas nigromaculans f. sp. zinniae in zinnia seed. Plant Dis Rep 63(10):869–873. http://babel.hathitrust.org/cgi/pt?id=uc1.31175005867711;view=1up; seq=377 Strider DL (1979b) Eradication of Xanthomonas nigromaculans f. sp. zinniae in zinnia seed with sodium hypochlorite. Plant Dis Rep 63(10):873–876. http://babel.hathitrust.org/cgi/pt?id=uc1. 31175005867711;view=1up;seq=377 Strider DL (1980) Control of bacterial leaf spot of zinnia with captan. Plant Dis 64(10):920–922. http://www.apsnet.org/publications/plantdisease/backissues/Documents/1980Articles/PlantDis ease64n10_920.pdf Strider DL, Jones RK, Haygood RA (1981) Southern bacterial wilt of geranium caused by Pseudomonas solanacearum. Plant Dis 65(1):52–53. doi:10.1094/PD-65-52 Szopińska D (2013) The effects of organic acids treatment on germination, vigour and health of zinnia (Zinnia elegans Jacq.) seeds. Acta Sci Pol Hortorum Cultus 12(5):17–29. http:// wydawnictwo.up.lublin.pl/acta/hortorum_cultus/2013/streszczenia2013_5/02%20Szopinska% 20Hort%2012_5_%202013.pdf Szopińska D (2014a) Alleviation of Zinnia elegans Jacq. seed deterioration using hydrogen peroxide and organic acids. Ecol Chem Eng S 21(2):309–326. doi:10.2478/eces-2014-0024 Szopińska D (2014b) Effects of hydrogen peroxide treatment on the germination, vigour and health of Zinnia elegans seeds. Folia Hort 26(1):19–29. doi:10.2478/fhort-2014-0002 Szopińska D, Tylkowska K, Deng CJ, Gao Y (2012) Comparison of modified blotter and agar incubation methods for detecting fungi in Zinnia elegans Jacq. seeds. Seed Sci Technol 40:32–42 Terry-Lewandowski VM, Stimart DP (1983) Multiple resistance in induced amphiploids of Zinnia elegans and Z. angustifolia to three major pathogens. Plant Dis 67:1387–1389. http://www. apsnet.org/publications/plantdisease/BackIssues/Documents/1983Articles/PlantDisease67n12_ 1387.pdf Terry-Lewandowski VM, Stimart DP (1985) The inheritance of resistance to powdery mildew in interspecific hybrids and induced amphiploids of Zinnia elegans Jacq. and Z. angustifolia HBK. Euphytica 34(2):483–487. http://link.springer.com/article/10.1007/BF00022945 Terry-Lewandowski VM, Bauchan GR, Stimart DP (1984) Cytology and breeding behavior of interspecific hybrids and induced amphiploids of Zinnia elegans and Zinnia angustifolia. Can J Genet Cytol 26(1):40–45. doi:10.1139/g84-007 Tesfagiorgis HB (2008) Studies on the use of biocontrol agents and soluble silicon against powdery mildew of zucchini and zinnia. PhD thesis. School of Agricultural Sciences and Agribusiness, Faculty of Science and Agriculture, University of KwaZulu-Natal, Pietermaritzburg, Republic of South Africa. pp. 182. http://siliconconference.org.za/fotos/Habtom%20PhD%20Thesis% 20Final.pdf#page=158 Thakur RN (1969) Sclerotium root rot of Zinnia elegans from Jammu and Kashmir. Labdev J Sci Tech 6(B):119–120 Thammakijjawat P, Thaveechai N, Kositratana W, Chunwongse C, Frederick RD, Schaad NW (2001) Genetic analysis of Ralstonia solanacearum strains from different hosts in Thailand using PCRrestriction fragment length polymorphism. Kasetsart J Nat Sci 35:397–408. http:// www.thaiscience.info/Article%20for%20ThaiScience/Article/2/Ts-2%20genetic%20analysis% 20of%20ralstonia%20solanacearum%20strains%20from%20different%20hosts%20in%20thai land%20using%20pcr-restriction%20fragment%20length%20polymorphism.pdf Trimboli D, Fahy PC, Baker KF (1978) Apical chlorosis and leafspot of Tagetes spp. caused by Pseudomonas tagetis Hellmers. Aust J Agri Res 29:831–839. doi:10.1071/AR9780831 Tsai WS, Shih SL, Green SK, Hanson P, Liu HY (2004) First report of the occurrence of Tomato chlorosis virus and Tomato infectious chlorosis virus in Taiwan. Plant Dis 88(3):311. doi:10.1094/PDIS.2004.88.3.311B Tsay TT, Wu ST, Lin YY (2004) Evaluation of Asteraceae plants for control of Meloidogyne incognita. J Nematol 36(1):36–41. http://www.ncbi.nlm.nih.gov/pmc/articles/PMC2620738/ pdf/36.pdf

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Vali RJ, Moorman GW (1992) Influence of selected fungicide regimes on frequency of dicarboximide-resistant and dicarboximide-sensitive strains of Botrytis cinerea. Plant Dis 76:919–924. doi:10.1094/PD-76-0919 Verma VS, Singh S (1973) Zinnia leaf curl virus. Die Gartenbauwissenschaft 38(20), No. 2:159–162. http://www.jstor.org/stable/43387245?seq=1#page_scan_tab_contents Wang K, Hiruki C (2001) Use of heteroduplex mobility assay for identification and differentiation of phytoplasmas in the aster yellows group and the clover proliferation group. Phytopathology 91:546–552 Wehlburg C (1969) Two leaf spot diseases of zinnia. Plant Pathology Circular 86, Florida Department of Agriculture, Division of Plant Industry. https://www.freshfromflorida.com/content/ download/11093/142849/pp86.pdf Williamson B, Tudzynski B, Tudzynski P, Van Kan JAL (2007) Botrytis cinerea: the cause of grey mould disease. Mol Plant Path 8(5):561–580. doi:10.1111/J.1364-3703.2007.00417.X Wu WS, Chou JK (1995) Chemical and biological control of Alternaria carthami on zinnia. Seed Sci Technol 23:193–200 Wu WS, Yang YH (1992) Alternaria blight, a seed-transmitted disease in Taiwan. Plant Pathol Bull 1:115–123 Yadahalli KB, Kulkarni S, Anahosur KH (1994) In vitro and in vivo evaluation of fungicides against leaf spot of Zinnia caused by Cercospora zinniae Ell. and Mart. Karnataka J Agric Sci 7 (3):363–365. http://14.139.155.167/test5/index.php/kjas/article/viewFile/5996/6223 Yourman LF, Jeffers SN (1999) Resistance to benzimidazole and dicarboximide fungicides in greenhouse isolates of Botrytis cinerea. Plant Dis 83:569–575. doi:10.1094/ PDIS.1999.83.6.569

Diseases of Azalea Robert G. Linderman

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-Like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Phytophthora Root Rot, Dieback, and Wilt (Phytophthora spp.) . . . . . . . . . . . . . . . . . . . . . 2.2 Pythium Root Rot (Pythium spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Cylindrocladium Blight and Wilt (Cylindrocladium scoparium Morgan) . . . . . . . . . . . . 2.4 Leaf and Flower Gall (Exobasidium spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Rhizoctonia Root Rot and Web Blight (Rhizoctonia spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Genetic Abnormalities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Witches’ Broom . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract

Florist azaleas (Rhododendron obtusum (Lindl.) Planch) are grown worldwide in greenhouses as a potted flower plant. They belong to the evergreen group but vary in their cold hardiness, so they must be protected against cold winter conditions. Many cultivars have been developed based largely on their horticultural traits, especially flower size and color. Some cultivars are prone to develop witches’ broom, thought to be a genetic disorder. However, most florist azaleas are susceptible to a number of diseases, but little attention has been paid to their resistance/susceptibility through breeding. Of concern to growers are diseases caused by species of Cylindrocladium, Pythium, Phytophthora, Rhizoctonia, and occasionally Exobasidium. Many of the diseases of florist azaleas, such as infections caused by Cylindrocladium, occur during the propagation phase of

R.G. Linderman (*) Plant Health, LLC, Corvallis, OR, USA e-mail: [email protected] # Springer International Publishing Switzerland 2016 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florist's Crops Diseases, Handbook of Plant Disease Management, DOI 10.1007/978-3-319-32374-9_29-1

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rooting cuttings, with surviving, infected-but-symptomless plants later succumbing to the disease.

Keywords

Phytophthora spp. • Pythium spp. • Cylindrocladium spp. • Exobasidium spp. • Rhizoctonia spp. • Witches’ Broom

1

Introduction

Evergreen azaleas originated in Japan and are classified in the genus Rhododendron, subgenus Tsutsutsi. These grow indoors and outdoors. Most azaleas bloom in spring, although evergreen florist’s azaleas can be forced into bloom under greenhouse conditions for gift giving at any time. Evergreen azaleas vary in cold hardiness with the particular cultivar, but the group spans US Department of Agriculture hardiness zones 5 through 9. A significant component of the florist industry involves the sale of potted evergreen azaleas (Rhododendron obtusum (Lindl.) Planch). Plants are grown in production greenhouses for protection against winter cold, since they vary in cold hardiness. Accordingly, they can be “forced” into flower any time by manipulating the environmental conditions to set flower buds that will bloom predictably to match sales for special occasions. Many cultivars have been selected, mainly for their predictable and extraordinary flowering, with variation in flower size and color. Cultivar development has paid little attention to disease susceptibility/resistance. Some diseases, such as Cylindrocladium leaf spot, blight, and/or root rot, have caused tremendous losses in the nursery industry over the years. Other diseases, common to many other greenhouse and nursery crops, include root rots caused by species of Pythium, Phytophthora, and Rhizoctonia. Some foliage and flower diseases, such as leaf and flower gall, caused by species of Exobasidium occur mainly on landscape plants and rarely on florist azaleas, mainly because it is readily controlled by judicious removal of galls and application of chemical sprays. Most of the more serious diseases are caused by soilborne pathogens that infect newly rooted cuttings. Surviving plants are infected but remain symptomless until some later time when poor growth, dieback, and mortality occur. Soilborne root pathogens are much more challenging to manage because of the multiple ways they can invade the greenhouse cultural system and the limitations on effective management. Nonetheless, numerous methods have been identified to block entry, largely by eliminating the pathogens from the sources. For example, sanitation of containers with heat, treatment of irrigation water, using pathogen-free growth media, chemically sanitizing cutting propagation material, and maintaining strict sanitation practices within the greenhouse environment can be effective measures for preventing entry into the greenhouse. Some pathogens like Cylindrocladium spp. are especially troublesome because they can disperse spores (inoculum) via an aerial phase within the greenhouse environment.

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Fungal and Fungus-Like Diseases

2.1

Phytophthora Root Rot, Dieback, and Wilt (Phytophthora spp.)

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Geographic occurrence and impact. Phytophthora spp. are known to infect azaleas and rhododendrons worldwide. These fungus-like organisms (Phylum Oomycota and genetically related to the brown algae) infect many other plants within the Ericaceae family as well (Jones and Benson 2001; Linderman and Benson 2014). Interestingly, some of those plants occur as wild species, but the diseases do not occur in natural settings, even when the pathogen is present, for unknown reasons. They cause root rot, crown canker, and branch dieback, depending on the species. Some species can cause all of those symptoms, while others are confined to the roots and crown. Most of the Phytophthora diseases occur in landscapes or in nursery field plantings, in ground or in containers. When the diseases occur in the nurseries, significant economic losses result. Beyond the nursery, after shipment and sale, further disease development, even mortality, may occur in the hands of the consumer. Those occurrences are frequently a result of earlier undetected infections or contamination that develops after time and/or a change in environmental and soil conditions. Sometimes, infected liner plants appear healthy and are shipped to other wholesale nurseries for growing on, and with further development of the infections, symptoms appear. Symptoms/signs. Root infections may develop at any time in the life of florist azaleas, from cutting propagation to field and landscape plantings. Most frequently involved are P. cinnamomi Rands and P. nicotianae Breda de Haan (syn. P. parasitica Dastur), but in some areas, other species such as P. citricola Sawada are recovered. Infected roots are brown and become necrotic. When enough roots are infected, the root system becomes impaired, and as a result leaves may become chlorotic and then necrotic but rarely wilt. Those leaves generally drop off the plant, causing partial defoliation (Fig. 1). Leaves on infected plants, as well as the plants themselves, are also often smaller than with healthy plants. With some Phytophthora species, root infections progress to the lower stem tissue where lethal crown cankers occur. These cankers are revealed by removal of the bark, exposing the internal wood which appears brown. Biology and Epidemiology. Root, crown, stem, and foliage infections caused by a number of species of Phytophthora can occur on florist azaleas when sufficient inoculum is present and environmental conditions are conducive. Those infections occur as a result of inoculum being disseminated by air movement, or by water splash of spores from the soil to aboveground tissues landing on susceptible sites. Free water is required for spore germination and infection. The source of inoculum (zoospores, sporangia, chlamydospores, or oospores, depending on the species) in greenhouse culture of florist azaleas is often unknown. Temperature conditions in the greenhouse systems are generally in the right range for infections by Phytophthora spp., as with Pythium spp. infections. So the reduction of infections by environmental conditions that might occur outdoors does not apply for florist azaleas.

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Fig. 1 Mortality of azalea (left) caused by Phytophthora citricola (R. G. Linderman)

Temperature is modified by heating and cooling to enhance plant growth. When flower bud setting is involved, temperatures are usually lowered. This allows plants destined for the florist trade to be programmed for forcing in time for special holiday sales. The source of Phytophthora spp. inoculum varies, depending a lot on the practices of the nursery. For soilborne, root-infecting pathogens, the medium, recycled containers, irrigation water, and plant propagules used for cutting propagation could be sources. Strict sanitation practices within the greenhouse, as well as the propagation area, must be maintained. Soilless growth media are used in growing florist azaleas, so they are least likely to be a source. Use of well or municipal water is also not likely to be a source. Likely sources are reused containers, recycled irrigation water, and source of propagation cuttings. Management. The primary management strategy for Phytophthora-caused diseases of florist azaleas is prevention. Eliminating the pathogen from the major sources of irrigation water, the growth medium, the containers, and even the cuttings can go a long way to preventing these diseases. Thereafter, application of preventative chemical fungicides provides some protection against invasion from a number of sources, including greenhouse workers. The chemicals used for Phytophthora control are the same as for Pythium control, and they generally target only those genera and some other fungus-like pathogens, but not fungal pathogens. There are preventative chemicals such as products containing etridiazole, mefenoxam, cyazofamid, fenamidone, and phosphonates. The insensitivity of some species of Pythium to mefenoxam has also been reported for Phytophthora spp. However, there can be variation in effectiveness of specific chemicals to different Phytophthora spp. (Linderman and Davis 2008a). Rotation or combinations with other chemicals should be practiced, as with management of Pythium spp.

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In addition to chemical agents, there are some biological control agents that target certain species of Phytophthora and Pythium. Species of Trichoderma and Streptomyces have been shown to have that activity but usually have not been tested on a range of pathogen species. Suppressive soils that prevent or limit some root diseases, including those caused by species of Phytophthora, have been reported. Often suppression is created by incorporating mixtures of microbes with great diversity into the growth medium. Within that microbial population are members with specific capacity to block the development of specific diseases. Some have the capacity to block sporangium development, and therefore reduce zoospore release (Linderman et al. 1983). Some are able to lyse the pathogen mycelium, while others produce other antibiotic compounds that inhibit growth of the pathogen in soil. With production of florist azalea, however, such practices generally are not used.

2.2

Pythium Root Rot (Pythium spp.)

Geographic occurrence and impact. Roots of many plant species can be infected by species of Pythium. Most often Pythium spp. are known as seedling damping-off pathogens, but they are root pathogens of many woody plants, including florist azaleas (Ivors 2014). They do not cause immediate death of plants but cause root rot that can debilitate plants to various degrees, depending on the time and extent of infection of the root system. Young rooted azalea cuttings can be killed if enough of the root system is involved. That will depend on when the pathogen begins to infect, relative to the rooting process. If infection occurs after a substantial number of roots have formed, the infections might not be detected, and the plants become infectedbut-symptomless until the percentage of infected roots increases. A number of species of Pythium can be involved in azalea root rot. Some may be more pathogenic than others (Weiland et al. 2013). Pythium spp. might also be secondary invaders to infections by species of Phytophthora. The net result and impact of Pythium root infections, therefore, is difficult to determine. If growing conditions in the greenhouse are favorable, Pythium infections might go unnoticed until some other stress occurs on the plants. Then the combination of Pythium root rot and stress causes the plants to show symptoms on the foliage, such as leaf chlorosis, small leaves, even dieback of young branches. Examination of the roots and detection of root rot suggest the role that Pythium played in the syndrome. Such plants, however, do not make it into the florist trade because they are discarded. Symptoms/signs. A range of symptoms of Pythium root rot can occur, including failure of cuttings to root, rooting followed by root rot, young plant mortality, stunting of plants at later growth stages, chlorosis of foliage, wilting under water deficiency, loss of vigor, necrotic larger roots, and leaf drop (Linderman and Benson 2014) (Fig. 2). Examination of the root ball of affected plants, compared to nonaffected healthy plants, usually shows a greatly reduced size. Often it is difficult to distinguish many of the aboveground symptoms from a nutritional disorder. Examining the root ball is critical. Over-watering or drought episodes may hasten

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Fig. 2 Symptoms of Pythium root rot of azalea: Left photo showing greater root browning of plant on left. Right photo showing aboveground symptoms of smaller plant, smaller leaves, and off-colored foliage, compared to plant on the-left (R. G. Linderman)

the development of aboveground symptoms, but plants without Pythium root rot generally show no or fewer symptoms. Biology and epidemiology. Identifying the species of Pythium involved in root rot of azalea is extremely difficult, even for the specialist. Ten species have been identified on rhododendrons and azaleas, but those affecting florist azaleas have not. It is possible, even likely, that more than one species can be involved. Further, some species/strains can be more pathogenic than others (Weiland et al. 2013). There can be many sources of inoculum of Pythium species that can invade the florist azalea production system: (1) Used and recycled containers have been shown to harbor Pythium species, even after rigorous washing. Baiting residual contaminated medium from propagation flats or larger containers reveals that Pythium spp. are present. It generally is impossible to predict just how pathogenic each might be without inoculation experiments. The variability of species or strains within a crop has been demonstrated, however (Weiland et al. 2013). (2) Contaminated media is another potential source of inoculum leading to Pythium root rot. Most growers rely on the soilless medium they buy or mix themselves being pathogen-free. However, Pythium has been found in some peat mosses, albeit infrequently. (3) Irrigation water is a likely source of Pythium where water is being recycled in the summer months. However, often well water is used in propagation, and it is believed to be pathogenfree. Water in irrigation reservoirs should be treated in order to prevent inoculum from being delivered into production pots. Generally chlorine compounds, such as hypochlorite or chlorine dioxide, or even chlorine gas have been used for that purpose. (4) Insect vectors, such as fungus gnats, have been suggested to carry propagules of Pythium from an infected pot to healthy pots. However, there is some degree of controversy in this regard, in that some studies show that the pathogen does not carry over from the larval to the adult stage that flies. However, there is no doubt that the larvae transmit the pathogen from root to root within a pot. Pythium root rot often occurs in combination with some environmental stress, such as high soluble salts in the medium, over-watering, under-watering (drought),

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or poor drainage. Water saturation of the medium can allow the pathogen to produce sporangia and then zoospores spread from root to root. Those conditions also stress plants in a physiological manner that makes them more susceptible, even for cultivars that generally are thought to be somewhat resistant. Having covered all of the primary sources of Pythium spp. of potting medium, used containers, plant propagules, or irrigation water, it is still essential to prevent the introduction of Pythium thereafter. Pythium spp. can be lurking around, just waiting for some lack of sanitation. So it is important to disinfect all bench surfaces, potting benches, tools, and equipments that will contact the potting mix. Management. Preventing Pythium root rot should be the primary management strategy. Eliminating the pathogen from the major sources of irrigation water, the growth medium, the containers, and even the cuttings can go a long way to managing this disease. Thereafter, application of preventative chemical fungicides provides some protection against invasion from a number of sources, including greenhouse workers. The chemicals used for Pythium control are the same as for Phytophthora control, and they generally target only those genera and some other fungus-like microorganisms, but not fungal pathogens. They are preventative chemicals, such as products containing etridiazole, mefenoxam, cyazofamid, fenamidone, and phosphonates. However, some species of Pythium are not sensitive to mefenoxam (Aegerter et al. 2002), so rotation or combinations with other chemicals should be practiced. In addition to chemical agents, there are some biological control agents that target Pythium spp., such as species of Trichoderma and Streptomyces. There are examples of suppressive soils that prevent or limit some root diseases, often created by incorporating organic materials, such as composts, that contain mixtures of microbes with great diversity. Within that microbial population are members with specific capacity to block the development of specific diseases. Some have the capacity to block sporangium development and therefore reduce zoospore release. Some are able to lyse the pathogen mycelium, while others produce other antibiotic compounds that inhibit growth of the pathogen in soil.

2.3

Cylindrocladium Blight and Wilt (Cylindrocladium scoparium Morgan)

Geographic occurrence and impact. A serious disease of evergreen, florist azaleas occurred in the USA and in European countries in the mid-1950s; within 10 years it appeared in major proportions in many parts of the USA and other countries. The cause of the disease was Cylindrocladium scoparium, a fungus previously known to attack primarily conifer and other tree species in seedling nurseries. Subsequently, the disease occurred on several other major nursery crops. The diseases varied in symptom expression, including root rot, stem lesions, leaf spots and flower blight, and wilt. In recent years to the present, serious Cylindrocladium diseases have occurred on azalea and mini rose, as well as other ericaceous plants, primarily in the early stages of cutting propagation. Subsequent sporulation of the pathogen under greenhouse conditions and spread by water splash or air movement of spores

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Fig. 3 Leaf spots on azalea plant caused by Cylindrocladium scoparium (left); close-up of spot on cutting during propagation (right) (R. G. Linderman)

results in devastating incidence and severity on many, often thousands, of plants. During that phase of sporulation and spread of the disease, more plants become infected, but remain symptomless, only to wilt and succumb later. Growers have sustained, and are still sustaining, severe economic losses from these diseases, often in the range of hundreds of thousands of dollars per year per grower. Symptoms/signs. There are multiple symptoms of Cylindrocladium infections of florist azaleas, depending on the production stage and environmental conditions (Linderman and Benson 2014). Leaf spot infections can occur if there is airborne inoculum in the greenhouse (Fig. 3). Often those spots go unnoticed because the infected leaf will abscise (due to the production of ethylene) and be lost on the pot surface among other leaves that had fallen for other reasons. Leaf spots are discrete spots that become necrotic as seen from the upper leaf surface. On the lower surface, however, mainly on red or pink cultivars, the veins radiating out from the spot are red. That symptom is absent or less pronounced on white cultivars. Flowers may also be infected by aerial inoculum, forming necrotic flecks that rapidly coalesce (Fig. 4). Infected fallen leaves become completely colonized by the pathogen in time as they remain moist. Internally, the pathogen forms microsclerotia in the leaf tissue that provide it long-term survival. Microsclerotia may also form in infected roots (Fig. 5). Under moist conditions, the sclerotia can generate abundant leaf-surface conidial sporulation that can contribute to the pathogen’s dispersal within the greenhouse (Fig. 6). Examination of such leaves, even with a hand lens, will reveal the diagnostic sporulation. Conidia released from fallen, infected leaves can wash into the medium and initiate root rot on newly formed roots. Infected roots are brown initially and then become necrotic. The extent of roots infected determines when the plant will succumb. Often, infected cuttings will die in pots where four cuttings were stuck

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Fig. 4 Flower infections on azalea caused by inoculation with Cylindrocladium scoparium (R. G. Linderman)

Fig. 5 Microsclerotia of Cylindrocladium scoparium formed in infected leaves (left and middle) or roots (right) (R. G. Linderman)

(Figs. 7 and 8). In many cases, however, the infections do not affect the appearance of the aboveground portions of the plant, mainly because new roots are forming fast enough to maintain the plant in spite of the infected roots. At a later stage, however, mortality may occur. Often, three or four rooted cuttings are transplanted into a single pot for growing on, and only one or two of them die (Fig. 9). That is a sign that the infection began at an earlier stage in propagation. Historically, however, when only one plant was transplanted to a larger pot, that plant eventually wilted due to the initial root infections on the cuttings. At that time, cutting into the lower stem would usually reveal an internal, brownish canker. That symptom alone, however, would not distinguish Cylindrocladium as the cause from cankers caused by Phytophthora,

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Fig. 6 Photo on the left shows sporulation of Cylindrocladium spp. on abscised azalea leaf. Photo in the center is a close-up of conidial clumps generated from microsclerotia imbedded in infected leaf. Photo on the right shows perithecia of Calonectria kyotensis (teleomorph) formed on infected azalea leaf (R. G. Linderman)

Fig. 7 Wilt symptoms on azalea cuttings (left) and older plant (right) where one of four cuttings was infected. Note distinctive defoliation from infection in photo on right (R. G. Linderman)

for example. Additional diagnostic tests need to be done to confirm Cylindrocladium blight. Biology and epidemiology. The main causal agent is Cylindrocladium scoparium Morgan, but other species with a perfect stage (teleomorph), such as Calonectria kyotensis and Calonecria theae, were also shown to be pathogenic (Alfieri et al. 1972). Those species with a Calonectria stage were shown to eject ascospores aerially when relative humidity declined (Linderman 1974a).

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Fig. 8 Root systems of azalea plants from one pot showing effects of root rot caused by Cylindrocladium (R. G. Linderman)

Fig. 9 Wilting of one azalea plant of four in the pot due to earlier infection by Cylindrocladium (left photo). Older plant wilting and death due to earlier root infection (right photo) (R. G. Linderman)

In nearly all cases, cutting and root rot of azaleas caused by species of Cylindrocladium begins during cutting propagation and spreads thereafter, depending on the greenhouse production system. It was demonstrated experimentally (Linderman 1973, 1974b) that the pathogen produced microsclerotia in infected leaves that dropped from plants with leaf spot infections. Those infections were often

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not noticed, and leaf abscission occurred as a result of ethylene production (Axelrood-McCarthy and Linderman 1981). The microsclerotia imbedded in the fallen leaves generated conidiophores on the surface, and conidia then splashed to other cuttings, causing root rot (Linderman 1974a). Some cutting roots became infected, but the plant remained symptomless. In the absence of stress under ideal growing conditions, liner plants moved through the production stages were sold and shipped only to die at some later time. Management. Aerially dispersed conidia that landed on plants from which cuttings would be taken can be contaminants, not yet causing infections. Therefore, chemical sanitization by dipping cuttings in solutions of fungicides or chlorine compounds, killing the contaminating conidia, is an effective management practice (Linderman, unpublished). Failure to sanitize cuttings has led to disastrous consequences from this disease. The source of airborne conidia in a greenhouse may be unknown, but infected plants with sporulation at the lower stem could be one source (Linderman, personal observation). Fallen, infected leaves on or below the production benches also can harbor the pathogen and become a source of dispersed conidia. Removal of such debris and flaming the areas under the benches and chemically sanitizing the benches is essential to removing sources of the pathogen inoculum. Assessment of the presence of Cylindrocladium spp. in soil or soilless media might also be needed, especially if the propagation medium is to be reused. Inserting azalea leaves into the medium and incubating for a week was shown to be effective in detecting the pathogen. The leaves were selective in being only infected by Cylindrocladium spp. (Linderman 1972). The method was also effective in isolating the pathogen from the stem by inserting a leaf into the cut end. If Cylindrocladium, or any other soilborne fungal pathogen, was detected, the medium must be sanitized with heat or a fumigant like Metam sodium prior to reuse (Linderman and Davis 2008b). Application of fungicides can also provide protection of plants exposed to aerial inoculum. Frequency of application is also critical due to the rate of sporulation of the pathogen as well as development of new, unprotected leaf surface from expansion. Fungicides containing thiophanate methyl have been shown to be effective.

2.4

Leaf and Flower Gall (Exobasidium spp.)

Geographic occurrence and impact. Leaf and flower gall disease of azaleas occurs widely throughout the world, mainly on landscape plants, but it can occur on florist azaleas under moist conditions. In the United States, the causal agent is generally considered to be Exobasidium vaccinii (Fuckel) Woronin. In China, Japan, and Korea, however, E. japonicum Shirai is considered to be the pathogen. Exobasidium rhododendri (Fuckel) C. Cramer has been reported on both azalea and rhododendron worldwide. The origin of the disease is not known, but likely first occurred on rhododendrons in their centers of origin. If enough galls appear on a plant, there can be some loss of vigor of the plant. Such severe infections rarely occur in commercial

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Fig. 10 Azalea leaf gall caused by Exobasidium vaccinii. Left: young galls; Middle: pathogen sporulation on gall; Right: older, necrotic galls (Courtesy of R. Rosetta)

greenhouse production, however. The impact of the disease comes from the time and cost of its management, involving labor and preventative chemical applications. Symptoms/signs. Leaf gall first appears as light green swellings on the edges and undersides of leaves. Both expanding leaves and flower buds and young, developing shoots are susceptible to infection. The infected tissue expands to form gall-like structures that generally involve the entire leaf (Fig. 10). As the galls mature, they turn white from surface sporulation of the fungal pathogen. The galls can be small or large and bladder shaped. The fungus causes the plant cells to divide and enlarge, making the tissue soft and succulent. Once galls turn white from sporulation, the pathogen can spread very rapidly by water splash or air movement to other plants, initiating new, secondary infections on leaves and even flowers. After sporulation occurs, the leaf galls slowly turn brown, shrink, and become quite hard. Biology and epidemiology. Spores of the pathogen apparently overwinter in bud scales. As buds break, the spores germinate and infect the young bud tissue as it expands. Humidity level in the greenhouse is a critical factor in the epidemiology of the leaf gall disease. High relative humidity and free water on the leaves and flowers is required for spore germination and initiation of new infections, so irrigation scheduling is very important. Overhead watering must be done early in the day so the foliage is not wet through the night. Because of the variation in relative humidity, often the lower leaves, where humidity is higher, are more susceptible than upper leaves where air circulation is better. Once infection has begun, the fungus grows in the intercellular spaces of the surface tissue, producing haustoria that penetrate cells to derive its nutrition. Eventually, often within a few days, the pathogen sporulates on the gall surface, causing it to turn white. Spores produced on a gall can be released and spread to other expanding buds, causing secondary infections. Some of the spores settle into protected sites in forming buds and are in position to initiate new infections. Management. The main strategy for managing azalea leaf gall is to diligently scout production plants for the first signs of gall development. Small galls must be removed before they turn white with sporulation. Prevention of galls also generally involves application of fungicides as buds break and new foliage appears. Plants should be sprayed before and after removing galls with chemicals such as mancozeb-

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Fig. 11 Hyphal webbing site of azalea cuttings in propagation caused by Rhizoctonia solani. Note discrete spots on leaf in lower right (R. G. Linderman)

containing fungicides, copper-containing materials, quaternary ammonium materials, and even oxidizing agents such as hydrogen peroxide. A good sticker may be needed to help retain the applied chemical. Application of any of these materials is strictly preventative, not curative.

2.5

Rhizoctonia Root Rot and Web Blight (Rhizoctonia spp.)

Geographic occurrence and impact. Rhizoctonia root rot and subsequent blighted foliage occurs occasionally on florist azaleas in the United States (Copes and Benson 2014). The disease also has been reported in Australia. This disease can occur on many different woody ornamental plants, including rhododendrons. Most often, the disease occurs during cutting propagation where moist, warm conditions prevail. Economic losses result from the loss of affected rooted cuttings, but some infections aren’t expressed until later when canopy growth is affected and some dieback/blight occurs. Symptoms/signs. When Rhizoctonia root rot occurs during propagation of cuttings, the affected cuttings fail to root or the new roots become infected, causing the entire cutting to collapse (Fig. 11). Often the infection site will expand and develop, spreading to adjacent cuttings. Webbing of aerial hyphae occurs, looking like a spider web moving from plant to plant. The pathogen moves both in the rooting medium and from leaf to leaf above the medium surface. Biology and epidemiology. Some propagators root cuttings under a reemay cover in order to retain heat and to reduce the frequency of misting from above. The high humidity under the reemay allows the fungus to move from one infected cutting to adjacent cuttings, often leading to a patch of infected cuttings. The occurrence of a web-blight pattern where the fungus becomes aerial and infects

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the lower leaves results in a distinct leaf spot at the infection site. The primary causal agent, Rhizoctonia solani J. G. Kuhn, is a nonsporulating Deuteromycete, so how the pathogen invades the propagation system is unknown. The possible sources could be the medium, reused rooting flats, or some cuttings becoming contaminated during cutting collection. The incidence of this disease is very infrequent and spread is by vegetative means. Spread may also occur in particulate material moved by air currents. Management. The infrequent occurrence of Rhizoctonia root rot and cutting death supports the idea that no active management is required, other than removal of affected cuttings at an infection site, including adjacent cuttings that usually show no signs of infection. Often that means removal of the entire affected plug flat. Addressing the possible sources of the pathogen, however, is in order. If propagation flats are being reused, then their sanitization with aerated steam (Linderman and Davis 2008b) is required to kill any carryover of the pathogen from a previous crop, even crops other than azalea. If the rooting medium is suspected as the source, then a decision would be needed as to steam treatment that is generally not a course of action unless other diseases have occurred that would also be curbed by medium pasteurization. Since R. solani can colonize organic components of the medium as a saprophyte, rigid sanitation in the propagation area is called for to prevent that from happening. Affected areas of the propagation area should also be treated chemically to avoid spread and carryover. Chemicals that specifically target Rhizoctonia pathogens should be used; broader spectrum chemicals might be used if other causal agents that can cause similar symptoms are known to occur, such as species of Pythium or Phytophthora. Since sporulation is not involved in disease initiation or spread, cuttings are less likely to be contaminated, so their sanitation generally is not necessary.

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Genetic Abnormalities

3.1

Witches’ Broom

Geographic occurrence and impact. Some cultivars of florist azaleas have a tendency to develop a cluster of small leaved branches in a tight configuration characterized as a witches’ broom. The occurrence of these structures has little impact on the growth of the plant, and plants that exhibit them are generally discarded as being offtype or the affected branch simply pruned off (Linderman 2014). Symptoms/signs. The witches’ broom branches occur generally on one location on the plant, often near the base of the plant. The tight branching and greatly reduced leaf size distinguish it from the rest of the plant (Fig. 12). Biology and epidemiology. Evergreen azaleas and rhododendrons occasionally exhibit abnormal, miniaturized vegetative growth in the form of a witch’s broom. Researchers generally agree that the condition is the result of a genetic change that

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Fig. 12 Witches’ broom on azalea (R. G. Linderman)

occurred, and that no biotic causal agent is involved. Therefore, there is no risk of transmission from affected plants to adjacent plants. Management. Since no biotic agent is involved in the development of witches’ broom, no management is required other than to either prune off the broom, or simply discard the plant as being off-type.

References Aegerter BJ, Greathead AS, Pierce LE, Davis RM (2002) Mefenoxam – resistant isolates of Pythium irregulare in an ornamental greenhouse in California. Plant Dis 86(6):692 Alfieri SA Jr, Linderman RG, Morrison RH, Sobers EK (1972) Comparative pathogenicity of Calonectria theae and Cylindrocladium scoparium to leaves and roots of azalea. Phytopathology 62:647–650 Axelrood-McCarthy PE, Linderman RG (1981) Ethylene production by cultures of Cylindrocladium floridanum and C. scoparium. Phytopathology 71:825–830 Copes WE, Benson DM (2014) Rhizoctonia damping-off and root rot; Rhizoctonia web blight. In: Linderman RG, Benson DM (eds) Compendium of rhododendron and azalea diseases and pests, 2nd edn. APS Press, St. Paul, pp 14–19 Ivors KL (2014) Pythium damping-off and root rot. In: Linderman RG, Benson DM (eds) Compendium of rhododendron and azalea diseases and pests, 2nd edn. APS Press, St. Paul, pp 13–14 Jones RK, Benson DM (eds) (2001) Diseases of woody ornamentals and trees in nurseries. APS Press, St. Paul Linderman RG (1972) Isolation of Cylindrocladium from soil or infected azalea stems with azalea leaf traps. Phytopathology 62:736–739 Linderman RG (1973) Formation of microsclerotia of Cylindrocladium spp. in infected azalea leaves, flowers, and roots. Phytopathology 63:187–191 Linderman RG (1974a) Ascospore discharge from perithecia of Calonectria thea, C. crotalariae, and C. kyotensis. Phytopathology 64:567–569 Linderman RG (1974b) The role of abscised Cylindrocladium-infected azalea leaves in the epidemiology of Cylindrocladium wilt of azalea. Phytopathology 64:481–485 Linderman RG (2014) Witches’ broom. In: Linderman RG, Benson DM (eds) Compendium of rhododendron and azalea diseases and pests, 2nd edn. APS Press, St. Paul, p 68

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Linderman RG, Benson DM (eds) (2014) Compendium of rhododendron and azalea diseases and pests, 2nd edn. APS Press, St. Paul Linderman RG, Davis EA (2008a) Evaluation of chemical agents for the control of Phytophthora ramorum and other species of Phytophthora on nursery crops [Online]. Plant Health Prog. doi:10.1094/PHP-2008-0211-01-RS Linderman RG, Davis EA (2008b) Eradication of Phytophthora ramorum and other pathogens from potting medium or soil by treatment with aerated steam or fumigation with metam sodium. HortTechnology 18(1):106–110 Linderman RG, Moore LW, Baker KF, Cooksey DA (1983) Strategies for detecting and characterizing systems for biological control of soilborne plant pathogens. Plant Dis 67:1058–1064 Weiland JE, Beck BR, Davis A (2013) Pathogenicity and virulence of Pythium species obtained from forest nursery soils on Douglas-fir seedlings. Plant Dis 97:744–748

Diseases of Begonia Cristina Rosa and Gary W. Moorman

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-Like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Fusarium Wilt (Fusarium foetens) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Gray Mold (Botrytis Blight; Botrytis cinerea Pers.: Fr.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Powdery Mildew (Oidium begoniae, Asexual Stage) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Pythium Root Rot (Pythium sp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Bacterial and Phytoplasma Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Bacterial Spot (Bacterial Blight; Xanthomonas campestris pv. begoniae (Takimoto) Dye) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Viral Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Clover Yellow Mosaic Potexvirus (ClYMV) Described by Johnson (1942) . . . . . . . . . 4.2 Cucumber Mosaic Virus (CMV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Impatiens Necrotic Spot Virus (INSV) and Tomato Spotted Wilt Virus (TSWV) . . . . 4.4 Tobacco Mosaic Virus (TMV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5 Tobacco Necrosis Necrovirus (TNV) and Carnation Mottle Carmovirus (CarMV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6 Tobacco Ringspot Virus (TRSV) and Arabis Mosaic Nepovirus (ArMV) . . . . . . . . . . . . 4.7 Broad Bean Wilt Virus (BBWV) Described by Stubbs (1947) . . . . . . . . . . . . . . . . . . . . . . . 4.8 Zucchini Yellow Mosaic Virus (ZYMV) Described by Lisa et al. (1981) . . . . . . . . . . . . 5 Nematode Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 Foliar Nematode (Aphelenchoides fragariae) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 Abiotic Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2 2 2 3 4 5 7 8 9 10 10 10 11 12 12 13 13 14 14 15 15

C. Rosa (*) • G.W. Moorman (*) Department of Plant Pathology & Environmental Microbiology, The Pennsylvania State University, University Park, PA, USA e-mail: [email protected]; [email protected] # Springer International Publishing Switzerland 2016 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florist's Crops Diseases, Handbook of Plant Disease Management, DOI 10.1007/978-3-319-32374-9_30-1

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Abstract

Begonias are susceptible to a wide variety of fungi, bacteria, and viruses, as well as nematodes and abiotic diseases. The systemic nature of some of the pathogens makes it likely that the diseases they cause can be found wherever vegetatively propagated begonias are shipped. Management of these pathogens is paramount for specialty propagators while growers purchasing plants must inspect incoming plants for symptoms and understand the biology of the pathogens involved in order to manage them effectively. Keywords

Fusarium wilt • Botrytis blight • Powdery mildew • Pythium root rot • Bacterial spot • Viruses • Foliar nematodes • Abiotic diseases

1

Introduction

Begonias are a very diverse and complex group of plants. The name “Begonia” was first used by Charles Plumier, a Franciscan monk and botanist, in 1703. Linnaeus described all the species known to him in 1753, and by the end of the 1700s, begonias were being cultivated widely in Europe. By the early 1800s, hybrids were being developed. Based on their dominant horticulture characteristics, begonias can be grouped in categories including cane-like, shrub-like, semperflorens, rhizotamous, rex cultorum, tuberous, and trailing-scandent (Thompson and Thompson 1981).

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Fungal and Fungus-Like Diseases

2.1

Fusarium Wilt (Fusarium foetens)

Geographic occurrence and impact. This disease has been found in Europe, Japan, and North America causing wilt and sometimes stem rot (Elmer and Vossbrinck 2004; Schroers et al. 2004; Sekine et al. 2008; Tian et al. 2012; Tschope et al. 2007; Van der Gaag and Raak 2010). Symptoms/signs. Leaves become dull green and the leaf veins may yellow. The leaves wilt and die. The vascular tissue of infected plants turns brown. A basal stem rot may develop as the disease process proceeds (Fig. 1). Large numbers of pale orange spores develop on the dying tissue, particularly if the tissue is placed on a wet surface in a container and incubated. Biology and epidemiology. Fusarium foetens is related to the Fusarium oxysporum complex that contains many form species responsible for vascular wilts of various plants. It is thought that the pathogen was introduced to Europe on plant material brought there for breeding (Schroers et al. 2004), indicating that the pathogen may be spread long distances associated with plant shipments.

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Fig. 1 Fusarium wilt on begonia (Photo courtesy of Wade Elmer)

Management. Infected plants should be destroyed. The various types and cultivars of begonia differ in relative susceptibility (Brand and Wienberg 2005; Tian et al. 2012; Tian and Zheng 2012). The biological control agents Bacillus subtilis, Streptomyces griseoviridis, Gliocladium catenulatum, Streptomyces lydicus, and Trichoderma harzianum have been reported to provide protection of begonias against Fusarium foetens in greenhouse experiments (Tian and Zheng 2013). To reduce the presence of the fungus in production systems, chlorine bleach, hydrogen peroxide, and quaternary ammonia sanitizers have been found effective (Elmer 2008).

2.2

Gray Mold (Botrytis Blight; Botrytis cinerea Pers.: Fr.)

Geographic occurrence and impact. This disease is very common and occurs wherever plants are grown if the environmental conditions are conducive to its development. The pathogen, Botrytis cinerea, can infect almost any plant in a greenhouse including begonias. Symptoms/signs. Cuttings rot at their base. Tan spots develop on leaves. Established plants rot at the crown or a dry shriveled area develops along a branch (Tompkins 1950). Abundant, dusty, gray masses of spores form on infected tissue if the humidity is high or if infected tissue is placed in a container with moisture. Biology and epidemiology. Seedlings and all above ground parts of mature plants are susceptible. The fungus can infect intact tissue or through wounds. Infection occurs most readily when the fungus has a food base from which it attacks, such as fading flowers or senescent leaves that have fallen onto healthy leaves. High relative humidity, wetness on the plant, and temperatures between 18  C and 25  C greatly favor Botrytis infection (Jarvis 1980). The spores are readily spread by air currents (Hausbeck and Pennypacker 1991).

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Fig. 2 Powdery mildew on begonia (Photo courtesy of The Pennsylvania State University Dept. of Plant Pathology and Environmental Microbiology)

Management. Maintain low humidity within the crop canopy by spacing plants well and venting in a greenhouse setting to improve air circulation. Remove dead and dying flowers and leaves from the area around the plants, from the potting soil surface, and from the plant. Remove this debris from the greenhouse promptly or put it in a closed waste container. Avoid unnecessarily damaging plants. Apply a fungicide preventively but note that the exclusive use of one class of chemical, particularly systemic chemicals, can result in the selection of fungicide resistance to the chemical class. Resistance to benzimidazole fungicides is very widespread in Botrytis populations Do not rely on only one chemical class (Elad et al. 1992; Gullino and Garibaldi 1987; Hausbeck and Moorman 1996; Moorman and Lease 1992).

2.3

Powdery Mildew (Oidium begoniae, Asexual Stage)

Geographic occurrence and impact. This disease is widespread, probably found wherever begonias are grown, and it significantly damages the aesthetic quality of affected plants. The obligate plant pathogenic organism exhibits dimorphism. That is, the asexual phase of growth looks significantly different from the sexual stage (Bélanger et al. 2002). The asexually reproducing phase of the pathogen is named Oidium begoniae (Fig. 2). The sexual stage is thought to be a member of the genus Microsphaera because the conidia of the asexual stage are not formed in chains (Quinn and Powell 1981), but the structures formed as a result of sexual reproduction (chasmothecia; formerly cleistothecium) are rarely found associated with begonia. It is known that Sphaerotheca fuliginea (Powell 1985) and Golovinomyces cichoracearum (formerly Erysiphe cichoracearum) (Sammons et al. 1982) may also cause powdery mildew in begonia. Symptoms/signs. White, mealy fungal growth develops on leaves, flowers, and stems. Tissue beneath the fungus may die.

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Biology and epidemiology. Disease develops most rapidly when temperatures are 20–21  C and is greatly inhibited if temperatures are above 28  C (Quinn and Powell 1982). Unlike essentially all other fungal plant pathogens, wetness has little influence on disease development of this powdery mildew. In fact, it flourishes when leaf surfaces are relatively dry. Severe powdery mildew outbreaks should be anticipated when cool, damp night conditions alternate with warm, dry days. Management. Examine plants carefully and frequently to detect the onset of disease. Maintain a fungicide program to protect plants. It is known that horticultural oil can kill powdery mildew fungi when it comes in direct contact with the pathogens. Once the oil dries on the plant surface, it has no activity against powdery mildew. That is, it has no residual activity. Applied under the wrong conditions or when the plant is wilting, oils are phytotoxic.

2.4

Pythium Root Rot (Pythium sp.)

Geographic occurrence and impact. Several species of Pythium have been reported to cause root rot and lower stem rot of begonia around the world including P. ultimum (Globisporangium ultimum) and P. splendens (Globisporangium splendens) (Farr et al. 1989). Phytopythium helicoides (formerly, Pythium helicoides) has also been found on begonias (Yang et al. 2013). Symptoms/signs. Infected seedlings die. Shiny tan, water-soaked areas develop on the stems and petioles of established plants at or just above the soil line as plants collapse and die (Middleton 1942). Under very high humidity conditions, the pathogen may be seen as a fluffy, white mass near the soil line but this is unusual to see. Biology and epidemiology. Pythium species are often found in field soil, sand taken from streams and rivers, and can be found in pond and lake sediments, and dead roots of previous crops (Ivors and Moorman 2014). Excessively wet potting mixes greatly favor the development of root rot. Management. Plant in pasteurized, not sterilized, potting media. If the potting mix has been sterilized (killing all living organisms in it) by heating it to too high a temperature or heating it too long, a biological vacuum will have been created. If Pythium then contaminates the sterile potting mix, it can cause very severe crop losses, because it has no competition and no natural inhibition by other microbes. Keep hose ends off the ground in order to avoid picking up Pythium-contaminated soil and then spraying it onto the crop. Use a well-drained potting mix and do not over-water the plants, particularly if plants are not utilizing a great deal of water because of prevailing weather conditions. Apply a fungicide or biological control agent at planting for the best protection. If the crop is grown for several months, any chemical or biological control agent will have to be applied repeatedly. Listed below are additional fungus or fungus-like organisms which have been associated with Begonia (Farr et al. 1989). For the most up-to-date listing, search http://nt.ars-grin.gov/fungaldatabases/fungushost/fungushost.cfm:

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Aecidium begoniae Agrobacterium tumefaciens Alternaria sp. Alternaria tenuis (Alternaria alternata) Armillaria mellea Bartalinia begoniae Botryosphaeria sp. Botryotinia fuckeliana (Botrytis cinerea) Ceratobasidium sp. Cercospora begoniae Cercospora sigesbeckiae Choanephora cucurbitarum Cladosporium inconspicuum Cladosporium sphaerospermum Coleosporium begoniae Colletotrichum capsici (Colletotrichum truncatum) Colletotrichum gloeosporioides Colletotrichum sp. Corticium solani (Rhizoctonia solani) Corynespora cassiicola Curvularia inaequalis Erysiphe begoniae Erysiphe begoniicola Erysiphe communis (Erysiphe pisi var. pisi) Erysiphe orontii (Golovinomyces orontii) Erysiphe polygoni Erysiphe polyphaga (Golovinomyces orontii) Fusarium begoniae Fusarium equiseti Fusarium roseum Fusarium solani Gloeosporium begoniae Glomerella cingulata (Colletotrichum gloeosporioides) Golovinomyces orontii Helminthosporium sp. Lasiodiplodia theobromae Macrophoma sp. Macrophomina phaseoli (Macrophomina phaseolina) Marssonina mali Meliola begoniae Microsphaera begoniae (Erysiphe begoniicola) Moniliopsis aderholdii (Rhizoctonia solani) Mycena citricolor Myrothecium roridum Oidium begoniae (Golovinomyces orontii)

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Oidium ellipticum Penicillium bacillisporum (Talaromyces bacillisporus) Pestalotiopsis sp. Phoma sorghina (Epicoccum sorghi) Phoma sp. Phomopsis sp. Phyllachora begoniae Phyllosticta begoniae Phyllosticta sp. Phyllostictina sp. Phytophthora cactorum Phytophthora cryptogea Phytophthora cryptogea f. sp. begoniae Phytophthora nicotianae var. nicotianae (Phytophthora nicotianae) Phytophthora niederhauserii Phytophthora parasitica (Phytophthora nicotianae) Pseudoidium sp. Pucciniastrum boehmeriae Pythium aphanidermatum Pythium debaryanum (Globisporangium debaryanum) Pythium intermedium (Globisporangium intermedium) Pythium irregulare (Globisporangium irregulare) Pythium vexans (Phytopythium vexans) Rhizoctonia solani Sclerotinia sclerotiorum Sclerotium rolfsii (Athelia rolfsii) Septonema sp. Septoria begoniae Shrungabeeja begoniae Sphaeropsis begoniicola Stemphylium sp. Thanatephorus cucumeris (Rhizoctonia solani) Thielaviopsis basicola Verticillium albo-atrum Verticillium dahliae

3

Bacterial and Phytoplasma Diseases

Where possible, the scientific names used are those accepted by the International Society of Plant Pathology Committee on the Taxonomy of Plant Pathogenic Bacteria (Bull et al. 2010).

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Fig. 3 Bacterial blight on begonia (Photo courtesy of The Pennsylvania State University Dept. of Plant Pathology and Environmental Microbiology)

3.1

Bacterial Spot (Bacterial Blight; Xanthomonas campestris pv. begoniae (Takimoto) Dye)

Geographic occurrence and impact. This disease, one of the most damaging to begonias, was first described in the 1920s (Powell 1985) (Fig. 3). It causes a leaf spot or blighting of some cultivars but can be systemic in Rieger elatior begonias (Harri et al. 1977). In some cases such as a slight infection, the disease is easily overlooked. Thus, the pathogen can be inadvertently shipped with plants if infection goes undetected. Symptoms/signs. Water-soaked circular lesions surrounded by yellow halos develop on leaves. In wax begonias (semperflorens types), the spots may be very small (1 mm diameter) (Daughtrey et al. 1995). Plants slowly die one leaf at a time. If infection begins at the leaf margin, wedge-shaped brown dead areas develop with the wide part of the wedge at the margin and the point of the wedge pointing toward the plant stem. When infected tissue is cut, placed in a clear drop of water on a microscope slide, and observed with a bright field microscope with the iris diaphragm mostly closed, bacteria can be seen to stream from the tissue. When Rieger begonias are infected systemically, they wilt and die. Biology and epidemiology. The bacteria can enter injured roots or leaves and become systemic in some cultivars of Rieger begonias. Eventually wilting and death of the plant occurs. In some other begonias such as Rex, the pathogen remains localized in leaf spots (Jodon and Nichols 1974). The pathogen can survive for extended periods of time if the leaf debris is dry. The pathogen can be spread from plant to plant on workers’ hands, clothing, and tools and has been shown to be disseminated in recycled irrigation water (Atmatjidou et al. 1991). Management. Purchase plants free of the pathogen. Discard infected plants, particularly Rieger-types which are systemically infected. Remove infected leaves from Rex and tuberous types because they are not systemically infected. Irrigate plants in a manner that keeps water off the foliage. Do not propagate from infected plants. Remove the debris from infected plants from the greenhouse or place it in a closed

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container. Workers should wash their hands thoroughly periodically when performing tasks on the plants and should disinfect any tools that come in contact with the plants. Additional Diseases with Bacterial Pathogens Crown gall (Rhizobium tumefaciens, formerly Agrobacterium tumefaciens) (Powell 1985) Bacterial fasciation (Rhodococcus fascians) (Powell 1985) Soft rot (Pectobacterium carotovorum, formerly, Erwinia carotovora) (Powell 1985) Additional Diseases with Phytoplasma Pathogens Phytoplasmas induce symptoms such as phyllody, shoot proliferation, small leaves and flowers, and stunted plant growth. These bacteria are vectored primarily by leafhoppers, planthoppers, and psyllids. “Candidatus Phytoplasma asteris” A phytoplasma in the 16SrI group was reported to infect begonia (Powell 1985). Shoot proliferation. A phytoplasma belonging to the 16SrIII group was associated with shoot proliferation in begonia in Brazil in 2006 (Ribeiro et al. 2006).

4

Viral Diseases

Viruses of begonia are associated with mild to severe mosaic and mottling, ringspots, leaf malformation, stem necrosis, chlorosis, and stunting. Generally viruses can be controlled by using clean propagative material, by discarding diseased plants and isolating surrounding plants and controlling vector populations for viruses that are vector transmitted. Clean potting soil needs to be used to control viruses whose vectors are soil inhabitants. The presence of viruses can be diagnosed by use of molecular tests such as reverse transcriptase PCR or immune-based assays such as ELISA or rapid immunostrips, or by indicator hosts. The EPPO Panel on “Certification of Ornamentals” developed procedures for the production of healthy carnation, pelargonium, lily, narcissus, chrysanthemum, tulip, crocus, iris, begonia, impatiens, rose, freesia, hyacinth, kalanchoe, and petunia. In the USA, certification programs are managed by the major commercial propagators. Pelargonium and other major flower crops are tested for the presence of the most prevalent viruses either by in house plant pathologists or by testing services contracted with by the propagators. In the USA, phytosanitary certificates are required for intra- and interstate plant movement. Data on pathogens of high consequence, including viruses, are recorded by the National Plant Diagnostic Network. Where available, information was taken from Adams and Antoniw (2006).

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4.1

C. Rosa and G.W. Moorman

Clover Yellow Mosaic Potexvirus (ClYMV) Described by Johnson (1942)

Geographic occurrence and impact. It is common in the Western USA and Canada (Agrawal et al. 1962; Pratt 1961). It has occasionally being imported to the UK, but it has not established in the EPPO region. Symptoms. ClYMV induces color variations and poor vigor. Biology and Epidemiology. ClYMV is in the family Alphaflexiviridae. ClYMV is transmitted by infected sap but no vector is known. Reports indicate that it is also seed transmitted (Hampton 1963). Clover yellow mosaic virus is serologically distantly related to White Clover mosaic virus. Management. Use of healthy propagative material and sanitation is recommended.

4.2

Cucumber Mosaic Virus (CMV)

Geographic occurrence and impact. CMV is found worldwide, has an extremely broad host range, and can infect species in more than 100 plant families (Zitter and Murphy 2009). Symptoms/signs. Symptoms observed on infected begonia are mosaic, leaf deformation and curling, and vein clearing. Biology and epidemiology. CMV (Jacquemond 2012; Palukaitis and GarcíaArenal 2003) belongs to the family Bromoviridae. CMV is a ss + RNA virus with a tripartite genome, is aphid transmitted in a noncirculative (nonpersistent) manner (Hoggan 1933; Simons 1955; Watson and Roberts 1939), and can be seed transmitted in some plant species (Neergaard 1977). Management. Control for this virus is particularly difficult, since CMV is a common virus on many plant Families and it can be transmitted by multiple aphid species. Because aphids transmit CMV readily during probing, aphid control is partially efficacious at reducing the spread of the virus (Zitter and Murphy 2009). Mineral oil has been proposed to delay viral symptoms in different crops (Simons and Zitter 1980).

4.3

Impatiens Necrotic Spot Virus (INSV) and Tomato Spotted Wilt Virus (TSWV)

Geographic occurrence and impact. INSV and TSWV are prevalent in many plant families and species, making their control particularly challenging. INSV can infect around 800 plant species and TSWV more than 1,000. Symptoms/signs. Symptoms associated with INSV and TSWV are mosaic, mottling, stem necrosis, ringspots on leaves, and leaf deformation (Fig. 4). Biology and epidemiology. INSV (Law and Moyer 1990) and TSWV (Samuel et al. 1930) are in the Family Bunyaviridae. These ssRNA viruses have tripartite

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Fig. 4 Impatiens necrotic spot virus on begonia (Photo courtesy of The Pennsylvania State University Dept. of Plant Pathology and Environmental Microbiology)

genomes, with negative or ambisense orientation. INSV and TSWV are transmitted by thrips in a circulative and propagative manner (Ullman et al. 1993; Wijkamp et al. 1993); thus, they can replicate in their plants as well as in their insect hosts. Management. One of the best ways to manage these viruses is to control thrips numbers, since thrips need a relatively long acquisition and transmission time, and since the viruses encounter a latent period in the vector prior to becoming transmissible. For instance, in the Netherlands, where INSV and TSWV have been reported on begonia, the incidence of disease has decreased significantly thanks to the implementation of a rigorous thrips management program. The Netherlands has also established a certification program for many ornamental plants, including begonia. To complicate matters, many thrips are insecticide resistant (Brødsgaard 1994; Zhao et al. 1995); thus, care needs to be taken to rotate different classes of insecticides. Weed management is also necessary (Bond et al. 1983; Cho et al. 1986; Kobatake et al. 1984), since weeds can serve as reservoirs for tospoviruses and thrips.

4.4

Tobacco Mosaic Virus (TMV)

Geographic occurrence and impact. TMV can infect a variety of plants in 30 Families worldwide (Shew and Lucas 1991). Symptoms/signs. Symptoms on begonia are reported to be yellowing, necrosis, mosaic, leaf distortion, and plant stunting. Biology and epidemiology. TMV belong to the Virgaviridae family. The virus has a ss + RNA genome. Symptoms attributed to TMV are often caused by other Tobamoviruses, and many Tobamovirus species were once classified as strains of TMV. TMV is occasionally transmitted by chewing insects, but most commonly it is mechanically spread (Harris and Bradley 1973; Lojek and Orlob 1969), in fact TMV virions are extremely stable. TMV can persist in the soil, probably on plant debris, and can infect roots. It can penetrate wounded embryos from infected seed coats (Broadbent 1965). TMV is a special concern in greenhouses, where it can be very hard to eradicate (Broadbent and Fletcher 1963).

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Management. Sanitation is the best TMV control method.

4.5

Tobacco Necrosis Necrovirus (TNV) and Carnation Mottle Carmovirus (CarMV)

Geographic occurrence and impact. TNV (Price 1940) can experimentally infect 88 species in 37 plant families and CarMV can infect members in more than 9 families of plants worldwide. Symptoms. On most hosts TNV causes necrotic lesions and rarely systemic symptoms. TNV can support coinfection with satellite viruses. Carnation mottle virus can infect Begonia elatior and Begonia x cheimantha where it is associated with symptoms of vein clearing, leaf curling, and flower breaking. Biology and epidemiology. These viruses are in the Family Tombusviridae. Members of this family have a ss + RNA monopartite genome. TNV (described first by Smith and Bald 1935 and more recently by Fraenkel-Conrat 1988) is transmitted by the zoospores of the fungus-like microorganism Olpidium brassicae (Kassanis and MacFarlane 1964; Teakle 1962; Teakle and Gold 1963), is not transmitted by seed or by pollen, and can be transmitted mechanically. CarMV is not known to be transmitted by a vector or by seed, but it is probably transmitted by plant to plant contact and mechanically during cultural practices. The virus is also stable in water and can be transmitted through irrigation. An unconfirmed report from India indicates that aphids could be vectors of this virus. Management. To control TNV, care should be taken in using clean irrigation water and potting soil, and in avoiding diseased propagative material. Control of CarMV consists of using certified virus-free material, strict sanitation, and care in handling infected plants.

4.6

Tobacco Ringspot Virus (TRSV) and Arabis Mosaic Nepovirus (ArMV)

Geographic occurrence and impact. ArMV (Smith and Markham 1944) has been detected in 13 States in the USA where it has been the object of quarantine, but it is not widespread or established outside Europe. ArMV has a wide host range. TRSV (Fromme et al. 1927) geographic distribution is mainly in northern USA and China, but it has also been found in Europe and Australia. More than 17 plant families are susceptible to TRSV (Price 1940). Symptoms/signs. Begonia yellow spot, caused by TRSV, shows symptoms of chlorotic local lesions and yellow mottle patterns, sometimes resembling natural variegation. Biology and epidemiology. ArMV and TRSV belong to the family Secoviridae. Their genomes consist of two linear ss + RNA segments. ARMV is nematode and seed transmitted and can be transmitted by Cuscuta sp. Nematodes lose the virus during molting and do not transmit it to their progeny. TRSV is transmitted by

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nematodes (McGuire 1964), pollen, and seeds as well as by insects (Dunleavy 1957; Komuro and Iwaki 1968; Messieha 1969; Schuster 1963) and mites (Thomas 1969) in a nonspecific manner. TRSV does not replicate in its nematode vector, is lost during molting, and is not transmitted to the nematode progeny. Management. TRSV is controlled by using clean potting soil, discarding infected propagative material, and by using clean seed.

4.7

Broad Bean Wilt Virus (BBWV) Described by Stubbs (1947)

Geographic occurrence and impact. The virus has been reported worldwide (Lisa and Boccardo 1996). The virus was reported in the USA in New York state, South Carolina and Minnesota in the early 1980s. BBWV was subdivided in two species, BBWV-1 and BBWV-2, in 2000. The two species are serologically distinct, with BBWV-1 more prevalent in Europe, while BBWV-2 is more prevalent in North America, Asia, and Australia. BBWV has a broad host range (Edwardson and Christie 1991). Symptoms/signs. Symptoms associated with BBWV on begonia are leaf mottling, faint ringspots, color-break in flowers, and generalized stunting. Symptoms tend to be more severe in winter. Biology and epidemiology. As ArMV and TRSV, BBWV is in the family Secoviridae, and its genome consists of two ss + RNA segments. In contrast to ArMV and TRSV, BBWV is aphid transmitted in a noncirculative (nonpersistent) manner (Lisa and Boccardo 1996). Management. Control of BBWV is difficult because its host range is broad and it is transmitted in a nonpersistent manner; thus, exclusion of aphids is advised, as well as of weeds that serve as virus reservoirs.

4.8

Zucchini Yellow Mosaic Virus (ZYMV) Described by Lisa et al. (1981)

Geographic occurrence and impact. The virus was detected on begonia in Taipei in 2008. As for CMV, another nonpersistently aphid transmitted virus, several plant families are susceptible to ZYMV and the virus is widely distributed, making its control difficult. Symptoms. Symptoms in begonia consist of faint ringspots on leaves at the early stage of infection; the spots become chlorotic and coalesce during disease progression. Biology and Epidemiology. ZYMV is a potyvirus in the family Potyviridae. Its genome consists of a ss + RNA filament and is monopartite. The virus is aphid transmitted in a noncirculative (nonpersistent) manner. Management. While the use of insecticides is not recommended to control nonpersistent viruses and not many experiments have been reported on control of ZYMV in begonia, strategies used to control this virus on other crops would be

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probably appropriate. These strategies are: the use of resistant material, the use of clean propagative material and adoption of good hygiene practices, the use of mineral oil, and the exclusion of aphids from greenhouses.

5

Nematode Diseases

5.1

Foliar Nematode (Aphelenchoides fragariae)

Geographic occurrence and impact. Aphelenchoides affects a large number of ornamental plants, including begonia. In some cases, the nematode invades the vascular tissue and the begonia remains mostly symptomless. Thus, the nematode can be inadvertently shipped from place to place in infected plants. Severe damage can occur in begonias. Symptoms/signs. Plants may be stunted. In some cases, excessive red pigmentation develops in infected leaves. Bronzed or water-soaked areas develop on leaves of some cultivars. Fibrous-rooted cultivars have small brown leaf spots. Some cultivars exhibit no symptoms despite heavy infection. If leaf or stem tissue is placed in a drop of water in a clear dish and teased into small pieces, numerous colorless nematodes can be seen with some magnification, vigorously moving in a snake-like motion in the water. The nematodes will tend to settle to the bottom of the dish. Biology and epidemiology. This nematode can enter stomata or wounds, continue to move in the plant, and feed on cells inside the plant (migratory endoparasitic life style). Or the nematode can remain on the surface of the plant, continue to move at will if there is water on the plant tissue, and feed on surface cells (migratory ectoparasitic life style). There is some suggestion that the nematode may have the ability to enter intact leaf surfaces directly (Riedel 1985). Peirson (1974) found that the “Aphrodite Rose” cultivar of Rieger begonia can harbor 12,000 larvae of this nematode per gram of fresh weight of leaves, yet exhibit no symptoms. Management. Purchase nematode-free plants. Irrigate plants in a manner that keeps water off the foliage so that movement of the nematodes on the leaf surface is inhibited and localized dispersal among closely spaced plants by splashing is avoided. Discard infected plants. A very high value stock plant can be heat-treated to eliminate nematodes (Guba and Gilgut 1938). Some experimentation is required to determine the best temperature and exposure duration for eliminating the nematodes while not killing the plant. An effective hot water dip may range from 1 min at 49  C (120  F) to 5 min at 45  C (115  F), depending upon the cultivar. A treated plant may be severely damaged, but it will recover over time and be free of the nematode. Additional Diseases with Nematode Pathogens Root-knot (Meloidogyne spp.) (Powell 1985)

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15

Abiotic Diseases

A variety of leaf symptoms develop as a result of nutrient deficiencies in begonia. If yellowing is the predominant symptom and the yellowing is between the veins, iron or magnesium may be lacking. Uniformly yellowing leaves indicates nitrogen or calcium deficiency. Dead tissue at the leaf margin may indicate potassium deficiency, while stunting of an otherwise green plant could be due to lack of phosphorus. A lack of boron causes the foliage to russet and cracks may develop in the leaf petioles and the plants may be very brittle (Nelson et al. 1977).

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Fraenkel-Conrat H (1988) Tobacco necrosis, satellite tobacco necrosis, and related viruses. In: The plant viruses. New York, Springer US, pp 147–161 Fromme FD, Wingard SA, Priode CN (1927) Ringspot of tobacco: an infectious disease of unknown cause. Phytopathology 17:321–328 Guba EF, Gilgut CJ (1938) Control of begonia leaf-blight nematode. Mass Agric Exp Stat Bull 348:1–12 Gullino ML, Garibaldi A (1987) Control of Botrytis cinerea resistant to benzimidazoles and dicarboximides with mixtures of different fungicides. Meded Fac Landbouwwet Rijksuniv Gent 52:895–900 Hampton RO (1963) Seed transmission of white clover mosaic and clover yellow mosaic viruses in red clover. Phytopathology 53:1139 Harri JA, Larsen PO, Powell CC (1977) Bacterial leaf spot and blight of Rieger elatior begonia: systemic movement of the pathogen, host range, and chemical control trials. Plant Dis Rep 61:649–653 Harris KF, Bradley RHE (1973) Importance of leaf hairs in the transmission of tobacco mosaic virus by aphids. Virology 52:295–300 Hausbeck MK, Moorman GW (1996) Managing Botrytis in greenhouse-grown flower crops. Plant Dis 80:1212–1219 Hausbeck MK, Pennypacker SP (1991) Influence of grower activity and disease incidence on concentrations of airborne conidia of Botrytis cinerea among geranium stock plants. Plant Dis 75:798–803 Hoggan IA (1933) Some factors involved in aphid transmission of the cucumber-mosaic virus to tobacco. J Agric Res 47:689–704 Ivors KL, Moorman GW (2014) Oomycete plant pathogens in irrigation water. In: Hong CX, Moorman GW, Wohanka W, B€ uttner C (eds) Biology, detection, and management of plant pathogens in irrigation water. APS Press, St. Paul, pp 57–64 Jacquemond M (2012) Cucumber mosaic virus. Adv Virus Res 84:439–504 Jarvis WR (1980) Epidemiology. In: Coley-Smith JR, Verhoeff K, Jarvis WR (eds) The biology of Botrytis. Academic, London, pp 219–250 Jodon MH, Nichols LP (1974) Bacterial leaf spot of begonia. Pa Flower Grow Bull 272:8–9 Johnson F (1942) The complex nature of white clover mosaic. Phytopathology 32:103–111 Kassanis B, MacFarlane I (1964) Transmission of tobacco necrosis virus by zoospores of Olpidium brassicae. J Gen Microbiol 36:79–98 Kobatake H, Osaki T, Inouye T (1984) Reservoirs of tomato spotted wilt virus in Nara Prefecture. Ann Phytopathol Soc Jpn 50:541–544 Komuro Y, Iwaki M (1968) Bean yellow mosaic virus and tobacco ringspot virus isolated from Crotalaria (Crotalaria spectabilis). Ann Phytopathol Soc Jpn 34:7–15 Law MD, Moyer JW (1990) A tomato spotted wilt-like virus with a serologically distinct N protein. J Gen Virol 71:933–938 Lisa V, Boccardo G (1996) Fabaviruses: broad bean wilt and allied viruses. In: Harrison BD, Murant AF (eds) Polyhedral virions and bipartite RNA genomes, vol 5, The plant viruses. Plenum Press, New York, pp 229–250 Lisa V, Boccardo G, D’Agostino G, Dellavalle G, D’Aquilio M (1981) Characterization of a potyvirus that causes zucchini yellow mosaic. Phytopathology 71:667–672 Lojek JS, Orlob GB (1969) Aphid transmission of tobacco mosaic virus. Science 164:1407–1408 McGuire JM (1964) Efficiency of Xiphinema americanum as a vector of tobacco ringspot virus. Phytopathology 54:799–801 Messieha M (1969) Transmission of tobacco ringspot virus by thrips. Phytopathology 59:943–945 Middleton JT (1942) Stem rot of tuberous begonia. Bull Torrey Bot Club 69:92–99 Moorman GW, Lease RJ (1992) Benzimidazole- and dicarboximide-resistant Botrytis cinerea from Pennsylvania greenhouses. Plant Dis 76:477–480 Neergaard P (1977) Seed-borne viruses, Chapter 3. In: Seed pathology, vol I. MacMillan Press, London/Madras, 839 pp

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Nelson PV, Krauskopf DN, Mingis NC (1977) Visual symptoms of nutrient deficiencies in Rieger elatior begonia. J Am Soc Hortic Sci 102:65–68 Palukaitis P, García-Arenal F (2003) Cucumoviruses. Adv Virus Res 62:241–323 Peirson DQ (1974) Epidemiology of a foliar disease of Rieger elatior begonias caused by Aphelenchoides fragariae (Ritzema Bos). MS, Ohio State University Powell CC (1985) Begonia. In: Strider DL (ed) Diseases of floral crops, vol 1. Praeger Scientific, New York, pp 423–445 Pratt MJ (1961) Studies on clover yellow mosaic and while clover mosaic viruses. Can J Bot 39:655–665 Price WC (1940) Comparative host ranges of six plant viruses. Am J Bot 27:530–541 Quinn JA, Powell CC (1981) Identification and host range of powdery mildew of begonia. Plant Dis 65:68–70 Quinn JA, Powell CC (1982) Effects of temperature, light, and relative humidity on powdery mildew of begonia. Phytopathology 72:480–484 Ribeiro LFC, Mello APD, Bedendo IP, Gioria R (2006) Phytoplasma associated with shoot proliferation in Begonia. Sci Agric 63(5):475–477 Riedel RM (1985) Nematode problems. Diseases of floral crops. In: Strider DL (ed) Diseases of floral crops, vol 1. Praeger Scientific, New York, pp 295–312 Sammons B, Rissler JF, Shanks JB (1982) Development of gray mold of poinsettia and powdery mildew of begonia and rose under split night temperatures. Plant Dis 66:776–777 Samuel G, Bald JG, Pittman HH (1930) Investigations on ‘spotted wilt’ of tomatoes. Aust Counc Sci Ind Res Bull 44:8–11 Schroers HJ, Baayen RP, Meffert JP, de Gruyter J, Hooftman M, O’Donnell K (2004) Fusarium foetens, a new species pathogenic to begonia elatior hybrids (Begonia  hiemalis) and the sister taxon of the Fusarium oxysporum species complex. Mycologia 96(2):393–406 Schuster MF (1963) Flea beetle transmission of tobacco ringspot virus in the Lower Rio Grande Valley. Plant Dis Rep 47:510–511 Sekine T, Kanno H, Aoki T (2008) Occurrence of a leaf and stem rot caused by Fusarium foetens in begoia elatior hybrids (Begonia  hiemalis). Jpn J Phytopathol 74:164–166 Shew HD, Lucas GB (1991) Compendium of tobacco diseases. APS Press, St. Paul Simons JN (1955) Some plant-vector-virus relationships of southern cucumber mosaic virus. Phytopathology 45:217–219 Simons JN, Zitter TA (1980) Use of oils to control aphid-borne viruses. Plant Dis 64:542–546 Smith KM, Bald JG (1935) A description of a necrotic virus disease affecting tobacco and other plants. Parasitology 27:231–245 Smith KM, Markham R (1944) Two new viruses affecting tobacco and other plants. Phytopathology 34:324–329 Stubbs IL (1947) A destructive vascular wilt virus disease of broad bean (Vicia faba L) in Victoria. J Dept Agric Vic 46:323–332 Teakle DS (1962) Transmission of tobacco necrosis virus by a fungus, Olpidium brassicae. Virology 18:224–231 Teakle DS, Gold AH (1963) Further studies of Olpidium as a vector of tobacco necrosis virus. Virology 19:310–315 Thomas C (1969) Transmission of tobacco ringospot virus by Tetranycus sp. Phytopathology 59:633–636 Thompson ML, Thompson EJ (1981) Begonias: the complete reference guide. Times Books, New York Tian X, Zheng Y (2012) Species susceptibility and biological control of Fusarium wilt of Hiemalis begonias in Canada. Can J Plant Pathol 34:345–346 Tian X, Zheng Y (2013) Evaluation of biological control agents for Fusarium wilt in Hiemalis begonia. Can J Plant Pathol 35:363–370 Tian XL, Dixon M, Zheng YB (2012) Susceptibility of various potted begonias to Fusarium foetens. Can J Plant Pathol 34:248–254

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Tompkins CM (1950) Botrytis stem rot of tuberous-rooted begonia. Hilgardia 19:401–410 Tschope B, Hey M, Wohanka W, Hennig F (2007) Characterisation and identification of Fusarium foetens, causative agent of wilting and stem rot of begonia elatior hybrids (Begonia  hiemalis) by its volatile compounds. Eur J Hortic Sci 72(4):152–157 Ullman DE, German TL, Sherwood JL, Westcot DM, Cantone FA (1993) Tospovirus replication in insect vector cells: immunocytochemical evidence that the nonstructural protein encoded by the S RNA of tomato spotted wilt tospovirus is present in thrips vector cells. Phytopathology 83:456–463 Van der Gaag D, Raak M (2010) Pest risk assessment Fusarium foetens. Plant Protection Service, Ministry of Agriculture, Nature and Food Quality, Wageningen. Bulletin 11-16495 Watson MA, Roberts FM (1939) A comparative study of the transmission of Hyoscyamus virus 3, potato virus Y and cucumber virus 1 by the vector Myzus persicae (Sulz.), M. circumflexus (Buckton) and Macrosiphum gei (Koch). Proc R Soc Ser B 127:543–576 Wijkamp I, van Lent J, Kormelink R, Goldbach R, Peters D (1993) Multiplication of tomato spotted wilt virus in its insect vector, Frankliniella occidentalis. J Gen Virol 74:341–349 Yang X, Richardson PA, Olson HA, Hong CX (2013) Root and stem rot of begonia caused by Phytopythium helicoides in Virginia. Plant Dis 97(10):1385 Zhao G, Liu W, Brown JM, Knowles CO (1995) Insecticide resistance in field and laboratory strains of western flower thrips (Thysanoptera: Thripidae). J Econ Entomol 88:1164–1170 Zitter TA, Murphy JF (2009) Cucumber mosaic. Plant Health Instructor. doi:10.1094/PHI-I-20090518-01

Diseases of Celosia Ann B. Gould

Abstract

Celosia (Celosia L.), generally known as cockscomb or woolflower, is a genus of ornamental and edible tender annuals in the Amaranthaceae. Ornamental cultivars are subject to few diseases, although fungal and bacterial leaf spots, Botrytis blight, Rhizoctonia root and stem rot, damping-off, and root-knot nematode may occur where humidity is high or where good growing practices, sanitation, and other cultural disease management practices that reduce inoculum and spread of propagules are not followed. Keywords

Alternaria spp. • Macrophomina phaseolina • Rhizoctonia solani • Meloidogyne spp.

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-Like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Alternaria Leaf Spot and Blight (= Nimbya Leaf Spot) [Alternaria sp.; A. alternata (Fr.) Keissl.; A. celosiicola Jun. Nishikawa & C. Nakashima (Basionym: Nimbya celosiae E.G. Simmons & Holcomb ( A. celosiae (E.G. Simmons & Holcomb) Lawrence, Park & Pryor)), A. gomphrenae Togashi] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Botrytis Blight, Botrytis Stem Rot (Botrytis cinerea Pers.:Fr.) . . . . . . . . . . . . . . . . . . . . . 2.3 Cercospora Leaf Spot (Cercospora celosiae Syd.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Charcoal Rot, Collar Rot (Macrophomina phaseolina (Tassi) Goid.) . . . . . . . . . . . . . . . 2.5 Fusarium Canker, Blight, Leaf Spot (Fusarium lateritium f. sp. celosiae (Abe) Matuo) [ Fusarium lateritium Nees) (tel. Gibberella baccata (Wallr.) Sacc]) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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A.B. Gould (*) Department of Plant Biology, School of Environmental and Biological Sciences, Rutgers, The State University of New Jersey, New Brunswick, NJ, USA e-mail: [email protected] # Springer International Publishing AG 2017 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florist's Crops Diseases, Handbook of Plant Disease Management, DOI 10.1007/978-3-319-32374-9_31-1

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Phytophthora Root and Stem Rot, Damping-off (Phytophthora cryptogea Pethybr. & Laff.; P. drechsleri Tucker; P. nicotianae Breda de Haan (syns. Phytophthora nicotianae var. parasitica, P. parasitica) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7 Powdery Mildew (Erysiphe celosiae Tanda; Leveillula taurica (Lév.) G. Arnaud) . 2.8 Pythium Blight, Damping-off (Pythium irregulare Buisman; P. ultimum Trow) . . . . 2.9 Rhizoctonia Root and Crown Rot, Blight, Damping-off [Rhizoctonia solani Kühn (tel. Thanatephorus cucumeris (A.B. Frank) Donk)] . . . . . . . . . . . . . . . . . . . . . . . . . 2.10 Verticillium Wilt (Verticillium albo-atrum Reinke & Berthold; Verticillium dahliae Kleb.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Bacterial and Phytoplasma Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Bacterial Leaf Spot (Dickeya chrysanthemi (syn. Erwinia chrysanthemi Burkholder et al.); Pseudomonas syringae pv. tabaci (Wolf & Foster) Young, Dye & Wilkie; Xanthomonas sp. Dowson) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Seedling Blight (Pseudomonas syringae van Hall) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Phytoplasma Diseases (“Candidatus Phytoplasma spp.”) . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Viral Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Cucumber Mosaic (Cucumber mosaic virus) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Beet Curly Top (Beet curly top virus) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Tomato Spotted Wilt, Impatiens Necrotic Spot (Tospovirus) . . . . . . . . . . . . . . . . . . . . . . . . . 4.4 Celosia Mosaic (Celosia mosaic virus) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Nematode Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 Root-Knot (Meloidogyne spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Introduction

Celosia L., also known generally as celosia or as woolflower (plumed flowers), cockscomb (flowers crested by fasciation), or spiked (wheat) cockscomb, is a genus of ornamental and edible herbaceous tender annuals in the Amaranthaceae (pigweed family). Of tropical origin, the genus is thought to originate in Africa or Asia. Cultivated species include C. argentea L. (silver cockscomb) (preferred scientific name), C. cristata L. (crested cockscomb), C. nitida Vahl (West Indian cockscomb), C. palmeri S. Watson (Palmer’s cockscomb), C. trigyna L. (woolflower), and C. virgata Jacq. (albahaca) (ITIS 2016). In the older literature and in trade, cockscombs may be listed as variants of C. argentea (e.g., C. argentea var. argentea, C. argentea var. cristata, C. argentea var. plumosa, C. argentea ‘Spicata’, or C. plumosa). The leaves and brightly colored flowers are edible: C. argentea is cultivated as a leafy green in tropical regions of Africa, South America, and Southeast Asia (National Research Council 2006). As an ornamental, celosia adds colorful interest in gardens (beds, borders), containers, as cut flowers or in dried floral arrangements, or indoors with sufficient light. Celosia is also an herbaceous annual weed in many cropping systems (e.g., groundnut, pearl millet, cotton, maize) in India (Nalini et al. 2015; Ravindra et al. 2008). Celosia is propagated from seed, and in warm weather the annual grows well in full-sun and in moist but well-drained soils, flowering until mid- to late-summer in temperate regions (Nau 1991; Gilman and Howe 1999). The annual is subject to few diseases; leaf spots, Botrytis bight, damping-off, and root, stem, and crown rots

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Fig. 1 Alternaria leaf spot and blight (Photo courtesy of R. J. McGovern)

(non-water molds or oomycetes) may occur under conditions of high moisture; other reported issues include powdery mildew, root-knot nematode, and phytoplasma and virus diseases (Blake and Williamson 2015; Farr et al. 1989; Horst 2008; Horst 2013; Pundt 1993; Raabe et al. 2009; Wehlburg et al. 1975; Wilson 1932; Tanne et al. 2000). Cold damage and high soluble salts in growing media may also affect celosia production (Chase and Daughtrey 2012). Most of the disease information in this chapter pertains to Celosia sp. grown as an ornamental. In some regions of the world, however, both edible and ornamental crops are cultivated; the distinction between these is made where appropriate.

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Fungal and Fungus-Like Diseases

2.1

Alternaria Leaf Spot and Blight (= Nimbya Leaf Spot) [Alternaria sp.; A. alternata (Fr.) Keissl.; A. celosiicola Jun. Nishikawa & C. Nakashima (Basionym: Nimbya celosiae E.G. Simmons & Holcomb ( A. celosiae (E.G. Simmons & Holcomb) Lawrence, Park & Pryor)), A. gomphrenae Togashi]

Geographic Occurrence and Impact Species of Alternaria have a wide host range, causing leaf spots and blights on numerous ornamental crops. One of several leaf spot diseases of celosia, Alternaria leaf spot (as one or more of the species listed above) in production (plug trays and finishing) and landscape has been reported in the USA (on Celosia argentea and C. argentea var. cristata, Farr et al. 1989, United States Agricultural Research Service 1960; on Celosia sp., Ivey and Iverson 2005), Japan (on C. argentea var. plumosa and C. argentea var. cristata, Nishikawa and Nakashima 2013), South America (on C. argentea, Cúndom and Cabrera 2002), Thailand (on Celosia sp., Seemadua et al. 2011), other parts of South and Southeast Asia and the Caribbean (on C. plumosa, Yuchida 1998), and in Africa on C. argentea

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grown as a vegetable (Yarger 2007). Holcombe (1978) reported that a species of Alternaria (not listed above) from alligatorweed, A. alternantherae, in Louisiana was pathogenic to ornamental C. argentea inoculated in the greenhouse, causing severe infection and moderate to severe defoliation. Outdoors, the disease is more common during wet seasons (Pirone 1978). Symptoms/Signs Leaf spotting appears on older leaves as small, reddish purple spots that turn tan to gray (concentric rings may be evident) with a red to purple border as they enlarge (up to 8 mm diameter) (Seemadua et al. 2011) (Fig. 1). Spots may also appear on bracts, petioles, or stems, and blighting of leaves, shoots, and flowers may occur; plants are most heavily diseased during cool, wet weather or during stress (Agrios 2005; Yuchida 1998). Although Alternaria leaf spot may occur during plug production, it is most evident at finishing or in landscape plantings (Chase and Daughtrey 2012). Biology and Epidemiology Abundant Alternaria spores (conidia) are produced in lesions during moist weather. Conidia are pale to brown, multicellular with longitudinal and transverse septa, and borne in chains on short conidiophores. The fungus survives adverse conditions as mycelium or spores associated with infected plant debris or seeds and is spread by moving air, splashing water (rain or irrigation), mechanically during handling, or movement of contaminated seeds or plant material (Agrios 2005; Gleason et al. 2009). Management Management of leaf spot diseases benefits from strategies that reduce inoculum: discard severely diseased plants, use pathogen-free seed, minimize moisture and humidity in the canopy (e.g., careful watering, avoiding overhead irrigation, proper spacing and ventilation), and consider a protectant fungicide or biological product if conditions conducive to disease development are expected. Sanitation practices include cleaning production areas or gardens of diseased plants and debris to reduce any surviving inoculum (Chase and Daughtrey 2012; Yuchida 1998). In addition, leaf spot diseases may be more severe on plants weakened by environmental factors or poor growing conditions (Douglas 2012).

2.2

Botrytis Blight, Botrytis Stem Rot (Botrytis cinerea Pers.:Fr.)

Geographic Occurrence and Impact Also known as “gray mold,” Botrytis blight is a very common and widely distributed disease of over 170 families of cultivated plants, including flowering crops (Elad et al. 2016a). Botrytis cinerea, a nonhost specific pathogen, readily colonizes nonliving organic matter as a saprophyte and, under the right conditions, can also attack leaf, floral, stem, bud, fruit, root, bulb, and crown tissues as a necrotroph. In celosia, the pathogen causes a stem canker and blight on the inflorescence and leaves when humidity is high, causing direct crop loss and significantly reducing yield or suitability as cut flowers or in dried arrangements.

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Fig. 2 Botrytis leaf and stem blight (Photo Courtesy of North Carolina State University)

Symptoms/Signs On hosts affected by gray mold, leaves and petals may appear water soaked at first, progressing to irregular, sunken spots or larger lesions that fade to tan to dark (Chase et al. 1995) (Fig. 2). Flowers will fail to open if infected during the bud stage. Under conditions of high humidity, the fungus produces a characteristic gray, fuzzy growth (mycelium and spores) on infected tissue (Agrios 2005). This aerial mycelium may also be evident on affected celosia stems, which turn gray to black and may break at the point of infection. Biology and Epidemiology Epidemics of gray mold are common in enclosed structures, such as greenhouses, and outdoors where moisture is high. Abundant conidia (asexual spores) and aerial mycelium of B. cinerea are associated with diseased tissues when humidity is high (>90%) and a thin film of water is present for 4 h or more (Carisse 2016). Spores are hyaline, ovoid, and borne in bunches (botryose) on branched conidiophores. These spores readily disperse by air currents throughout the greenhouse or outdoors, penetrating host tissues, especially floral organs, most often through wounds. Spores may also lodge on stems and other plant parts, readily penetrating wounds to cause stem cankers or cutting rots during propagation. On many hosts, the fungus produces sclerotia: black, elliptical (up to 5 mm/0.2 in. diameter) survival structures that are associated with dead, colonized tissues. Sclerotia, which consist of a compact mass of hyphae with a melanized rind, are resistant to adverse conditions and serve as an inoculum source for new infections. The fungus also survives the noncropping period as mycelium in plant debris, and can remain asymptomatically as an endophyte in plant tissues (Elad et al. 2016b). Management • Cultural practices – Botrytis diseases benefit from strategies that reduce moisture (tissue wetness and humidity) and wounding. Since moisture is required for spore germination and penetration, careful watering, avoiding overhead irrigation or watering early in the day, and proper spacing and ventilation in production areas are required (Elad 2016). In shops, keep flowers and cooler floors dry

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(Wright 2011). Avoid unnecessary mechanical injury during production, harvest, and transport to reduce the number of infection courts for the fungus. In celosia marketed as cut flowers, plants are harvested 3–4 week after the flower matures. To avoid issues with gray mold after harvest, these hardened stems must be dry and kept at a constant temperature at storage and during transport to prevent condensation (Zuck 2015). Stems that are wounded by poor handling at harvest or by inadequate packing prior to transport may become colonized by the fungus when moisture becomes available. Process flowers quickly and carefully, ensuring good air flow to keep blooms dry. Control insects during production, which create wounds in plants. Sanitation practices reduce inoculum; these include cleaning production areas or outdoor beds of diseased plants and debris where the fungus may have survived in absence of its host (Chase and Daughtrey 2012; Elad 2016). In addition, senescent and dead plant tissues are readily colonized and may serve as a “base” from which the fungus spreads to healthy tissue. Proper management of Botrytis blight includes monitoring. Purchase clean stock and ask suppliers about Botrytis management practices and shipping conditions. Look for symptoms (including fluffy gray mold on infected plant parts) when weather conditions favor the pathogen (wet, overcast, and very humid weather). If symptoms are present but sporulation is not evident, place affected tissue in a moist chamber for 1–2 days to encourage the development of aerial mycelium. Avoid plant stress. High salts, compacted planting media, uneven watering, and fertilization that does not meet the needs of the plant reduce the ability of plants to resist disease. • Chemical management of Botrytis diseases on ornamental hosts, including celosia, includes application of preventive fungicides when environmental conditions are conducive to disease development. Chemical applications combined with cultural management strategies to reduce inoculum may be necessary to control Botrytis blight in many cropping situations, especially for high value plants (Fillinger and Walker 2016). Due to the genetic plasticity of the Botrytis fungus (Carisse 2016), fungicide resistance has developed to several commonly used classes of fungicides worldwide (Beever et al. 1989; Bollen and Scholten 1971; Katan 1982; Leroch et al. 2011; Leroux et al. 2002; Lamondia and Douglas 1997; Yourman and Jeffers 1999; Zhao et al. 2010). A treatment of fungicide mode of action, resistance development, and fitness costs in Botrytis sp. may be found in reviews compiled by Fillinger and Walker 2016, Kretschmer et al. 2009, and Leroux 2007. As some populations of Botrytis may have partial resistance to antifungal compounds, strategies to reduce fungicide resistance development include reducing inoculum through cultural practices, alternating or tank-mixing compounds with differing modes of action, and development of new chemistries to maintain a diversity of mode of actions (Beever and Brien 1983; Fillinger and Walker 2016; Rosslenbroich and Stuebler 2000).

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• Biological control – In addition to synthetic fungicides, the use, efficacy, and durability of biological control agents and biopesticides for gray mold management have been assessed (Nicot et al. 2016) in an effort to identify materials that can be applied to Integrated Crop Management (ICM) or organic cropping methods. Many of these products inhibit the activity of B. cinerea in the greenhouse or laboratory, but these vary in efficacy and mode of action and few are commercialized. Those biologics with activity against Botrytis in ornamental plants include formulations containing Reynoutria sachalinensis, Bacillus subtilis QST 713, Gliocladium catenulatum J1446, Streptomyces griseoviridis K61, and S. lydicus WYCD108. Other materials include neem, soybean oils, ammonium chloride, hydrogen dioxide, and potassium bicarbonates. Although these products have general labels for ornamental plants, none include celosia-specific phytotoxicity or application information.

2.3

Cercospora Leaf Spot (Cercospora celosiae Syd.)

Geographic Occurrence and Impact Cercospora spp. cause a leaf spot or blight on a wide variety of host plants (Agrios 2005). C. celosiae has been reported on celosia in the USA (on Celosia cristata, Ellett 1989; on C. argentea and C. argentea var. cristata, Farr et al. 1989), Asia (on C. cristata, C. plumosa, Braun and Sivapalan 1999; on C. argentea var. cristata, Groenewald et al. 2013, Nguanhom et al. 2015; on C. argentea, To-Anun et al. 2011), South America (on Celosia sp., Standen 1952), Africa (on C. argentea grown as a vegetable [Legos spinach], Yarger 2007), Malaysia (on C. cristata, Turner 1967), and the Indian subcontinent (on C. argentea, Kew Royal Botanic Gardens 2015). As with other leaf spot diseases of celosia, the disease is more common during wet seasons (Pirone 1978). Symptoms/Signs Leaf spots on celosia appear as irregularly shaped, pale to brown lesions (up to 7 mm/0.3 in. diameter) surrounded by a red to dark brown border (Groenewald et al. 2013); infected tissues sometimes drop away from the leaf, leaving holes (known as shot-holes). Biology and Epidemiology C. celosiae spores (conidia) are hyaline, straight to curved, multiseptate, and are borne on pale to olive-brown groupings of conidiophores within small, brown compact masses of hyphae (fruiting structures) called stromata (Groenewald et al. 2013). Conidia are easily detached and spread by moving air. Moisture or high humidity and warmer temperatures favor disease development, and the fungus survives adverse conditions associated with seed or within stromata in old leaf lesions (Agrios 2005). Management Management of leaf spot diseases benefits from strategies that reduce inoculum as described for Alternaria leaf spot in section 2.1. In addition, use of disease-free seed and potting material is helpful (Agrios 2005).

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Charcoal Rot, Collar Rot (Macrophomina phaseolina (Tassi) Goid.)

Geographic Occurrence and Impact Macrophomina phaseolina is a widely distributed soilborne pathogen that affects the roots and lower stems of more than 500 host plants, notably soybean, maize, cotton, and sunflower, causing a root rot, charcoal rot, collar rot, and damping-off and seedling blight worldwide (Ijaz et al. 2013; Sinclair 1982; Smith and Carvil 1997). In a pot culture experiment, Singh et al. (1990) reported that a M. phaseolina isolate from chickpea was pathogenic to a Celosia argentea, a new host. The fungus was also reported in Pakistan in an outdoor planting of C. argentea in 2011 (Rehman et al. 2015). In the USA, charcoal rot (as Sclerotium batatitcola) of celosia (C. argentea var. cristata) has been reported in Texas (US Agricultural Research Service 1960). Symptoms/Signs In general, disease symptoms caused by M. phaseolina include a rot of the roots and lower stem. Infected seedlings exhibit sunken cankers at the soil line. Aboveground, leaf yellowing and premature leaf drop and plant death are common. Rehman et al. (2015) reported symptoms of charcoal/collar rot on C. argentea as chlorosis and wilt of the lower leaves during warm dry weather. In addition, the roots and stem collar were shrunken and shredded. In C. argentea grown as a vegetable, infection by M. phaseolina results in dark spots on the leaves (Yarger 2007). Biology and Epidemiology M. phaseolina produces numerous small, black, opaque microsclerotia (50–150 μm diameter) just under the epidermis that serve as survival structures in soil and crop residues and as a source of primary inoculum (Gupta et al. 2012; Sinclair 1982). Typically, development of charcoal rot is favored by hot, dry weather. Rehman et al. (2015) noted that charcoal/collar rot of celosia in Pakistan was more severe during warmer, dry conditions (30 C/82 F and 95%) was followed by lower relative humidity. Like other Peronospora spp. (Pinckard 1942), sporangial release of coleus downy mildew followed a daily periodicity with sporangial counts highest between 1000 and 1500 h. Management • Cultural control – Coleus should be scouted on a regular basis for signs and symptoms of downy mildew. Infected plants should be kept separate and isolated from healthy plants, as the sporangia can be airborne. Intervals between scouting events should be shortened during periods of high relative humidity. Environmental conditions that may limit downy mildew include keeping relative humidity below 85% and temperatures at or over 25  C/77  F. Limiting extended leaf wetness periods by watering in the morning or early afternoon will ensure that the foliage does not remain wet overnight. Increased plant spacing will ensure better ventilation and shorter leaf wetness periods. • Sanitation – Removing downy mildew-infected plant material is essential (Harlan and Hausbeck 2011). Although Peronospora is an obligate pathogen, needing live plant material to reproduce, the sporangia on abscised leaves may be viable for a short time. • Fungicides and biocontrols – If effective fungicides are used, downy mildew on coleus can be adequately controlled (Harlan and Hausbeck 2010). Fungicide active ingredients that have been shown to be highly effective in limiting Peronospora infection on coleus include mefenoxam, dimethomorph, fluopicolide, azoxystrobin, fosetyl-al, mandipropamid, fluoxastrobin, and fenamidone (Harlan and Hausbeck 2010). Research on other Peronospora species has shown that drench applications of mefenoxam, fluopicolide, and oxathiapiprolin have offered long-term protection from infection. In particular,

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B.R. Harlan and M.K. Hausbeck

these applications, when applied to greenhouse plants, have offered protection from downy mildew long after transplanting into the landscape (Harlan and Hausbeck 2016). Due to the prevalence of the coleus downy mildew pathogen in recent years, preventive fungicide applications are warranted. Biocontrol products have been tested against Peronospora spp. with limited success. An extract of Reynoutria sachalinensis, when applied as a preventive spray, has shown some efficacy against coleus downy mildew (Harlan and Hausbeck 2010; Ivors et al. 2011). In a trial conducted by Warfield et al. (2008), spray applications of a phosphorous acid salts product were highly effective. • Resistance – With the vast number of different coleus cultivars commercially sold and the varying colors and shapes observed among these cultivars, it is not surprising that a wide range of susceptibility to downy mildew has been observed. Daughtrey et al. (2014) tested 147 cultivars over a multiyear study and rated 32% of the cultivars with low susceptibility, while 27.2% were rated as highly susceptible to downy mildew. In particular, all 11 cultivars in the Wizard series were rated as having medium to high susceptibility, while seven of the nine Fairway series cultivars were rated as having a low or medium susceptibility. When growing highly susceptible cultivars, it is recommended that preventive fungicides be applied prior to disease development.

2.3

Pythium Root Rot (Pythium spp.)

Geographic occurrence and impact. This disease has been reported in Florida, Mississippi, California, and Maryland (Alfieri et al. 1984; Anonymous 1960; Parris 1959). Symptoms/signs. Pythium is a water mold that can “nibble” the feeding roots of plants, resulting in wilting and stunted growth. This reduced plant vigor can eventually affect marketability (Garzon et al. 2011). In severe cases, stems of seedlings or recent transplants collapse at the soil line and die (Pscheidt and Ocamb 2016). Plants that survive the initial infection phase are likely to remain stunted. Both cutting- and seed-propagated cultivars of coleus are susceptible. If Pythium root rot is suspected on a plant showing aboveground symptoms, the root system should be inspected. Infected roots will be brown or water-soaked and sparse when compared to a healthy root system (Fig. 2). Biology and epidemiology. Pythium species are soilborne oomycete organisms that prefer wet conditions for infection and reproduction. This preference for wet conditions explains Pythium’s characterization as a water mold. Three distinct spore types are produced by Pythium: oospores, sporangia, and zoospores. Oospores are thick-walled sexually produced spores that can lay dormant for extended periods. Oospores can withstand long periods of drying, and once environmental conditions are conducive, or a suitable host is presented, they can germinate and infect via fungal-like strands. Sporangia and zoospores are the most common spore type associated with Pythium epidemics as they are produced in large numbers. Zoospores, in particular, are associated with the spread of the pathogen in ebb-and-flow

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Fig. 2 Pythium root rot symptoms: Infected roots are brown, water soaked, and sparse (Photo courtesy of M. Daughtrey)

irrigation systems as this particular spore type is motile in water (Koike and Wilen 2009). Management • Cultural control – Overwatering, creating soil conditions that are overly saturated for extended periods, favors Pythium development. In particular, soil moisture conditions of 70% or higher are conducive to infection by Pythium. Keep fungus gnats and shore flies under control as they can spread the pathogen (Koike and Wilen 2009). • Sanitation – This pathogen can become a greenhouse “resident” that hibernates on dirty plant containers, benches, hoses, and greenhouse walkways (Hausbeck and Harlan 2013). The reuse of flats, pots, and containers is not recommended; however, if they are reused, they should be thoroughly disinfected with a cleaning agent such as quaternary ammonium chloride salts or chlorine bleach. (Refer to chapter “▶ Sanitation in Florists’ Crops Disease Management” for additional information about disinfectants.) Plants with a suspected Pythium infection should be isolated from healthy plants to reduce the further spread of the pathogen. • Fungicides and biocontrols – Unfortunately, there are very few effective fungicides registered against Pythium root rot. When attempting to control root rot pathogens with fungicide drenches, it is important to make applications to the soil. Applications of the low risk fungicide mefenoxam can be highly effective in limiting Pythium infection. However, resistance of the pathogen to mefenoxam is an issue (Del Castillo Munera and Hausbeck 2016). Etridiazole is another product that is highly effective against Pythium. Other products, including fluopicolide, fenamidone, and phosphorous acid salts, have shown promise in managing Pythium (Hausbeck and Harlan 2013). Although using biocontrol against Pythium in a greenhouse can be helpful, studies have shown the level of control to be inconsistent. For instance, in a

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study published by Daughtrey and Tobiasz (2001), four popular biocontrol products were tested against Pythium, and all received ratings statistically similar to the untreated control plants. However, Baysal-Gurel and Miller (2013) found applications of biocontrol products effective in limiting Pythium damping-off in the greenhouse. • Resistance – At this time there is no published information on the resistance of coleus to Pythium spp.

2.4

Rhizoctonia Root and Crown Rot (Rhizoctonia solani)

Geographic occurrence and impact. This disease has been reported in Florida, Illinois, New York, and Texas (Alfieri et al. 1984; Anonymous 1960). Symptoms/signs. Rhizoctonia can cause pre- and postemergence damping-off of coleus. Younger plants are more susceptible (Wright 2013). Infected stems will appear blackened and water-soaked (Pscheidt and Ocamb 2016). In severe cases, Rhizoctonia can move into the plant canopy, causing spots on leaves and petioles. Root rot symptoms are similar to those caused by Pythium, and a diagnostic clinic may be necessary to differentiate with which pathogen you are dealing. Biology and epidemiology. Rhizoctonia is a saprophytic fungal pathogen in the phylum Basidiomycota that is associated with several greenhouse crops throughout the world. Rhizoctonia is not host-specific, meaning one isolate can infect several different species of plants (Wright 2013). Unlike many of the other pathogens associated with greenhouse production, spores are not the usual method of dissemination for Rhizoctonia. Mycelia, the hairlike fungal strands produced by Rhizoctonia, are spread by the movement of infested soil particles, often by workers, which grow and infect nearby hosts. Sclerotia, the resting structure produced by Rhizoctonia, can survive for extended periods in soil particles. Rhizoctonia can thrive under almost all greenhouse conditions, although warmer, more humid conditions may enhance its spread. Prolonged periods of high humidity can result in the pathogen growing from the soil, up the stem, and into the foliage canopy. Management • Cultural control – One of the primary sources of Rhizoctonia inoculum in the greenhouse are infested trays (Gutierrez et al. 2001). The reuse of trays, pots, and other containers is not recommended; however, if they are reused, they should be thoroughly disinfected with a cleaning agent such as quaternary ammonium chloride salts or chlorine bleach. • Sanitation – Between crops, clean soil, and organic material from benchtops and floors as this material can harbor the pathogen for extended periods. Because Rhizoctonia does not produce spores, one of the more common methods of transmission is by workers. All pruning tools should be disinfected between crops and foot baths should be installed between greenhouses. Greenhouse carts should also be thoroughly cleaned. • Fungicides and biocontrols – Moorman and Lease (1999) conducted a study looking at soil drenches to control Rhizoctonia rot on coleus and found that

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azoxystrobin and thiophanate-methyl were effective at limiting infection. Other products that have consistently reduced Rhizoctonia on ornamental plants include fludioxonil, PCNB, metconazole, and pyraclostrobin. The type of symptoms observed should determine the fungicide application method; drench applications should be used for root and crown rots, while high volume sprays or “sprenches” should be used to control foliar symptoms. The biopesticide, polyoxin D zinc salt, is highly effective against Rhizoctonia (Hausbeck and Harlan 2015). Using biocontrols in potting mixes may be helpful in managing damping-off when the plants are still young and more susceptible to the pathogen. • Resistance – At this time there is no published information on the resistance of coleus to Rhizoctonia.

3

Viral Diseases

3.1

Bromoviridae

Geographic occurrence and impact. Viruses in the Bromoviridae are some of the most economically important and widely distributed viruses in the world. One member of this group has been reported in coleus: Cucumber mosaic virus (CMV, genus: Cucumovirus). Cucumber mosaic virus has been reported in Louisiana (Holcomb and Valverde 1991). Symptoms/signs. Symptoms of CMV infection of coleus include ring-spotting, mosaic, and oak-leaf symptoms. Biology and epidemiology. CMV is spread by more than 80 aphid species (Hemiptera: Aphididae) in a non-circulative (nonpersistent) manner. When these insects feed on CMV-infected plants, the virus moves into the mouthparts of the vector and can spread when the insect moves and feeds on adjacent plants. Although seed transmission has been reported on other crops (Neergaard 1977), it has not been reported or studied on coleus. Management. There is no cure or chemical control for plant viruses. New shipments of plants should be inspected for symptoms of viruses upon arrival. Vector control is the most important management aspect associated with CMV on coleus. This management plan should include monitoring insect populations and applying chemical controls when needed. Insect populations can be tested for CMV infection. Stock plants should be tested regularly for CMV, and infected plants should be discarded. Refer to chapter “▶ Insect Management for Disease Control in Florists’ Crops” for additional virus management information.

3.2

Bunyaviridae

Geographic occurrence and impact. Viruses belonging to the Bunyaviridae include those most commonly associated with ornamental crops. Impatiens necrotic

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spot virus (INSV, genus: Tospovirus) has been reported on coleus in New York (Catlin 2014), Michigan (Byrne 2008), and France (Lambert 2005). Symptoms/signs. Ring spots are the most typical symptom associated with INSV on coleus (Fig. 3). Other symptoms of INSV include leaf mottle, stem or petiole lesions, irregularly shaped necrotic lesions, or plant stunting (Catlin 2014). Biology and epidemiology. INSV is spread by the western flower thrips (Frankliniella occidentalis) as well as two closely related species F. fusca and F. intonsa. Unlike viruses in the Bromoviridae family, INSV enters the gut of the vector where it replicates. Thrips must be in their juvenile stage (first and early second larval stages) to acquire the virus, and only immature thrips that acquire these viruses or adults derived from such immatures are vectors. Management. There is no cure or chemical control for plant viruses. New shipments of plants should be inspected for symptoms of viruses upon arrival. Western flower thrips populations should be closely monitored. In particular, flowers should be inspected, as thrips are likely to reside in such protected areas. If thrips are detected, chemical controls might be necessary. Insect populations can be tested for INSV incidence. Since almost all herbaceous plants associated with horticulture can harbor INSV, greenhouses should be scouted regularly for signs of infection. Any plants showing symptoms of INSV should be disposed of immediately.

3.3

Potyviridae

Geographic occurrence and impact. Potyviruses are fairly common and infect many crops. One member of this group has been recently discovered in coleus:

Fig. 3 Symptoms of INSV: ring spots and irregularly shaped necrotic spots (left, Photo courtesy of L. Pundt; right, Photo courtesy of M. Daughtrey)

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Tobacco etch virus (TEV, genus: Potyvirus). TEV has been reported on coleus in Missouri and Minnesota (Lockhart et al. 2010). Symptoms/signs. Symptoms of TEV are similar to those associated with coleus vein necrosis virus and include foliar and veinal lesions. Biology and epidemiology. At this time, a number of aphid species are the only known vectors of TEV (Sikora 2004). The virus has a broad host range (>120 species in 19 dicot families) including a number of solanaceous crops such as pepper, tomato, and tobacco. Management. There is no cure or chemical control for plant viruses. New shipments of plants should be inspected for symptoms of viruses upon arrival. Greenhouses should be scouted regularly for aphids. If aphids are detected, chemical controls might be necessary. Due to the rarity of this virus on coleus, plants showing symptoms of a viral infection should first be tested for INSV and CMV.

3.4

Betaflexivirdae

Geographic occurrence and impact. Of the virus families associated with coleus, viruses in the Betaflexivirdae are the least common and economically important. A new virus was observed on coleus in 2005 and was named coleus vein necrosis virus (CVNV, genus: Carlavirus) by Mollov et al. (2007). Symptoms/signs. Symptoms of CVNV on coleus include abnormal leaf coloration, vein necrosis, and ring patterns (Fig. 4). Biology and epidemiology. It is unknown how this virus is spread in the environment. In laboratory studies, mechanical viral transmission was successful. However, when aphids (Myzus persicae) were tested as a possible vector, transmission of CVNV was not observed (Mollov et al. 2007). Management. There is no cure or chemical control for plant viruses. New shipments of plants should be inspected for symptoms of viruses upon arrival. Due Fig. 4 Symptoms of CVNV: abnormal leaf coloration and ring patterns (Photo courtesy of M. Daughtrey)

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to the rarity of CVNV, if virus symptoms are observed on coleus, testing for INSV and CMV should be completed as they are much more common. At this time, there is not a commercially available test for CVNV.

4

Viroid Diseases

4.1

Pospiviroidae

Geographic occurrence and impact. Viroids are small, single-stranded sections of RNA that do not have protein coats. The coleus blumei viroids (CbVd, genus: Coleviroid) are widely distributed and have been classified as six different species, CbVd-1 through CbVd-6. CbVd-1 was first detected in Brazil in 1989 and is the most common species and has been reported worldwide (Fonseca et al. 1989; Tsushima and Sano 2015). CbVds has been reported in Germany, China, India, Indonesia, and Japan (Spieker et al. 1990; Hou et al. 2009; Jiang et al. 2013; Tsushima and Sano 2015). Symptoms/signs. A wide range of symptoms can be observed on CbVd-infected coleus. In fact, CbVd is often detected on plants that appear to be symptomless and healthy (Fu et al. 2011). Stunting has been identified as a symptom, although yellowing or chlorosis-type symptoms are the most common (Tsushima and Sano 2015). This yellowing can be observed on the leaves in patches, often irregular in shape (Adkar-Purushothama et al. 2013). Biology and epidemiology. Very little is known on how this pathogen infects and spreads on coleus. Jiang et al. (2014) did show that seed transmission is possible and vegetative cuttings are likely to spread the viroid. Management. There is no cure or chemical control for plant viroids. Getting seed and cuttings from stock plants that have been tested for CbVd will reduce the spread of the disease.

References Adkar-Purushothama CR, Nagaraja H, Sreenivasa MY, Sano T (2013) First report of Coleus blumei viroid infecting coleus in India. Plant Dis 97:149 Alfieri SA Jr, Langdon KR, Wehlburg C, Kimbrough JW (1984) Index of plant diseases in Florida. Bulletin No 11 (revised). Florida Department of Agriculture and Consumer Services, Division of Plant Industry, Gainesville, p 389 Anonymous (1960) Index of plant diseases in the United States. USDA handbook no 165. Washington, DC, p 531 Anonymous (2008) Coleus downy mildew. Plant Clinic News, July. Central Science Laboratory. http://fera.co.uk/news/resources/documents/PCN%20-%20plantClinicNews0708.pdf. Accessed 25 Mar 2016 Anonymous (2015) Floriculture crops 2014 summary. USDA National Agricultural Statistics Service. http://usda.mannlib.cornell.edu/MannUsda/viewDocumentInfo.do?documentID= 1072. Accessed 7 Oct 2016

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Baysal-Gurel F, Miller SA (2013) Evaluation of fungicides and biorational products for management of Pythium and Rhizoctonia damping-off in greenhouse-produced vegetables. Phytopathology 103(Suppl 2):S2.13 Byrne J (2008) Diagnostic update for greenhouse samples. Michigan State University Extension News for Agriculture-Floriculture, Mar 28. http://msue.anr.msu.edu/news/diagnostic_update_ for_greenhouse_samples. Accessed 7 Oct 2016 Byrne JM, Hausbeck MK, Sconyers LE (2005) Influence of environment on atmospheric concentrations of Peronospora antirrhini sporangia in field-grown snapdragon. Plant Dis 89:1060–1066 Catlin N (2014) INSV on coleus. e-GRO Alert 3(37). http://www.e-gro.org/pdf/337.pdf. Accessed 7 Oct 2016 Daughtrey ML, Tobiasz M (2001) Biocontrol effects against Pythium root rot of seedling geraniums, 2001. Biol Cult Tests 17:O09 Daughtrey ML, Harlan B, Linderman S, Hausbeck MK (2014) Coleus cultivars and downy mildew. Special research report #136. American Floral Endowment, Disease Management. http://endowment.org/wp-content/uploads/2013/03/136-ColeusDM-Cv-2014.pdf. Accessed 7 Oct 2016 Daughtrey ML, Holcomb GE, Eshenaur B, Palm ME, Belbahri L, Lefort F (2006) First report of downy mildew on greenhouse and landscape coleus caused by a Peronospora sp. in Louisiana and New York. Plant Dis 90:1111 Del Castillo Munera J, Hausbeck MK (2016) Characterization of Pythium species associated with greenhouse floriculture crops in Michigan. Plant Dis 100:569–576 Denton GJ, Beal E, Denton JO, Clover G (2015) First record of downy mildew, caused by Peronospora belbahrii, on Solenostemon scutellarioides in the UK. New Dis Rep 31:14 Fonseca MEN, Boiteaux LS, Singh RP, Kitajima EW (1989) A small viroid in Coleus species from Brazil. Fitopatol Bras 14:94–96 Fu FH, Li SF, Jiang DM, Wang HQ, Liu AQ, Sang LW (2011) First report of Coleus blumei viroid 2 from commercial coleus in China. Plant Dis 95:494 Garzon CD, Molineros JE, Yanez JM, Flores FJ, Del Mar Jimenez-Gasco M, Moorman GW (2011) Sublethal doses of mefenoxam enhance Pythium damping-off of geranium. Plant Dis 85:1233–1238 Gutierrez WA, Shew HD, Melton TA (2001) Rhizoctonia diseases in tobacco greenhouses. North Carolina State University Plant Pathology Extension TB07, Tobacco Disease Note 7. http:// www.ces.ncsu.edu/depts/pp/notes/Tobacco/tdin007/tb07.html. Accessed 7 Oct 2016 Harlan BR, Hausbeck MK (2010) Evaluation of foliar sprays and soil drenches of fungicides for the control of downy mildew of coleus, 2009. Plant Dis Manag Rep 4:OT018 Harlan BR, Hausbeck MK (2011) Is coleus downy mildew here to stay? Michigan State University Extension News for Agriculture-Floriculture, Feb 8. http://msue.anr.msu.edu/news/is_coleus_ downy_mildew_here_to_stay. Accessed 7 Oct 2016 Harlan BR, Hausbeck MK (2013) Understanding coleus downy mildew. Special research report #134. American Floral Endowment, Disease Management. http://endowment.org/wp-content/ uploads/2014/03/134ColeusDMDisease2013.pdf. Accessed 7 Oct 2016 Harlan BR, Granke L, Hausbeck MK (2012) Epidemiology and management of downy mildew, a new pathogen of coleus in the United States. Acta Hortic 952:813–818 Harlan BR, Hausbeck MK (2016) Epidemiology and management of impatiens downy mildew in the United States. Acta Hortic 952:813–818 Hausbeck MK, Harlan BR (2010) Evaluation of registered and unregistered fungicides for control of Botrytis blight of geranium, 2009. Plant Dis Manag Rep 4:OT011 Hausbeck MK, Harlan BR (2013) Pythium root rot in the greenhouse. Michigan State University Extension News for Agriculture-Floriculture, Oct 7. http://msue.anr.msu.edu/news/pythium_ root_rot_in_the_greenhouse. Accessed 7 Oct 2016 Hausbeck MK, Harlan BR (2015) Greenhouse disease control update. In: Greenhouse session summaries, proceedings of the 2015 Michigan Greenhouse Expo, Grand Rapids, pp 1–7.

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http://glexpo.com/summaries/2015summaries/Greenhouse_DiseaseControl.pdf. Accessed 7 Oct 2016 Hausbeck MK, Moorman GW (1996) Managing Botrytis in greenhouse-grown flower crops. Plant Dis 80:1212–1219 Hausbeck MK, Quackenbush W, Linderman SD (2002) Evaluation of biopesticide for control of Botrytis blight of geranium, 2002. Fungicide Nematicide Tests 58:OT028 Holcomb GE, Valverde RA (1991) Identification of a virus causing a mosaic on coleus. Plant Dis 75:1183–1185 Hou WY, Li SF, Wu ZJ, Jiang DM, Sano T (2009) Coleus blumei viroid 6: a new tentative member of the genus Coleviroid derived from natural genome shuffling. Arch Virol 154:993–997 Ito Y, Takeuchi T, Matsushita Y, Chikuo Y, Satou M (2015) Downy mildew of coleus caused by Peronospora belbahrii in Japan. J Gen Plant Pathol 81:328–330. http://link.springer.com/ article/10.1007/s10327-015-0601-3. Accessed 25 Mar 2016 Ivors K, Lacy LW, Milks DC (2011) Evaluation of fungicides for the control of downy mildew on coleus, 2010. Plant Dis Manag Rep 5:OT019 Jarvis WR (1980) Epidemiolgy. In: Coley-Smith JR, Verhoeff K, Jarvis WR (eds) The biology of Botrytis. Academic, London, pp 219–250 Jiang DM, Li SF, Fu FH, Wu ZJ, Xie LH (2013) First report of Coleus blumei viroid 5 from Coleus blumei in India and Indonesia. Plant Dis 97:561 Jiang D, Rui Gao R, Qin LV, Wu Z, Xie L, Hou W, Li S (2014) Infectious cDNA clones of four viroids in Coleus blumei and molecular characterization of their progeny. Virus Res 180:97–101 Koike ST, Wilen CA (2009) Pythium root rot – UC IPM pest management guidelines: floriculture and ornamental nurseries. UC ANR publication 3392. http://www.ipm.ucdavis.edu/PMG/ r280100211.html. Accessed 7 Oct 2016 Lambert L (2005) Virus: plantes sensibles. Reseau D’Avretissements Phytosanitaires Bulletin d’information Cultures En Serres No 2. https://www.agrireseau.net/horticulture-serre/docu ments/68143. Accessed 7 Oct 2016 Lockhart BEL, Mason SL, Johnson DA, Mollov DS (2010) First report of tobacco etch virus infection in coleus in the United States. Plant Dis 94(7):921 Mollov DS, Hayslett MC, Eichstaedt KA, Beckman NG, Daughtrey ML, Lockhart BE (2007) Identification and characterization of a carlavirus causing veinal necrosis of coleus. Plant Dis 91:754–757 Moorman GW, Lease RJ (1992) Benzimidazole- and dicarboximide-resistant Botrytis cinerea from Pennsylvania greenhouses. Plant Dis 76:477–480 Moorman GW, Lease RJ (1999) Control of Rhizoctonia root rot of coleus, 1999. Fungicide Nematicide Tests 56:OT5 Neergaard P (1977) Seed pathology, vol 1. MacMillan, London, p 839 Palmateer AJ, Harmon PF, Schubert TS (2007) Downy mildew of coleus (Solenostemon scutellarioides) caused by Peronospora sp. in Florida. New Dis Rep 16:11 Parris GK (1959) A revised host index of Mississippi plant diseases. Mississippi State University, Botany Department Miscellaneous Publication 1, pp 146 Pedley R, Pedley K (1974) Coleus: a guide to cultivation and identification. Bartholomew and Son, Edinburgh, p 116 Pinckard JA (1942) The mechanism of spore dispersal in Peronospora tabacina and certain other downy mildew fungi. Phytopathology 32:505–511 Pscheidt JW, Ocamb CM (2016) Pacific northwest plant disease management handbook. http:// pnwhandbooks.org/plantdisease/. Accessed 7 Oct 2016 Rivera Y, Salgado-Salazar C, Windham AS, Crouch JA (2016) Downy mildew on coleus (Plectranthus scutellarioides) caused by Peronospora belbahrii sensu lato in Tennessee. Plant Dis. doi:10.1094/PDIS-10-15-1120-PDN Shaw CG (1981) Taxonomy and evolution. In: Spencer DM (ed) The Downy mildews. Academic, San Francisco, pp 17–29

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Sikora EJ (2004) Tobacco etch virus. Alabama cooperative extension ANR-869. http://www.aces. edu/pubs/docs/A/ANR-0869/ANR-0869.pdf. Accessed 7 Oct 2016 Spieker RL, Haas B, Charng Y-C, Freimuller K, Sanger HL (1990) Primary and secondary structure of a new viroid ‘species’ (CbVd1) present in the Coleus blumei cultivar ‘Bienvenue.’. Nucleic Acids Res 18:3998 Tsushima T, Sano T (2015) First report of Coleus blumei viroid 5 infection in vegetatively propagated clonal coleus cv. ‘Aurora Black Cherry’ in Japan. New Dis Rep 32:7 Warfield CY, Sugar JC, Sugar KJ (2008) Evaluation of fungicides for the control of downy mildew on coleus, 2007. Plant Dis Manag Rep 2:OT004 Webster BJ, Hausbeck MK (2004) Evaluation of reduced risk fungicides and biopesticides for control of Botrytis blight of geranium, 2004. Fungicide Nematicide Tests 60:OT009 Westcott C (1979) Westcott’s plant disease handbook, 4th edn. Van Nostrand Reinhold Company, New York, p 112 Wright J (2013) Greenhouse diseases 101: Rhizoctonia. Greenhouse grower. http://www.greenhou segrower.com/production/crop-inputs/greenhouse-diseases-101-rhizoctonia/. Accessed 2 Feb 2016

Diseases of Gardenia A. J. Palmateer and A. R. Chase

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-Like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Gardenia Canker (Diaporthe gardeniae = Phomopsis gardeniae) . . . . . . . . . . . . . . . . . . . 2.2 Myrothecium Leaf Spot (Myrothecium roridum) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Powdery Mildew (Erysiphe polygoni, Oidium sp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Rhizoctonia Root Rot (Rhizoctonia solani, Rhizoctonia sp.) . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Phytophthora Root and Stem Rot (Phytophthora cinnamomi and P. nicotianae) . . . . 2.6 Pythium Root Rot (Pythium splendens, P. spinosum) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Bacterial and Phytoplasma Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Bacterial Leaf Spot (Xanthomonas maculifoliigardeniae) . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Abiotic Disorders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Iron Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Bud Drop . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2 2 2 4 5 6 7 8 10 10 12 12 12 13

Abstract

Gardenias are a very important crop for both potted flowers for the florist trade and the landscape industries. Gardenias are susceptible to root rot, and the most common and damaging pathogens include Phytophthora and Rhizoctonia. One of the most devastating diseases affecting gardenias is canker caused by Diaporthe gardeniae, which can lead to unsightly cankers and galls on the lower stem causing plant decline and eventual death. Several leaf spot pathogens A.J. Palmateer (*) Department of Plant Pathology, University of Florida, Tropical Research and Education Center, Homestead, FL, USA e-mail: ajp@ufl.edu A.R. Chase Chase Agricultural Consulting LLC, Cottonwood, AZ, USA e-mail: [email protected] # Springer International Publishing AG 2016 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florist's Crops Diseases, Handbook of Plant Disease Management, DOI 10.1007/978-3-319-32374-9_33-1

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are reported on gardenia with Myrothecium and Xanthomonas being among the most common. Disease management practices for gardenia depend on good sanitation, judicious use of pesticides, and cultural practices. Keywords

Gardenia • Phytophthora • Myrothecium • Pythium • Rhizoctonia • Gardenia canker • Xanthomonas

1

Introduction

Gardenia jasminoides is a popular flowering shrub in the Rubiaceae family that can be found growing in tropical and semitropical areas in the landscape worldwide (Fig. 1). It is also grown as a potted flowering crop by the florist industry in temperate climates using protected culture. Patio gardenias are normally not grafted and cuttings are grown in pots. Some cultivars are also grown as standards where the apical meristem is pruned to form a head. Landscape plantings are mostly grafted to the root stock Gardenia thunbergia for resistance to root-knot nematodes and Rhizoctonia spp.

2

Fungal and Fungus-Like Diseases

2.1

Gardenia Canker (Diaporthe gardeniae = Phomopsis gardeniae)

Geographic occurrence and impact. This disease has been reported within the continental USA; gardenia canker has been reported in California (Hansen and Barrett 1938), Washington, Nebraska, Kansas, Ohio, Massachusetts, and Florida, and has also been reported in Europe (Italy) (Alfieri 1967) and South America. Symptoms/signs. Initial symptoms of gardenia canker include wilting, yellowing, shriveling, and defoliation of leaves and frequent premature bud drop of unopened flowers. The wilting can affect a portion of the plant or the entire plant where it can appear as a sudden wilt. The cankers are at first small, circular to oblong, swollen brownish lesions on the main stem and branches. Cankers can become as large as 5 cm (2 in.) in diameter and cause partial to complete girdling of stems and branches. Even partial girdling can result in stunting and wilt. The diseased area decays and begins to separate from trunk or branch tissue exposing the wood. In some cases, the periphery of the canker becomes calloused and appears rough and corrugated. The cankers become overgrown with corky callous tissue extending longitudinally in both directions, increasing in size and forming galls. These galls typically increase two to three times in size and produce deep longitudinal cracks. Branches in close proximity to galls begin to lose vigor compared to those further away or originating from unaffected stem tissue. Stem cankers are more common at

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Fig. 1 Healthy Gardenia jasminoides with showy white blooms

the base of the plant affecting the crown, but they can occur anywhere on the plant especially where wounded from mechanical injury. The foliage of older infected plants appears dull eventually turning yellow or drying out and becoming brown before defoliating. Flower buds will abscise before opening. Disease severity and wilting is usually more pronounced under cooler temperatures. During warmer conditions diseased plants may continue to live, but remain stunted and appear unhealthy. Sometimes signs of the imperfect stage of the fungus (Phomopsis gardeniae) are present consisting of black fruiting bodies on the lesions (pycnidia) that contain both alpha and beta conidia. Biology and epidemiology. The fungal pathogen is widely considered to be a wound parasite; however, once infection occurs and the fungus becomes established in host tissue, disease develops and progresses forming mature cankers and subsequent pycnidia. Under conditions of high relative humidity and warm temperatures, an abundance of spores forms within the partially submerged pycnidia and is easily spread by windblown rain and splash dispersion in irrigation water. The fungus is most prevalent on or near cankers and overwinters from one season to the next on the diseased portions of infected plants. The fungus is notorious for entering gardenias through leaf joints at the base of cuttings. Thus, freshly cut leaf bases act as an excellent point of entry for the fungus. Management • Cultural practices – Disease-free stock plants should be used for propagation. Pathogen elimination through sanitation by steam-pasteurizing rooting media, sand, and peat is highly recommended. All cuttings should be taken with sharp blades avoiding rough and jagged edges where spores of the pathogen are more likely to gain entry. All tools contacting plant material should be properly disinfected to minimize contamination and potential spread of the pathogen. Plant injury due to careless cultural practices should be avoided. The productivity

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of affected plants has been shown to be enhanced or lengthened by piling soil up around the stem base covering basal cankers to encourage new root formation above the diseased portions of stems. • Fungicides and biological controls – Preventative applications of fungicides containing chlorothalonil (FRAC M5), mancozeb (FRAC M3), propiconazole (FRAC 3), or thiophanate methyl (FRAC 1) ensuring excellent coverage of all plant parts may minimize disease outbreaks. In addition, the combination of pyraclostrobin (FRAC 11) and boscalid (FRAC 7) has been effective on similar pathogens on other crops. It is important to rotate fungicides from different classes in order to help prevent the development of resistance. Scientific literature reports good efficacy against Phomopsis on other plant species using biological control products containing Gliocladium virens. These products were applied as a soil drench and as a wound treatment.

2.2

Myrothecium Leaf Spot (Myrothecium roridum)

Geographic occurrence and impact. This disease was found and described from Pennsylvania in 1941 (Fergus 1957) and has since been found in California (Barrett and Hardman 1947) and Florida. Symptoms/signs. The leaf spots are mostly circular and vary in size up to 2 cm (~1 in.) in diameter (Fergus 1957). Initial spots appear water soaked and turn brown with age. Often the centers of the lesions fall out giving a shot hole appearance. Under wet conditions, the spots coalesce and large portions of the leaves can rot away. Characteristic greenish-black pillow-shaped sporodochia with white setal margins form on necrotic tissue and can be seen on both the upper and lower leaf surface (Fig. 2). Biology and epidemiology. Myrothecium leaf spot on gardenia is most common under humid conditions such as those found when attempting to root cuttings. Pathogenicity of M. roridum isolates is reported to vary greatly with most isolates being wound pathogens on gardenia. Management • Cultural practices – Monitor nutritional programs closely, because overfertilization has been shown to increase leaf spots and rots caused by Myrothecium in some tropical foliage plants. Imbalanced fertilization programs, damage from pesticides, heat or cold injury, and leaf necrosis caused by water stress should be avoided. Wounded or weakened tissue allows Myrothecium to form a colony in the canopy, which serves as the focal point for disease spread. • Fungicides and biological control – Apply fungicides containing azoxystrobin or pyraclostrobin (FRAC 11), chlorothalonil (FRAC M5), fludioxonil (FRAC 12), iprodione (FRAC 2), or mancozeb (FRAC M3). Good coverage is very important as this fungus can colonize both sides of the leaf surface. No effective biocontrol agents have been reported for this disease.

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Fig. 2 Myrothecium roridum on gardenia leaves

2.3

Powdery Mildew (Erysiphe polygoni, Oidium sp.)

Geographic occurrence and impact. This disease has been reported in Texas and tropical America but is not common. Symptoms/signs. Powdery mildew of gardenia primarily affects young leaves and shoots where symptoms of infection include leaf yellowing, slight deformation of leaf and bud tissue, and leaf drop. Powdery mildew fungi typically grow on the surface of plants and infect plant cells using a haustorium to absorb nutrients. Signs of infection include a white to gray colored fungal growth that appears like powder and is most often found on the upper leaf surface but may occur on the underside of leaves when conditions are highly favorable. Biology and epidemiology. Powdery mildew fungi can survive in leaf litter. High relative humidity and moderate to warm temperatures favor outbreaks of powdery mildew. Shade especially during periods of high relative humidity and moderate temperatures are most favorable conditions for powdery mildew. Management • Cultural practices – Proper location of plants in full to partial sun and avoiding heavily shaded areas will help to prevent outbreaks of powdery mildew. Space plants apart to allow for good air movement through the plant canopy. Overhead irrigation may actually reduce the spread of powdery mildew, because the water, unlike with other fungi, actually inhibits spore germination. Closely monitor plants and examine new shoot growth for signs of powdery mildew. • Fungicides and biological controls – The use of fungicides may be necessary in production situations when favorable conditions persist. Horticultural oil or plantbased oil such as neem oil can be used to control powdery mildew. Be certain not to apply oil within 14 days of a sulfur spray or plant injury may occur. Never

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Fig. 3 Darkly stained stem cankers on gardenia in propagation due to Rhizoctonia solani

apply oil when ambient temperatures exceed (30  C/86 F) or when plants are water-stressed. Sulfur-based products are commonly used for controlling powdery mildew and should be used only as a preventative before signs of powdery mildew appear. Fungicides with efficacy for controlling powdery mildew include thiophanate methyl (FRAC 1); copper (FRAC M1); myclobutanil, propiconazole, triadimefon (FRAC 3); azoxystrobin, pyraclostrobin, trifloxystrobin (FRAC 11); chlorothalonil (FRAC M5); and Bacillus spp. (FRAC 44).

2.4

Rhizoctonia Root Rot (Rhizoctonia solani, Rhizoctonia sp.)

Geographic occurrence and impact. This disease has been reported wherever gardenia is grown and is considered to be one of the most common and economically important diseases affecting gardenia. Specific reports from the USA (California, Florida, New Jersey), Greece, and Japan have been made. Symptoms/signs. Common symptoms in cutting beds include damping-off, hypocotyl rot, root rot, stunting, yellowing, and death. On older plants, Rhizoctonia causes the roots to turn light to dark brown and macerates the tissue; but the pathogen has been reported to affect all portions of the plant. Light to dark brown irregularly shaped spots form anywhere on the stem and foliage during propagation (Fig. 3) or in the landscape. Mycelium of the fungal pathogen is frequently present and looks like a light brown web colonizing affected tissue giving rise to the common name web blight. Biology and epidemiology. Rhizoctonia can survive in soil and plant debris as mycelium and sclerotia that can be stimulated to germinate by plant exudates. Rhizoctonia stem rot and aerial blight or leaf spot occurs in the warmer months especially during the rainy season when there is an abundance of moisture. Such favorable conditions can occur at anytime throughout the year when plants are produced in covered structures that employ overhead irrigation. Under favorable conditions severe outbreaks are common and can occur in as little 7 days.

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Management • Cultural practices – Prevention is the most important step for managing Rhizoctonia. Start with high quality, healthy cuttings, use new or sterilized pots and new potting media. Placing plants on hard surfaces, ground clothes, or benches so that they are not in direct contact with soil is crucial. Due to the quick onset of this disease it is recommended to scout plants weekly for early symptom detection. • Fungicides and biological controls – Specific trials on fungicides and biocontrol agents have not been reported on gardenia for Rhizoctonia. On other woody ornamental crops, strobilurins (FRAC 11); fludioxonil (FRAC 12); and thiophanate methyl (FRAC 1) are among the most effective fungicides. Additionally, the biocontrol agent Trichoderma harzianum T22 can be very effective against root rot but less so against stem rot or cutting rot.

2.5

Phytophthora Root and Stem Rot (Phytophthora cinnamomi and P. nicotianae)

Geographic occurrence and impact. Phytophthora cinnamomi is typically the most common species associated with container and field grown gardenias in the Southeastern USA, but other species have been widely reported to cause root and stem rot of gardenia. P. nicotianae has been reported from the USA in North Carolina (Olson and Benson 2011), California, and Florida and also Greece. Symptoms/signs. Phytophthora usually reduces the overall volume of roots that are present, thus greatly inhibiting the plant’s ability to take up water and nutrients for proper growth. The roots of diseased plants appear brittle and brown to reddish brown in color (Fig. 4a). The pathogen usually colonizes the crown often girdling the stem at or just above the soil line. A brown to reddish brown discoloration of the tissues occurs just below the bark layer and may extend up the stem above the soil. Affected tissue is most often wet in appearance. Aboveground symptoms can easily be confused with those caused by a nutritional disorder, overwatering, drought stress, and other abiotic disorders. Slight yellowing of the leaves followed by wilting and possibly plant death (Fig. 4b) are common symptoms associated with Phytophthora. Landscape or field grown gardenias may show symptoms of general decline for up to 1 year or more before succumbing to root rot caused by Phytophthora. Biology and epidemiology. Phytophthora species are fungus-like water molds or Oomycetes that most often survive as resting structures consisting of chlamydospores, oospores, and mycelia in diseased roots, crowns, and other plant debris. These structures are released into the soil or potting medium from infested crop debris and can be easily spread throughout propagation and container production areas in windblown plant and potting material and by splashing rain and irrigation water. Heaviest losses occur in poorly drained areas where there is standing water. Overwatering in the nursery and landscape are contributing factors to losses from Phytophthora.

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Fig. 4 (a) Brittle and brown diseased roots due to infection by Phytophthora nicotianae. (b) Necrotic and wilted leaves on gardenia with stem infection from Phytophthora nicotianae

Management • Cultural practices – An integrated approach is necessary for success in controlling Phytophthora root rot. Prevention and sanitation are key, because once symptoms appear it is often too late to adequately control the disease. Providing for proper plant establishment and following good horticultural practices including adequate nutrition and irrigation will help to reduce potential losses from this disease. Recycling irrigation water does not appear to be a significant source of disease in gardenias. • Fungicides and biological controls – Fungicides should be used as a preventative measure, and those used for managing Phytophthora include fosetylaluminum, phosphorous acid (FRAC 33); fluopicolide (FRAC 43); mefenoxam (FRAC 4); dimethomorph, mandipropamid (FRAC 40); chlorothalonil (FRAC M5); azoxystrobin, pyraclostrobin, fenamidone (FRAC 11); etridiazole (FRAC 14); propamocarb hydrochloride (FRAC 28); cyazofamid (FRAC 21); oxathiapiprolin (FRAC U5); and Bacillus spp. (FRAC 44).

2.6

Pythium Root Rot (Pythium splendens, P. spinosum)

Geographic occurrence and impact. Pythium root rot is common in most gardenia production areas although losses are usually minimal and damage is not as severe when compared to that caused by Phytophthora. Symptoms/signs. The aboveground symptoms often include yellowing of the oldest leaves first followed by defoliation and in severe cases wilting (Fig. 5a). The plants typically appear unhealthy for an extended period of time and may not respond to fertilization due to Pythium attacking the feeder roots and inhibiting the plant’s ability to take up water and nutrients. Healthy roots are white whereas diseased roots appear brown and discolored, or may be completely missing (Fig. 5b). The outer cortical tissue can be easily removed when pulling on the roots (Fig. 5c), leaving behind the threadlike core or stele of the water conducting tissue.

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Fig. 5 (a) Pythium root rot exhibiting severe leaf chlorosis and wilting caused by P. splendens. (b) Pythium root rot on gardenia showing typical sparse, necrotic, and discolored roots. (c) Outer cortex of roots infected with Pythium spp. often deteriorates leaving the light colored central core

Biology and epidemiology. The biology and epidemiology is very similar to Phytophthora. Disease development is highly favored by any factor that encourages wet soil conditions, including poor drainage and over watering. Planting gardenias too deep also contributes to the disease. Management • Cultural practices – An integrated approach is necessary for success in controlling Pythium root rot. Closely monitoring irrigation and using a potting medium that allows for adequate drainage or aeration is key. Providing for proper plant establishment and following good horticultural practices including adequate nutrition and irrigation will help to reduce potential losses from this disease. Recycling irrigation water does not appear to be a significant source of disease in gardenias. • Fungicides and biological controls – Fungicides should be used as a preventative measure and those used for managing Pythium include fosetyl-aluminum,

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phosphorous acid (FRAC 33); fluopicolide (FRAC 43); mefenoxam (FRAC 4); dimethomorph, mandipropamid (FRAC 40); chlorothalonil (FRAC M5); azoxystrobin, pyraclostrobin, fenamidone (FRAC 11); etridiazole (FRAC 14); propamocarb (FRAC 28); cyazofamid (FRAC 21); oxathiapiprolin (FRAC U5); and Bacillus spp. (FRAC 44).

3

Bacterial and Phytoplasma Diseases

3.1

Bacterial Leaf Spot (Xanthomonas maculifoliigardeniae)

Geographic occurrence and impact. This disease was reported for the first time in California in 1941 (Burkholder and Pirone 1941) on greenhouse grown gardenias. It was later described by Ark and Barrett in 1946. It is commonly found in production and the landscape in Florida and California. Symptoms/signs. Very young tender leaves are the first to develop symptoms which consist of small yellow spots. The yellow spots are angular to circular and gradually increase in size where the center initially appears water soaked. As the leaves age, the center portion of the spots turns reddish brown and necrotic. Lesions are most often surrounded by a wide chlorotic halo (Fig. 6). Spots eventually coalesce and produce larger necrotic areas. Premature leaf drop is common on severely affected plants. Biology and epidemiology. Conditions providing high relative humidity and temperature, especially in greenhouse production where gardenias are induced to bloom, favor disease development. Symptoms may appear rather suddenly on gardenias growing outdoors during warm and wet periods. Management • Cultural practices – Overhead irrigation should be avoided or timed for periods when leaves dry quickly as bacterial cells are readily dispersed in irrigation water increasing the incidence of diseased plants and plant parts. The disease is most severe during periods of prolonged leaf wetness. • Bactericides and biological controls – As a preventative measure apply a bactericide containing copper hydroxide, copper oxychloride, copper sulfate, or other forms (FRAC M1); mancozeb (FRAC M3); or streptomycin sulfate (FRAC 25). The quaternary ammonium (QA) product DDAC also shows good control of bacterial diseases on other ornamentals. Possible biological control agents include species of Bacillus such as B. amyloliquifaciens and B. subtilis which have both shown good efficacy against related bacterial pathogens on other ornamentals and agronomic crops. These are less common diseases that are known to affect gardenia: Botrytis cinerea – California, Florida Hawaii, US Virgin Islands, Puerto Rico, Greece, Saudi Arabia, and China

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Fig. 6 Xanthomonas leaf spot on gardenia caused by Xanthomonas maculifoliigardeniae

Capnodium spp. (sooty mold) – California, Georgia, US Gulf Coast States, and Venezuela Cercospora (and Pseudocercospora) – Indonesia, Venezuela, and Florida Colletotrichum spp. (anthracnose) – Hong Kong, Cina, India, and Florida Pestalotia (and Pestalotiopsis) – Alabama, Florida, Cuba, Japan, and China Phyllosticta gardeniae – Japan, Republic of Georgia, India, and the USA (Michigan, North Carolina, New Jersey, and Texas) Phytoplasma – China (Sun and Zhao 2012) Tomato spotted wilt virus – USA

4

Abiotic Disorders

4.1

Iron Deficiency

Geographic occurrence and impact. Gardenias are an acid loving plant, so when soil pH is greater than 6.0 the potential for micronutrient deficiencies increase. The most common nutrient deficiency affecting gardenias is iron, and when soil pH is above 7 the iron may be in a form that is not readily plant-available.

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Fig. 7 Iron deficiency shows stunted chlorotic leaves when plants are subjected to high pH

Symptoms/signs. Most often iron deficient gardenias are stunted and have pale green to yellow leaves. Symptoms are most obvious in younger leaves which turn completely yellow except for dark green veins. Older leaves may only turn yellow along the edges (Fig. 7). Management • Cultural practices – Anything that can be done to acidify (lower) the pH of the soil will help. Some relevant soil amendments include aluminum sulfate, iron sulfate, or wettable sulfur. Gardenias growing in soils with a high buffering capacity such as calcareous soils require foliar applications of iron chelate or ferrous sulfate (FeSO4.2H20) can be very effective for correcting iron deficiency in gardenias. Allow the soil to remain evenly moist but not completed saturated.

4.2

Bud Drop

Geographic occurrence and impact. Bud drop is a prevailing issue with gardenias and is especially common after a change in location or growing conditions. It can occur in both container and landscape gardenias. Symptoms/signs. When gardenias are under stress unopened flower buds may prematurely drop off from the plant (Fig. 8). Numerous things can cause stress, but some of the most common include insects feeding, diseases including root feeding nematodes, inadequate fertilization, irrigation and light, and temperature fluctuations such as unusually cold or hot conditions. Management • Cultural practices – Anything that can be done to minimize plant stress. Gardenias do not respond well to changes in growing conditions, so avoid moving or

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Fig. 8 Irregular watering or temperature fluctuations causes premature bud abortion and drop

relocating plants that are thriving. Monitor irrigation so that the soil remains evenly moist, but not saturated.

References Alfieri SA Jr (1967) Gardenia canker, vol 54, Division of plant industry circular. Florida Department of Agriculture, Gainesville Ark PA, Barrett JT (1946) A new bacterial leaf spot of greenhouse-grown gardenias. Phytopathology 36:865–868 Barrett JT, Hardman DA (1947) Myrothecium leaf spot and canker of Gardenia. Phytopathology 37:360 Burkholder WH, Pirone PP (1941) Bacterial leaf spot of gardenia. Phytopathology 31:192–194 Fergus CL (1957) Myrothecium roridum on Gardenia. Mycologia 49:124–127 Hansen HN, Barrett JT (1938) Gardenia canker. Mycologia 30:15–19 Olson HA, Benson DM (2011) Characterization of Phytophthora spp. on floriculture crops in North Carolina. Plant Dis 95:1013–1020 Sun XC, Zhao WJ (2012) First report of a group 16Sri phytoplasma associated with Gardenia jasminoides in China. Plant Dis 96:1576

Diseases of Geranium Cristina Rosa and Gary W. Moorman

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Alternaria Leaf Spot (Alternaria Sp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Black Root Rot (Thielaviopsis basicola (Berk. And Br) Ferris) . . . . . . . . . . . . . . . . . . . . . . 2.3 Cercospora Leaf Spot (Cercospora Sp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Gray Mold (Botrytis Blight; Botrytis cinerea Pers.: Fr.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Pythium Root Rot and Blackleg (Pythium Sp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 Rhizoctonia Root and Crown Rot (Thanatephorus cucumeris (Rhizoctonia solani Kuhn) Rhizoctonia Sp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7 Rust (Puccinia pelargonii-zonalis Doidge, Puccinia granularis, Puccinia morrisoni) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.8 Sclerotinia Crown Rot or Cottony Stem Rot (Sclerotinia sclerotiorum (Lib.) de Bary) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.9 Verticillium Wilt (Verticillium albo-atrum Reinke and Berthe or V. dahliae Kleb.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Bacterial and Phytoplasma Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Bacterial Blight (Xanthomonas hortorum Pv. pelargonii; Formerly, Xanthomonas campestris Pv. pelargonii) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Bacterial Fasciation (Rhodococcus fascians, Formerly Known as Corynebacterium fascians, Bacterium fascians, and Phytomonas fascians) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Bacterial Wilt or Southern Wilt (Ralstonia solanacearum, Formerly Known as Burkholderia solanacearum, and Pseudomonas solanacearum) . . . . . . . . . . . . . . . . . . . 3.4 Crown Gall (Rhizobium tumefaciens, Formerly Agrobacterium tumefaciens) . . . . . . . . 3.5 Pseudomonas Leaf Spot (Bacterial Leaf Spot; Pseudomonas cichorii) . . . . . . . . . . . . . . . 3.6 “Candidatus Phytoplasma Asteris” . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Viral Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Arabis Mosaic Virus (ArMV), Tobacco Ringspot Virus (TRSV), Tomato Ringspot Virus (ToRSV), and Artichoke Italian Latent Virus (AILV) . . . . . . . . . . . . . . . . . . . . . . . . . . .

2 3 3 4 5 5 7 9 9 10 11 13 13 15 16 18 19 20 20 20

C. Rosa (*) • G.W. Moorman Department of Plant Pathology & Environmental Microbiology, The Pennsylvania State University, University Park, PA, USA e-mail: [email protected]; [email protected] # Springer International Publishing AG 2016 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florist's Crops Diseases, Handbook of Plant Disease Management, DOI 10.1007/978-3-319-32374-9_34-1

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C. Rosa and G.W. Moorman 4.2 4.3 4.4 4.5 4.6

Beet Curly Top Virus (BCTV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Impatiens Necrotic Spot Virus (INSV) and Tomato Spotted Wilt Virus (TSWV) . . . . . Pelargonium Zonate Spot Virus (PZSV), Cucumber Mosaic Virus (CMV) . . . . . . . . . . Tobacco Mosaic Virus (TMV) and Tobacco Rattle Virus (TRV) . . . . . . . . . . . . . . . . . . . . . . Tomato Bushy Stunt Virus (TBSV), Tobacco Necrosis Virus (TNV), Moroccan Pepper Virus (MPV), and Pelargonium Leaf Curl Virus (PLCV) . . . . . . . . . . . . . . . . . . . . 4.7 Pelargonium Flower Break Virus (PFBV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Nematode Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 Abiotic Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1 Edema . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2 Heat Stress . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract

Geraniums are susceptible to a wide variety of fungi, bacteria, and viruses as well as nematodes and abiotic diseases. The systemic nature of some of the pathogens makes it likely that the diseases they cause can be found wherever vegetatively propagated geraniums are shipped. Management of these pathogens is paramount for specialty propagators, while growers purchasing plants must inspect incoming plants for symptoms and understand the biology of the pathogens involved in order to manage them effectively. Keywords

Botrytis cinerea • Pythium Spp. • Ralstonia solanacearum • Xanthomonas campestris • Virus • Edema • Culture Indexing • Virus Indexing • Certification

1

Introduction

Species of Geranium (described by Linnaeus) are often grown as herbaceous perennials outdoors, while Pelargonium (described by L’Héritier) are generally grown indoors for long periods of time or as annuals or perennials outdoors. Emphasis in this chapter is placed on the pathogens and diseases of Pelargonium graveolens, P. peltatum, and the complex hybrids Pelargonium X domesticum and Pelargonium X hortorum. Common names for these Pelargonium include florists, garden, bedding, zonal, ivy, horseshoe, rose, and scented and regal geranium, among others depending upon the species. It is thought that Pelargonium was brought into cultivation in Europe from plants initially obtained in South Africa in the early 1600s and are well documented as having been grown in many European gardens by the mid-1700s (Laughner 1993). At one time, Pelargonium was the mainstay of the bedding plant industry, and the majority of plants were propagated vegetatively. While vegetatively propagated varieties are still grown extensively, many other varieties are grown from seed. Disease susceptibility varies greatly with the variety and the pathogen involved, and some pathogens pose more of a problem in greenhouse production than when plants are grown outdoors. The geographic distribution

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of particular pathogens reported in the literature may provide clues to the environmental conditions that favor the disease, but the worldwide trade in millions of Pelargonium cuttings greatly increases the potential for any pathogen to be found anywhere Pelargonium is grown. Crucial to the production of Pelargonium varieties through vegetative propagation is the establishment of stock plants that are free of the plant pathogens that infect systemically or stay closely associated with plant tissues including Verticillium, Xanthomonas campestris pv. pelargonii, Ralstonia solanacearum, Agrobacterium tumefaciens, Rhodococcus fascians, and all the viruses noted here. Culture indexing is used to detect those plant pathogens that readily grow in broth containing required nutrients, and virus indexing employs various methods of testing plant tissues for the presences of specific viruses (De Boer et al. 1996; Huttinga 1996; Oglevee-O’Donovan 1993). Plants free of selected pathogens are then maintained as stock plants from which numerous cuttings are taken. This “culture/virus indexing” procedure has been central in the production of “pathogen-free” Pelargonium, but note that they are “free” of selected pathogens. Other pathogens may be present in or on the plants.

2

Fungal and Fungus-like Diseases

For a list of pathogens of this plant in the USA, see Farr, et al. (1989). For the most up-to-date listing, search http://nt.ars-grin.gov/fungaldatabases/fungushost/ fungushost.cfm.

2.1

Alternaria Leaf Spot (Alternaria Sp.)

Geographic occurrence and impact. This disease occurs when conditions are not well suited to geranium growth, particularly if the temperatures are too high or low or when other factors favor plant senescence. Generally, economic losses are minor (Engelhard 1993; Munnecke 1956). Symptoms/signs. Water-soaked spots initially formed on the underside of the leaf enlarge to 5–10 mm (1/4–1/2 in.). Leaves may have numerous spots, and some spots may have a yellow halo. Concentric dark rings form in the spots creating a target-like pattern where the fungus has formed dark brown, club-shaped, multi-celled spores on the surface of the leaf (Fig. 1). The spots may merge and occupy most of the leaf. Diagnosis is facilitated if infected leaves are placed in a container with moisture, and the spores are allowed to develop (Engelhard 1993). Biology and epidemiology. Warm, wet conditions greatly favor infection and development of symptoms particularly on older leaves. The fungus survives on dead leaves on the soil surface, and spores are dispersed by air currents. The disease generally begins on older leaves, particularly if they are senescing, and progresses upward on the plant. Management. Irrigate plants in a manner that keeps water off the foliage. Ensure good air circulation among plants so that the humidity within the canopy is kept low.

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Fig. 1 Alternaria leaf spot (Photo courtesy of The Pennsylvania State University Dept. of Plant Pathology and Environmental Microbiology)

Remove and destroy infected leaves and any crop debris in pots, or on benches, and walkways. Apply a fungicide to protect plants.

2.2

Black Root Rot (Thielaviopsis basicola (Berk. And Br) Ferris)

Geographic occurrence and impact. This disease can occur in Pelargonium and Geranium production (Linderman 1993). In Pelargonium, seed-type geraniums vary in susceptibility. While individual growers may sustain serious losses, the occurrence of this disease appears to be very sporadic and rare, particularly with the wide use of Thielaviopsis-free, peat-based potting mixes that have replaced field soilbased mixes. Symptoms/signs. Infected plants may be symptomless or may be stunted and have yellowed older foliage. Older infected roots may be very dark brown, almost black, while infected young roots may be only slightly darkened or may appear healthy unless observed microscopically. Thielaviopsis forms very dark brown, thick-walled, cigar-shaped spores in root cells that are readily found through microscopic observation and a thin-walled colorless spore on the surface of plant tissues (Linderman 1993). Biology/epidemiology. Thielaviopsis survives well in soil. If infested soil is brought into contact with plants, infection is likely to occur if environmental conditions and the physiological status of the plant favor disease development. It appears that when plants are under high temperature and low soil moisture stress, disease is likely to develop. Though rare, Thielaviopsis has been found in peat-based potting mixes (Graham and Timmer 1991). Management: • Cultural practices – Start with healthy, pathogen-free cuttings and seedlings. If the potting mix contains soil, pasteurize it before use. Black root rot is less severe in soil with a pH of 5–5.5, but growers should exercise caution when lowering the pH outside of the optimal range for plant growth.

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Fig. 2 Cercospora leaf spot on Pelargonium (Photo courtesy of Robert McGovern)

• Fungicides – They are available for suppressing Thielaviopsis but should be used routinely for this pathogen only if the operation has a history of crop losses from it. Discard infected plants.

2.3

Cercospora Leaf Spot (Cercospora Sp.)

Geographic occurrence and impact. The pathogen is reported to infect Pelargonium in North America and the Philippines (Daughtrey et al. 1995) and has been found on Geranium. Symptoms/signs. Initially small (1–5 mm; 1/8–1/4 in.) light green spots enlarge and become gray (Fig. 2). Dark fruiting structures develop within the gray area. As the spots enlarge, they may have a yellow area surrounding them, and the spots may merge. Infected flowers often do not open (Engelhard 1993). Biology and epidemiology. Little is known of the disease cycle of this pathogen in Pelargonium, but it is known that the pathogen is wind disseminated and that infection is greatly favored by wetness of the foliage (Daughtrey et al. 1995). Management. Irrigate plants in a manner that keeps water off the foliage. Remove infected foliage. Apply a fungicide to protect plants. The chlorothalonil and mancozeb fungicides are effective against Cercospora diseases. Check production guides for your region.

2.4

Gray Mold (Botrytis Blight; Botrytis cinerea Pers.: Fr.)

Geographic occurrence and impact. This disease is probably the most common and widespread one in Pelargonium production, and the pathogen infects many additional plants (Daughtrey et al. 1995). In fact, the very high susceptibility of Pelargonium results in Pelargonium being a significant harbor of Botrytis for other greenhouse crops (Hausbeck and Pennypacker 1991a; Hausbeck and Pennypacker 1991b).

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Fig. 3 Botrytis infected petal has fallen on a leaf. Botrytis grows from this food base into the healthy leaf (Photo courtesy of The Pennsylvania State University Dept. of Plant Pathology and Environmental Microbiology)

Fig. 4 Botrytis-infected Pelargonium cuttings (Photo courtesy of The Pennsylvania State University Dept. of Plant Pathology and Environmental Microbiology)

Symptoms/signs. Flowers turn dark and fall prematurely. Where infected petals land on leaves, Botrytis grows from this nutrient source, and an irregular leaf spot forms on the green leaf (Fig. 3). Cuttings develop a dark brown to black rot (Fig. 4) near the base (also see Pythium root rot, blackleg). Abundant, dusty, gray masses of spores form on infected tissue if the humidity is high or if infected tissue is placed in a container with moisture. Biology and epidemiology. All aboveground parts of the plant are susceptible, and the fungus can infect intact tissue or through wounds. Seedlings and mature plants are susceptible. Infection occurs most readily when the fungus has a food base from which it attacks, such as fading flowers or senescent leaves. High relative humidity, moisture on the plant, and temperatures between 18  C and 25  C (64  F and 77  F) greatly favor Botrytis infection (Jarvis 1980). The spores are readily spread by air currents, and any activity in the greenhouse that changes the relative humidity quickly or disturbs the plants can result in the release of spores (Hausbeck and Pennypacker 1991a, Hausbeck and Pennypacker 1991b). Management: • Cultural practices – Heat and ventilate plants in a greenhouse setting, and space plants to ensure good air circulation and low humidity within the crop canopy. It

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Fig. 5 Pythium root and blackleg (Photo courtesy of Robert McGovern)

is best to not crowd plants and to not hang plants above Pelargonium if those plants tend to drop faded flowers or if irrigating the pots above will result in excess irrigation water dripping onto the geraniums. Avoid unnecessarily damaging plants. Do not leave large stubs of tissue on the stock plant when taking cuttings. Remove and destroy fading flowers and leaves from the plant, the surface of the potting soil, from benches and greenhouse floors. Remove this material promptly from the greenhouse or place it in a closed container. • Fungicides – Apply a fungicide. Exclusive use of one class of chemical, particularly systemic chemicals, can result in the selection of resistance to the chemical class. Resistance to benzimidazole fungicides is very widespread in Botrytis populations. Do not rely on only one chemical class (Elad et al. 1992; Gullino and Garibaldi 1987; Hausbeck and Moorman 1996; Moorman and Lease 1992).

2.5

Pythium Root Rot and Blackleg (Pythium Sp.)

Geographic occurrence and impact. Many species of Pythium are known to be pathogens of Pelargonium. P. aphanidermatum, P. cryptoirregulare, P. irregulare, P. myriotylum, and P. ultimum are recovered the most frequently, but P. complectens (Phytopythium vexans), P. debaryanum (Globisporangium debaryanum), P. deliense, P. mamillatum (Globisporangium mamillatum), P. splendens (Globisporangium splendens), and P vexans (Phytopythium vexans) have reported. These species can be found worldwide, but a particular species may be more important in a given region than other species. Pythium root rot and blackleg (rot of the base of cuttings) can cause extensive crop losses. Symptoms/signs. Blackleg, the coal black rot of cuttings, first develops as brown water-soaked rot at the cut end of the tissue. The rot proceeds up the stem and kills the cutting rapidly. Seedling geraniums yellow and collapse when root rot occurs (Fig. 5). In the case of root rot of established plants, root tips appear translucent and

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water-soaked. The outer layers of root tissue strip off when plants are pulled from soil leaving the central core of vascular tissue bare. Usually, spherical spores can be found in root cells when examined microscopically. If Pythium is suspected but no spherical spores are observed in the tissue, plate the tissue on plain water agar, and Pythium usually grows out within 24 h. There are simple kits available commercially that can be used easily to detect the presence of Pythium sp. These immunoassay kits provide a result in less than 30 min. Biology/epidemiology. Pythium species are often found in field soil, sand taken from streams and rivers and can be found in pond and lake sediments, and dead roots of previous crops (Ivors and Moorman 2014). While some species are a problem in hydroponic systems, species of Pythium present in the clear water (not the sediment) of ebb and flood irrigation systems may not all be serious plant pathogens (Lanze 2015). It is important to know what species is present. Pelargonium most susceptible are those being grown at high fertilizer levels (Gladstone and Moorman 1989, Gladstone and Moorman 1990). Management: • Cultural practices – Pythium root rot is difficult to control once rot has begun. Every effort should be directed toward preventing the disease before it begins by using a heat-pasteurized potting mix. The entire pile must be heated to 75 –80  C (165 –180  F) and held at that temperature for 30 min. Potting mixes heated for very long times or temperatures higher than 80  C will result in the death of most beneficial organisms in the soil and create a “biological vacuum” that is readily filled by Pythium if the mix becomes contaminated in some way. If the commercial mix is purchased in sealed bags or bales, try to keep the covering on until used in order to prevent contamination. Pythium is easily introduced into pasteurized soil or soilless mixes by using dirty tools, dirty pots or flats, and dirty loading equipment, by walking on or allowing pets to walk on the mixes, and by dumping the mixes on benches or potting shed floors that have not been thoroughly cleaned and disinfected. Discard infected cuttings since affected rooted cuttings later develop root rot. Keep hose ends off the ground to prevent picking up contaminated soil. If pond or stream water is used for irrigation, be certain the intake pipe is well above the bottom so that sediment is not drawn in. If a known plant pathogenic species is found in the water, treatment may be required [Refer to Chapter 8, “▶ Sanitation in Florists’ Crops Disease Management,” and pertinent chapters in Hong et al. (2014)]. For example, slow sand filtration has been shown to be an effective, simple, and inexpensive method for removing Pythium from water. Heat, ultraviolet light, ozone, and chlorination can also be effective. The best of these methods is the one that can be used consistently and efficiently in an operation. Disinfect all bench surfaces, potting benches, tools, and equipment that will contact the potting mix. Periodically, thoroughly clean and disinfect ebb and flood reservoirs, benches, and flood and drain floors. • Fungicides and biocontrols – In a greenhouse operation with a history of Pythium root rot, apply a fungicide or a biological control agent as early in the cropping cycle as possible. Biological agents are generally applied to the potting

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mix before, during, or immediately after transplanting. However, read the label of the product to obtain information on exactly when the agent should be applied. In some cases, the agent must be in the potting mix for several days before plants are put in the mix in order to avoid phytotoxicity. Some biological control agents can be applied to plants in plug trays before transplanting. If chemical fungicides are also to be used, a general guideline is to not apply any chemical pesticides to the potting mix 10 d before and 10 d after applying the biological control agent. Biological control agents and fungicides may have to be applied more than once in order to maintain adequate protection for several weeks. They do not cure a plant once the plant is infected. Populations of some Pythium species have resistance to fungicides (Moorman and Kim 2004; Moorman et al. 2002).

2.6

Rhizoctonia Root and Crown Rot (Thanatephorus cucumeris (Rhizoctonia solani Kuhn) Rhizoctonia Sp.)

Geographic occurrence and impact. Although the pathogen is well known worldwide, this disease is not widespread in Pelargonium production, particularly with the use of soilless potting mixes. Symptoms and signs. A brown, dry rot develops at the base of cuttings (Manning et al. 1973) or seedlings are killed (Powell 1993). When infected tissue is incubated on a moist surface in a closed container, threads of fungal hyphae grow from the infected tissue directly down to the moist surface in 24 h. Biology and epidemiology. Little is known of this disease. It is reported that drought and high soluble salts renders plants more susceptible to the pathogen (Powell 1993). This species infects a wide range of host plants. Management. Every effort should be directed toward preventing the disease before it begins by using a heat-pasteurized potting mix. The entire pile must be heated to 75 –80  C (165 –180  F) and held at that temperature for 30 min. If this pathogen has been a problem before in production, the application of a fungicide is recommended in order to protect plants not yet infected.

2.7

Rust (Puccinia pelargonii-zonalis Doidge, Puccinia granularis, Puccinia morrisoni)

Geographic occurrence and impact. The pathogen was first found in South Africa (Doidge 1926) but now can be found sporadically wherever Pelargonium is grown. However, because it is readily recognized and prevented, it is generally not a major problem. Symptoms/signs. Chlorotic specks on the upper leaf surface appear directly opposite pustules of rust-colored spores on the underside of leaf. Spores erupt in concentric rings forming a “target” spot (Fig. 6). Biology/epidemiology. This rust requires only one host (autoecious; there is no alternate host plant), and it produces only two different types of spores: brown

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Fig. 6 Geranium rust on Pelargonium (Photo courtesy of The Pennsylvania State University Dept. of Plant Pathology and Environmental Microbiology)

single-celled urediniospores and pale brown, two-celled teliospores. When temperatures are between 16  C and 21  C (61  F and 70  F) and there is water on the leaves, the fungus can continuously produce urediniospores that germinate and infect plants through the stomata (Harwood and Raabe 1979). The urediniospores can be spread by wind or splashing or on workers’ hands and clothing. Apparently, teliospores form rarely. Management. Purchase rust-free cuttings. Irrigate plants in a manner that keeps water off the foliage. Discard unwanted geraniums at season’s end unless they are to be kept under observation and treated with fungicides if necessary. When infected plants are found, discard them and treat the remaining plants with a fungicide.

2.8

Sclerotinia Crown Rot or Cottony Stem Rot (Sclerotinia sclerotiorum (Lib.) de Bary)

Geographic occurrence and impact. This pathogen infects many different bedding plants in the seeding stage, including Pelargonium, where growing conditions are hot and humid. It is generally not a problem in greenhouses in temperate regions. Symptoms/signs. White, cottony fungal growth develops quickly near the soil line. As the mycelium ages, black sclerotia (up to 2–3 mm in size) develop on the infected tissue. Biology and epidemiology. The pathogen is carried in soil or on infected plants. It survives as sclerotia in crop debris and in soil. Management. If soil is a component of the potting mix, the potting mix should be heat pasteurized first. Benches, potting areas, and equipment used to move potting mix should be cleaned and disinfected periodically. Crop debris should be collected and removed from the growing area, particularly between cropping cycles. Infected plants should be discarded (Strider 1985).

Diseases of Geranium

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11

Verticillium Wilt (Verticillium albo-atrum Reinke and Berthe or V. dahliae Kleb.)

Geographic occurrence and impact. This is a problem primarily where plants are grown outdoors exposed to soil. But if infected plants or infested soil are shipped from place to place, this is an avenue for long distance spread to wherever Pelargonium is grown. Symptoms/signs. Middle and upper leaves yellow, collapse, dry, and fall. Vascular tissue of affected stems is browned or blackened. Symptoms are readily confused with those of bacterial blight. Sometimes, infected plants are severely stunted before any other symptoms are exhibited (Strider 1985). Biology and epidemiology. The pathogen survives well in soil and in the tissues of many species of infected plants. Management. Purchase culture-indexed cuttings. Use pasteurized potting mixes. Destroy infected plants. Fungus or fungus-like organisms reported to be associated with Pelargonium [See Farr et al. (1989) and Farr, and Rossman Fungal Databases, Systematic Mycology and Microbiology Laboratory, ARS, USDA. Retrieved from http://nt. ars-grin.gov/fungaldatabases/]: Aecidium pelargonii. Alternaria alternata. Alternaria pelargonii. Alternaria tenuis (Alternaria alternata). Alternaria tenuissima. Armillaria fuscipes. Armillaria heimii. Armillaria mellea. Ascochyta sp. Aspergillus fischeri. Aspergillus fischerianus (Aspergillus fischeri). Bipolaris maydis. Botryosphaeria berengeriana (Botryosphaeria dothidea). Botryosporium pulchrum. Botryotinia fuckeliana (Botrytis cinerea). Botrytis pelargonii. Calonectria morganii. Cladosporium fumago (Fumago salicina). Cladosporium sphaerospermum. Coleroa circinans. Colletotrichum gloeosporioides. Coniella australiensis. Coniothyrium trabutii. Cryptovalsa ampelina. Cylindrocarpon olidum (Thelonectria olida).

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Cylindrocladiella camelliae. Cylindrocladiella parva. Cylindrocladium scoparium. Cyphella pelargonii. Diaporthe medusae (Diaporthe rudis). Diaporthe rudis. Discohainesia oenotherae (Pilidium lythri). Drechslera setariae (Bipolaris setariae). Erysiphe communis (Erysiphe pisi var. pisi) Fibroidium pelargonii. Fusarium oxysporum. Fusarium pelargonii. Fusarium semitectum (Fusarium incarnatum). Gloeosporium pelargonii. Glomerella cingulata (Colletotrichum gloeosporioides). Helicobasidium purpureum. Leptosphaeria elaoudi. Leptosphaeria pelargonii. Macrophomina phaseoli (Macrophomina phaseolina). Macrosporium geraniaceae. Macrosporium pelargonii. Oidium sp. Patellaria atrata (Lecanidion atratum). Peroneutypa heteracantha. Pestalotia pelargonii. Pestalotia versicolor (Pestalotiopsis versicolor). Pestalotiopsis sp. Phaeosolenia pelargonii. Phyllosticta geraniicola. Physalospora geranii. Phytophthora cactorum. Phytophthora drechsleri. Phytophthora palmivora var. palmivora (Phytophthora palmivora) Phytophthora x pelgrandis. Pilidium concavum (Pilidium lythri). Pleosphaerulina sp. Pleospora herbarum. Pleospora phaeocomoides. Rhizoctonia solani. Schizophyllum commune. Sclerotiopsis sp. Sclerotium sp. Septoria canberrica. Septoria geranii. Septoria pelargonii.

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Sphaeropsis sp. Sphaerotheca fugax (Podosphaera fugax). Sphaerulina pelargonii. Sporotrichum epiphyllum. Stemphylium solani. Tuberculina pelargonii. Uredo pelargonii.

3

Bacterial and Phytoplasma Diseases

Where possible, the scientific names used are those accepted by the International Society of Plant Pathology Committee on the Taxonomy of Plant Pathogenic Bacteria (Bull et al. 2010).

3.1

Bacterial Blight (Xanthomonas hortorum Pv. pelargonii; Formerly, Xanthomonas campestris Pv. pelargonii)

Geographic occurrence and impact. At one time, this was the most serious disease of Pelargonium worldwide. The pathogen is spread in Pelargonium production and throughout the world wherever infected stock plants or cuttings are shipped. Losses can approach 100% in some cultivars. Since first reported in the early 1900s, this disease has increased or decreased depending upon the vigilance of growers, diligence in eliminating infected stock plants, and acceptance of growing plants from culture-indexed cuttings. Over the decades of Pelargonium production, outbreaks of this disease have severely damaged the reputations of specialty propagators. Seedling geraniums are susceptible to the pathogen but are free of the disease unless grown in close proximity to vegetatively propagated Pelargonium. Although perennial Geranium is not severely affected; they can be an ongoing source of the pathogen for Pelargonium (Nameth et al. 1999). Symptoms/signs (Figs. 7, 8, and 9). Small spots [less than 6.4 mm (1/4 in.)] develop on the underside of the leaf and become sunken and well defined. The leaf wilts and dies as the bacteria spread through water-conducting vessels of veins and petioles. V-shaped areas form with the wide part of V on the leaf margin and point of the V on veins. These symptoms are seen on both Pelargonium and Geranium. On Pelargonium, the leaf spotting may occur late in disease development. Prior to leaf spotting, lower leaves may wilt at the margins, while the blade and petiole remain turgid. As wilt progresses, the entire leaf collapses. The vascular tissue of the main stem on the side of the affected portion of the plant is discolored gray to brown. Lower leaves die and fall. Note that although many of these symptoms can be confused with those caused by Ralstonia (bacterial or southern wilt), Ralstonia does not cause leaf spots in Pelargonium. There are simple, rapid test immunological kits commercially available for testing plants for both Xanthomonas hortorum pv. pelargonii and for Ralstonia solanacearum, and there are molecular methods

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Fig. 7 Bacterial blight leaf spot (Xanthomonas) on Geranium (Photo courtesy of The Pennsylvania State University Dept. of Plant Pathology and Environmental Microbiology)

Fig. 8 Bacterial blight on ivy geranium (Pelargonium peltatum) (Photo courtesy of The Pennsylvania State University Dept. of Plant Pathology and Environmental Microbiology)

for identifying the presence of Xanthomonas hortorum pv. pelargonii in plant tissues (Sulzinski et al. 1996; Sulzinski et al. 1997). Note that other species are known to cause leaf spots on Pelargonium but that they are relatively minor problems (Rockey et al. 2015). Biology and epidemiology. The pathogen survives in infected plants and therefore is primarily a problem in Pelargonium propagated vegetatively. Disease develops most rapidly when temperatures are between 20  C and 30  C (68  F and 86  F). When the pathogen is splash dispersed and enters leaves through hydathodes, bacteria enter the vascular tissue and move throughout the plant. The only tissues not infected are the meristem tips where the vascular tissue is not fully differentiated. Viable bacteria survive on moist plant surfaces and within infected tissue and in the vascular tissue. At temperatures between 10  C and 15  C (50  F and 50  F), bacterial activity is greatly suppressed, and infected plants may remain symptomless. This can lead growers to believe that stock plants are not infected. However, as

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Fig. 9 Bacterial blight – vascular discoloration (Xanthomonas) in Pelargonium (Photo courtesy of The Pennsylvania State University Dept. of Plant Pathology and Environmental Microbiology)

temperatures rise, symptoms become apparent. In the meantime, if cuttings are taken from infected stock plants, the cuttings may root but will eventually exhibit symptoms. In addition to splash dispersal, the bacteria can be spread on workers’ hands and on knives used for taking cuttings (Munnecke 1954). Management. Purchase culture-indexed cuttings or grow plants from seed. Immediately discard infected plants after a positive diagnosis is made. Irrigate plants in a manner that keeps water off the foliage. Discard all unwanted geraniums at season’s end if they will not be examined periodically for symptoms. If cuttings are taken with knives, the knives should be washed and thoroughly disinfested after use on one plant and before use on another plant. If branches to be used for cuttings are broken off plants by hand, workers should wash their hands frequently or wear disposable gloves and disinfect them after use on one plant and before use on another plant.

3.2

Bacterial Fasciation (Rhodococcus fascians, Formerly Known as Corynebacterium fascians, Bacterium fascians, and Phytomonas fascians)

Geographic occurrence and impact. Although the bacteria can persist in infested soil, the probable main source of the pathogen in Pelargonium production is through the propagation from infected stock plants. For that reason the disease may appear anywhere Pelargonium is produced vegetatively propagation is grown (Strider 1985). Symptoms/signs. Short, thick, fleshy, leafy galls formed at base of main stems at or below soil level are pale green or green-yellow (Fig. 10). The rest of the plant appears healthy. Biology and epidemiology. Rhodococcus fascians survives associated with live plants or in soil. Many plants in addition to Pelargonium are susceptible to this

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Fig. 10 Bacterial fasciation on Pelargonium (Photo courtesy of The Pennsylvania State University Dept. of Plant Pathology and Environmental Microbiology)

pathogen. It is believed that Rhodococcus fascians does not require a wound to enter the plant. The pathogen is very difficult to grow in culture from infected plant tissues. Management. Purchase culture-indexed plants. Discard infected stock plants and cuttings and infested media.

3.3

Bacterial Wilt or Southern Wilt (Ralstonia solanacearum, Formerly Known as Burkholderia solanacearum, and Pseudomonas solanacearum)

Geographic occurrence and impact. Ralstonia solanacearum attack almost 200 plant species in 33 different plant families. The common name for the diseases this organism causes varies with the host that is attacked and geographic location. It is sometimes called southern wilt (in the northern hemisphere) or bacterial wilt. This bacterium is noted for diseases caused outdoors in land areas bounded by 45 N and 45 S latitudes where rainfall averages above 100 cm/year (39 in./year), the average growing season exceeds 6 months, the average winter temperatures are not below 10  C (50  F), the average summer temperatures are not below 21  C (70  F), and the average yearly temperature does not exceed 23  C (72  F) (Lucas 1975). Symptoms/signs. Lower leaves wilt, yellow, and fall. Vascular tissue of affected stems turns brown or black; roots may be discolored and rotted (Figs. 11 and 12). Note that although these symptoms can be confused with those caused by Xanthomonas (bacterial blight), Ralstonia does not cause leaf spots in Pelargonium. Slimy, sticky ooze forms tan-white to brownish beads where the vascular tissue is cut. When an infected stem is cut across and the cut ends held together for a few seconds, a thin thread of ooze can be seen as the cut ends are slowly separated. If one of the cut ends is suspended in a clear container of clean water, bacterial ooze will form a thread in the water. Back lighting of the container helps reveal the ooze. Biology and epidemiology. Although the primary location of survival in the environment is in crop and weed hosts, the bacteria can also survive in soil. They can be readily spread through the movement of contaminated soil and infected

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Fig. 11 Bacterial wilt (Ralstonia) (Photo courtesy of Robert McGovern)

Fig. 12 Bacterial wilt (Ralstonia) (Photo courtesy of Robert McGovern)

vegetatively propagated plants, in contaminated irrigation water, and on the surfaces of tools (cutting knives) and equipment used to work with the plants, and on soiled clothing. Populations within this genus and species can be divided into races and biovars based on differing host ranges, biochemical properties, susceptibility to bacteria-infecting viruses (phages), and serological reactions. Race 1 is endemic to North America where it attacks many floricultural and vegetable bedding plant crops including Pelargonium, Catharanthus, Impatiens, Ageratum, Chrysanthemum, Gerbera, Tagetes, Zinnia, Salvia, Capsicum, Lycopersicon, Nicotiana, Petunia, Solanum melongena (eggplant), Tropaeolum (nasturtium), and Verbena. Race 3 is tropical in distribution and does not occur naturally in North America. Race 3 biovar 2 infects potato (Williamson et al. 2002) and other hosts including Pelargonium, tomato, peppers, eggplant, bean, and beet. Weed hosts include black nightshade, climbing nightshade, horsenettle, Jimson weed, purslane, mustards, lamb’s-quarters,

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and bitter gourd. The bacteria can infect through roots and through any fresh wounds. The bacterium can be difficult to work with in the laboratory because it quickly loses pathogenicity and viability in artificial culture. Race 3 biovar 2 is considered a quarantinable pathogen. Management. Growing and propagating from pathogen-free plant material is the main way to avoid problems with Ralstonia, regardless of the race and biovar involved. It is imperative that propagators use pathogen-free potting soil or other media, establish stock plants that are tested and known to be free of the bacteria, train workers handling the stock plants in methods and procedures that prevent the pathogen from contaminating the potting soil or coming in contact with the stock plants, and then maintaining this system throughout the propagation phase of crop production. Do not bring ground-planted geraniums into the production area or propagate from them. Destroy infected plants. There are no chemicals or biological agents that adequately control Ralstonia. Infected plants MUST be discarded as soon as possible. The purchaser of cuttings or prefinished plants should isolate all new, incoming plants as if the health of the plants were unknown, even if the plants have been certified as healthy. New plants must not be commingled or dispersed among other plants in the greenhouse from other sources. This procedure is crucial because keeping plants originating from one source together limits the area that may need to be quarantined, sanitized, or isolated should Ralstonia solanacearum be found. If the pathogen is found in plants being irrigated using a recycling water system (such as ebb and flood), all susceptible plants in the system are at risk to infection. New, incoming plants should not be put into such a system being used for any other plants until their health is verified, and it is known that they do not harbor the pathogen.

3.4

Crown Gall (Rhizobium tumefaciens, Formerly Agrobacterium tumefaciens)

Geographic occurrence and impact. The bacteria can persist in infested soil, but the probable main source of the pathogen in Pelargonium is through the propagation of new plants from infected stock plants. Although this disease may appear anywhere Pelargonium are produced by vegetative propagation, it is usually recognized by the propagator and infected stock plants, and cuttings are discarded. Thus, the disease is very seldom a problem except where a local grower maintains the pathogen in a favorite stock plant (Strider 1985). An understanding of this pathogen’s biology and its disease cycle has significantly contributed to modern agriculture in the area of biotechnology and genetic engineering (Nester et al. 2005). Symptoms/signs. Spherical white- to light tan-colored galls develop near the soil line on the roots and stems of infected plants. Biology and epidemiology. Rhizobium tumefaciens survives in infested soil and in infected plants. The pathogen is spread through the movement of these materials from place to place. Wounding of plants through taking cuttings allows entry of the pathogen.

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Fig. 13 Pseudomonas leaf spot on Geranium (Photo courtesy of The Pennsylvania State University Dept. of Plant Pathology and Environmental Microbiology)

Management. Growing and propagating from pathogen-free stock plants is the main way to avoid crown gall. Propagators must use pathogen-free potting soil or other media, establish stock plants that are known to be free of the bacteria, and destroy infected plants. Ground-planted geraniums should not be brought into the production area nor should cuttings be taken from them.

3.5

Pseudomonas Leaf Spot (Bacterial Leaf Spot; Pseudomonas cichorii)

Geographic occurrence and impact. This disease was first described on Pelargonium X hortorum being grown in Florida, USA, under warm, very humid conditions (Engelhard et al. 1983). In greenhouses or outdoors, the disease can be very damaging particularly if plants are irrigated in a manner that puts water directly on the foliage. Long distance spread of the pathogen occurs when infected plants are shipped from place to place (Daughtrey et al. 1995). The pathogen can infect many different plants in addition to Pelargonium. Symptoms. Water-soaked spots, 5–10 mm (1/4–1/2 in.) in diameter, form on leaves (Fig. 13). Spots become dark brown to black and irregularly shaped. A yellow halo may surround each spot (Engelhard et al. 1983). Under ideal conditions, spots merge, and the entire leaf may be affected. Infected flower buds and sometime the entire inflorescence are blackened. Biology and epidemiology. Spotting severity increases with temperature between 16  C and 28  C (61  F and 82  F) but is greatly inhibited at temperatures above 28  C. High relative humidity greatly favors infection and spot enlargement (Jones et al. 1984). Sprinkler irrigation and rain splash the pathogen among leaves and flowers and provide the water on the plant tissues required by the bacteria for infection.

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Management. Purchase pathogen-free cuttings. It is of utmost importance to water plants in a manner that keeps leaf surfaces dry and to maintain low relative humidity among the plants. Discard infected plants and treat the remaining plants with a bactericide.

3.6

“Candidatus Phytoplasma Asteris”

Yellows diseases caused by phytoplasma can occur on Pelargonium. For instance, Candidatus phytoplasma asteris (Group 16SRI) was reported to infect geranium in Pakistan for the first time by Fahmeed et al. (2009).

4

Viral Diseases

The EPPO Panel on “Certification of Ornamentals” developed procedures for the production of healthy carnation, pelargonium, lily, narcissus, chrysanthemum, tulip, crocus, iris, begonia, impatiens, rose, freesia, hyacinth, kalanchoe, and petunia. A combination of thermotherapy, meristem tip culture and virus indexing can be used in geranium to eliminate viruses (Horst et al. 1977; Horst and Klopmeyer 1993). In the USA, certification programs are managed by the major commercial propagators. Pelargonium and other major flower crops are tested for the presence of the most prevalent viruses either by in-house plant pathologists or by companies that provide testing services. In the USA, phytosanitary certificates are required for intraand interstate plans movement. Data on pathogens of high consequence, including viruses, are recorded in the USA by the National Plant Diagnostic Network (https:// www.npdn.org/home). Where available, references for viruses are from Adams and Antoniw (2006).

4.1

Arabis Mosaic Virus (ArMV), Tobacco Ringspot Virus (TRSV), Tomato Ringspot Virus (ToRSV), and Artichoke Italian Latent Virus (AILV)

Geographic occurrence and impact. ArMV (Smith and Markham 1944) has been detected in 13 states in the USA where it has been the object of quarantine, but it is not widespread or established outside Europe. ArMV has a wide host range. TRSV (Fromme et al. 1927) geographic distribution is mainly in northern USA and China, but it has been found in Europe and Australia. More than 17 plant families are susceptible to the virus (Price 1940). ToRSV (Price 1936) is originally from the west coast of the USA (Frazier et al. 1961), and it is probably distributed to ornamental plants in many parts of the world. It can infect plants in 35 families. AILV (Majorana and Rana 1970; Vovlas et al. 1971) is found in Southern Italy and Bulgaria. AILV can infect a number of woody and herbaceous plants, including weeds.

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Symptoms/signs. ArMV is associated with mosaic symptoms in pelargonium. Pelargonium ringspot is a disease associated with both TRSV and ToRSV, but little evidence is found to confirm if the disease is the result of double or single infections. In fact, since the two viruses can infect the same hosts in the same geographic area, they have been often confused. ToRSV has also been associated with ringspots and mosaic symptoms in pelargonium. AILV in Pelargonium zonale produces severe leaf malformations and reduction in size, elongation of petioles, and stunted plant growth (Vovlas 1974). Biology and epidemiology. These three viruses are Nepoviruses in the family Secoviridae. Their genomes consist of two linear ss+RNA segments. ArMV is nematode transmitted (Fritzsche and Schmidt 1963; Harrison and Cadman 1959; Jha and Posnette 1959) and seed transmitted (Lister and Murant 1967, Murant and Lister 1967) and can be transmitted by dodder (Cuscuta sp.). Nematodes lose the virus during molting and do not transmit it to their progeny. TRSV is transmitted by the nematode Xiphinema americanum (McGuire 1964), by pollen and seeds, as well as by many insects (Dunleavy 1957; Komuro and Iwaki 1968; Messieha 1969; Schuster 1963) and mites (Thomas 1969) in a nonspecific manner. TRSV does not replicate in its nematode vector, is lost during molting, and is not transmitted to the nematode progeny. TRSV and ToRSV are not serologically related, and serological tests can be used to confirm their identification. ToRSV is transmitted by the nematode Xiphinema americanum (Téliz et al. 1966), but seed transmission has also been reported in some plant species. Small ssRNA satellite viruses have been associated with some strains of ToRSV, but scant information is available on the presence of these satellite viruses in pelargonium. AILV is transmitted by the nematode Longidorus apulus (Rana and Roca 1976), and it is not serologically related to any other Nepovirus. Management. Management of these viruses involves the use of virus-free seed and other propagative material, nematode avoidance through soil and water treatment where necessary, and elimination of weedy hosts.

4.2

Beet Curly Top Virus (BCTV)

Geographic occurrence and impact. The virus originated from the Eastern Mediterranean basin and is common in the Western part of the USA from Mexico to Canada (Bennett and Tanrisever 1957). Today it is present also in Africa, Asia, and Central and South America. BCTV has a broad host range and can infect plant species in 44 families (Bennett 1971). Symptoms/signs. The disease caused by BCTV on pelargonium, called leaf cupping, is very severe, with pronounced tissue yellowing, leaf curling, and distortion. Biology and epidemiology. BCTV is a member of the family Geminiviridae with an ss+DNA genome contained in geminate particles. Two species of leafhoppers from the arid and semi-arid regions, Circulifer tenellus described by Ball (1909) and C. opacipennis (Kheyri and Alimoradi 1969; Stahl and Carsner 1923), are vectors of

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Fig. 14 Symptoms of INSV (Photo courtesy of Jay Pscheidt)

the virus, and the virus is phloem limited and does not multiply in its insect vectors. The virus is also not transmitted through the vector eggs and requires a latent period before transmission to plants. The virus is present in the seed but cannot invade the embryo. Management. Control of this virus consists of controlling the insect vector, excluding infected plants, and using clean propagative material. Usually plants infected with this virus do not survive and thus are not found commercially.

4.3

Impatiens Necrotic Spot Virus (INSV) and Tomato Spotted Wilt Virus (TSWV)

Geographic occurrence and impact. INSV (Law and Moyer 1990) and TSWV (Samuel et al. 1930) together with their thrips vectors are prevalent worldwide in many plant families and in hundreds plant species, making their control particularly challenging. Symptoms/signs. Symptoms associated with INSV and TSWV are mosaic, mottling, stem and leaf necrosis, ringspots on leaves, leaf deformation, and stunted growth (Fig. 14). Symptomatology is driven by plant phenology and environmental conditions. Biology and epidemiology. INSV and TSWV are in the family Bunyaviridae. These ssRNA viruses have tripartite genomes, with negative or ambisense orientation. INSV and TSWV are transmitted by thrips in a propagative (persistent manner) (Ullman et al. 1993; Wijkamp et al. 1993); thus, they can replicate in their plants as well as in their insect hosts. Management. One of the best ways to manage these viruses is to exclude thrips from greenhouses and to reduce their numbers, since thrips need a relatively long acquisition and transmission time, and since the viruses encounter a latent period in the vector prior to becoming transmissible. Unfortunately, thrips are now resistant to many insecticides (Brødsgaard 1994; Zhao et al. 1995). Weed management is also necessary (Bond et al. 1983; Cho et al. 1986; Kobatake et al. 1984). (For additional

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information, refer to Chapter 3, “▶ Insect Management for Disease Control in Florists’ Crops”).

4.4

Pelargonium Zonate Spot Virus (PZSV), Cucumber Mosaic Virus (CMV)

Geographic occurrence and impact. PZSV is geographically restricted to Southern Italy. The virus can infect tomato and artichoke and is common in weeds. CMV is found worldwide, has an extremely broad host range, and can infect species in more than 100 plant families (Zitter and Murphy 2009). Symptoms/signs. Symptoms induced by PZSV are leaf malformations, puckering, and zonate yellow bands (Martelli and Cirulli 1969; Quacquarelli and Gallitelli 1979). CMV in geranium causes flower breaking disease. Biology and epidemiology. PZSV and CMV (described by Doolittle 1916; Jagger 1916, and more recently by Jacquemond 2012; Palukaitis and Garcia-Arenal 2003) have tripartite ss+RNA genomes. Both viruses are in the family Bromoviridae. PZSV belong to the Anulavirus genus (Gallitelli 1982) and CMV to the Cucumovirus genus (Wildy 1971). PZSV is seed and pollen transmitted but apparently not vector-transmitted. Thrips can serve as carriers of the infected pollen from host to host. CMV is aphid transmitted (Kennedy et al. 1962) in a non-propagative (nonpersistent) manner (Hoggan 1933; Simons 1955; Watson and Roberts 1939), and it can be seed transmitted in some plant species (Neergaard 1977). Management. PZSV can be controlled using clean propagative material. Control for CMV is particularly difficult, since the virus is common on many plant families, and it can be transmitted by multiple aphid species. Because aphids transmit CMV readily during probing, aphid control is particularly important for reducing the spread of the virus. In open field, the use of mineral oil to discourage aphid feeding has been used especially in Europe, but thus far its use in a greenhouse setting has been limited.

4.5

Tobacco Mosaic Virus (TMV) and Tobacco Rattle Virus (TRV)

Geographic occurrence and impact. These viruses can infect a variety of plants in 30 families for TMV (Shew and Lucas 1991) and more than 50 (Horváth 1978; Noordam 1956; Schmelzer 1957; Uschdraweit and Valentin 1956) for TRV in Europe, Japan, New Zealand, and North America. Symptoms/signs. Symptoms attributed to TMV (Siegel and Wildman 1954) are often caused by other tobamoviruses, and many tobamovirus species were once classified as strains of TMV. Symptoms on pelargonium are reported to be mosaic and small light colored or brown lesions on leaves. Biology and epidemiology. TMV and TRV belong to the Virgaviridae family. These viruses have an ss+RNA genome. TMV is occasionally transmitted by chewing insects, but most commonly, it is mechanically spread (Harris and Bradley

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1973; Lojek and Orlob 1969); in fact TMV virions are extremely stable. TMV can persist in the soil, probably on plant debris, and can infect roots. It can penetrate wounded embryos from the infected seed coat (Broadbent 1965). TMV is a special concern in greenhouses, where it can be very hard to eradicate (Broadbent and Fletcher 1963). TRV is transmitted by nematodes, Paratrichodorus spp. and Trichodorus spp., (Taylor and Brown 1997), and pelargonium can become infected with this virus if grown in soil infested with viruliferous nematodes. TRV is also transmitted mechanically and by seed in certain hosts. Management. Sanitation is the best TMV and TRV control method. Sanitation practices include the use of clean tools and pots, disinfected soil, virus-free seeds, and prompt removal of infected plants. Control of nematodes is essential to limit TRV spread; thus, the use of clean irrigation water and sanitized soil is recommended.

4.6

Tomato Bushy Stunt Virus (TBSV), Tobacco Necrosis Virus (TNV), Moroccan Pepper Virus (MPV), and Pelargonium Leaf Curl Virus (PLCV)

Geographic occurrence and impact. TBSV has a restricted natural host range and causes damage primarily on agricultural crops, while it can experimentally infect more than 120 plant species. It is geographically distributed in Europe, the Americas, and North Africa. TNV has a natural broad host range (Price 1940) that include tobacco, cucumber, melon, European strawberry, citrus, apple, and pear. It can experimentally infect more than 37 plant families, and it is distributed worldwide. PLCV (Hollings and Stone 1965) is common in the USA and Mediterranean region. It was detected in India in 2013 (Kumar et al. 2013). Susceptible host species are Pelargonium zonale, common bean, Datura, and Chenopodium. Some of these species are considered experimental hosts. MPV is present in Morocco, Iran, USA, Europe, and Central Asia (Wintermantel and Hladky 2013). Its host range includes tomato, lettuce, pepper, pelargonium, jimson weed, and lisianthus. Symptoms/signs. These viruses induce Pelargonium leaf curl or crinkle, and Pelargonium necrotic spot, with stunted growth, leaf deformation and splitting, extensive spots first hyaline and then necrotic, and yellowing and necrosis of leaves. Symptoms are strongly influenced by temperature. In pelargonium, TNV can cause symptomless infection that turns symptomatic under different light and temperature conditions. PLCV is the causal agent of leaf curl disease. The disease symptoms are leaf curl and yellow stellate spots that are more pronounced in winter and at lower temperatures and diminish in summer at higher temperatures. Biology and epidemiology. These are monopartite ss+RNA viruses and are members of the Tombusvirus genus in the family Tombusviridae. TBSV (Ainsworth 1936, Bawden and Pirie 1938 Smith 1935) is soilborne and is transmitted by infected seed and soil. Defective interfering RNAs (DI-RNAs) and satellite RNAs have been associated with TBSV in its naturally infected plants (Galetzka et al. 2000, Gallitelli

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Fig. 15 Symptoms of PFBV (Photo courtesy of Robert Wick)

and Hull 1985), but were not investigated in pelargonium. TNV (Babos and Kassanis 1963; Bawden 1941; Smith and Bald 1935) is a soilborne virus and is transmitted by zoospores of the chytrid fungus Olpidium brassicae (Kassanis and MacFarlane 1964, Teakle 1962, Teakle and Gold 1963). TNV can support the multiplication of satellite viruses in some plants and serologically can be divided into groups, but group serology does not coincide with symptomatology. PLCV is serologically related to TBSV. MPV (Makkouk et al. 1981) is distantly serologically related to TBSV. PLCV and MPV were described to be associated with soil infested by Olpidium spp. in Iran in 2011 (Rasoulpour and Izadpanah 2011). PLPV is an unassigned species in the Tombusviridae. Management. Virus control requires the use of clean propagative material and soil, avoiding the spread of the vectoring zoospores by removing infested soil and water, and good sanitation practices. Research on virus-resistant varieties is needed.

4.7

Pelargonium Flower Break Virus (PFBV)

Geographic occurrence and impact. Virus described by Stone and Hollings (1973). It infects six plant families and has limited host range. The virus was reported to be the most common viral pathogen of Pelargonium sp. in Western Europe (Krczal et al. 1995). Symptoms/signs. It can infect Pelargonium domesticatum and can cause flowerbreaking in several cultivars, but it remains usually symptomless on P. peltatum and P. zonale. Other symptoms can include flower streaking, stunting, and chlorotic spotting (Fig. 15). Seedlings of pelargonium-type “Nittany Lion” appear to be immune to PFBV (Kemp 1969). Biology and epidemiology. PFBV is an ss+RNA virus belonging to the genus Carmovirus in the family Tombusviridae. PFBV is transmitted mechanically and by vegetative propagation, but does not seem to be transmitted by seed. Transmission of the virus in recirculating irrigation systems and by thrips via pollen has been

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demonstrated (Krczal et al. 1995). The virus is not serologically related to other viruses. Management. Use of clean propagative material and discarding of infected plants is recommended. Hollings and Stone (1974) reported that Pelargonium can be freed from PFBV by thermotherapy, but plant survival rate was low. Other viruses associated with Pelargonium: Tombusviridae family: Pelargonium chlorotic ring pattern virus. Pelargonium leaf curl virus. Pelargonium line pattern virus. Pelargonium necrotic spot virus. Pelargonium vein banding virus is a virus related to viruses in the Badnavirus genus, not yet approved as a species. Pelargonium vein clearing is in the genus Nucleorhabdovirus. It has a monopartite ss-RNA genome and replicates in the plant cell nucleus. Other viruses: Cherry leaf roll virus. Lilac chlorotic leaf spot virus. Tomato black ring virus.

5

Nematode Diseases

Nematodes, including dagger nematode (Xiphinema sp.), foliar nematode (Aphelenchoides sp.), and root-knot nematode (Meloidogyne sp.), have been recorded to occur on Pelargonium but do not appear to be a widespread problem (Strider 1985).

6

Abiotic Diseases

A myriad of factors including air pollutants, misuse of plant growth regulators, and individual cultivar responses to environmental conditions can result in symptoms on Pelargonium (Freeman 1993).

6.1

Edema

Geographic occurrence and impact. This problem can occur wherever Pelargonium is grown. It is most damaging to ivy geranium (Pelargonium peltatum). (Balge et al. 1969; Digat and Albouy 1976)

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Fig. 16 Edema on ivy geranium (Pelargonium peltatum) (Photo courtesy of The Pennsylvania State University Dept. of Plant Pathology and Environmental Microbiology)

Symptoms/signs. Small water-soaked pimples or blisters form on the underside of lower leaves (Fig. 16). The blisters become corky and brown as they enlarge. Severely affected leaves fall. Biology and epidemiology. Edema development is favored when the weather is cool, cloudy, and humid while the soil is wet, particularly if mites are feeding on the plants. Although over the decades scientists have referred to this as a physiological disorder and have not linked it to a biotic pathogen, it is interesting to note that symptoms can be induced in otherwise healthy plants by injecting them with sap taken from a plant with edema (Digat and Albouy 1976). Management. Spacing plants to provide good air circulation, using a welldrained potting mix, not overwatering during cool, cloudy weather, and mite suppression, are reported to help in preventing the development of edema.

6.2

Heat Stress

Geographic occurrence and impact. This problem can occur wherever Pelargonium is grown. Sensitivity to high temperatures varies greatly among cultivars (Strider 1985). Symptoms/signs. Upper leaves and stems completely lose chlorophyll. Sometimes, only the center of the leaf is bleached. Usually only a few leaves are affected, but in severe cases, most of the leaves and stems may appear bleached. Biology and Epidemiology. Pelargonium exposed to temperatures above about 28  C/82  F for more than 12 h develop symptoms. While some cultivars exhibit severe symptoms, others in the same greenhouse may be completely free of symptoms. Management. Monitor temperatures in the area where cultivars known to be heat sensitive are grown. If it appears that temperatures could persist above 28  C for more than 12 h, mist the sensitive cultivars so that evaporative cooling occurs.

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However, many of the pathogens detailed elsewhere in this section are favored when moisture is on the foliage. Apply only enough moisture to slightly dampen the leaf surface and then evaporate completely in minutes. Do not mist to the extent that water begins to drip off the foliage.

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Engelhard AW (1993) Foliar diseases: fungal leaf spots. In: White JW (ed) Geranium IV. Ball Publishing, Geneva, pp 228–230 Engelhard AW, Mellinger HC, Ploetz RC, Miller JW (1983) A leafspot of florists’ geranium incited by Pseudomonas cichorii. Plant Dis 67:541–544 Fahmeed F, Rosete YA, Pérez KA, Boa E, Lucas J (2009) First report of ‘Candidatus Phytoplasma asteris’ (group 16SrI) infecting fruits and vegetables in Islamabad, Pakistan. J Phytopathol 157:639–641 Farr DF, Bills GF, Chamuris GP, Rossman AY (1989) Fungi on plants and plant products in the United States. APS Press, St. Paul Frazier NW, Yarwood CE, Gold AH (1961) Yellow bud virus endemic along California coast. Pl Dis Rep 45:649–651 Freeman RN (1993) Physiological and environmental disorders. In: White JW (ed) Geraniums IV. Ball Publishing, Geneva, pp 351–362 Fritzsche R, Schmidt HB (1963) Xiphinema paraeolongatum Altherr und Xiphinema n. sp., zwei Vektoren des Arabis-Mosaikvirus. Naturwissenschaften 50:163 (In German) Fromme FD, Wingard SA, Priode CN (1927) Ringspot of tobacco; an infectious disease of unknown cause. Phytopathology 17:321–328 Galetzka D, Russo M, Rubino L, Krczal G (2000) Molecular characterization of a tombusvirus associated with a disease of statice [Goniolimon tataricum (L.) Boiss.]. J Plant Pathol 82:151–155 Gallitelli D (1982) Properties of a tomato isolate of pelargonium zonate spot virus. Ann Appl Biol 100:457–466 Gallitelli D, Hull R (1985) Characterization of satellite RNAs associated with tomato bushy stunt virus and five other definitive tombusviruses. J Gen Viro 66:1533–1543 Gladstone LA, Moorman GW (1989) Pythium root rot of seedling geraniums associated with various concentrations of nitrogen, phosphorus, and sodium chloride. Plant Dis 73:733–736 Gladstone LA, Moorman GW (1990) Pythium root rot of seedling geraniums associated with high levels of nutrients. HortSci 25:982 Graham JH, Timmer NH (1991) Peat-based media as a source of Thielaviopsis basicola causing black root rot on citrus seedlings. Plant Dis 75:1246–1249 Gullino ML, Garibaldi A (1987) Control of Botrytis cinerea resistant to benzimidazoles and dicarboximides with mixtures of different fungicides. Med Fac Landbouww Rijksuniv Gent 52:895–900 Harwood CA, Raabe RD (1979) The disease cycle and control of geranium rust. Phytopathology 69:923–927 Harris KF, Bradley RHE (1973) Importance of leaf hairs in the transmission of tobacco mosaic virus by aphids. Virology 52:295–300 Harrison BD, Cadman CH (1959) Role of a dagger nematode (Xiphinema sp.) in outbreaks of plant diseases caused by Arabis mosaic virus. Nature (London) 184:1624–1626 Hausbeck MK, Moorman GW (1996) Managing Botrytis in greenhouse-grown flower crops. Plant Dis 80:1212–1219 Hausbeck MK, Pennypacker SP (1991a) Influence of grower activity and disease incidence on concentrations of airborne conidia of Botrytis cinerea among geranium stock plants. Plant Dis 75:798–803 Hausbeck MK, Pennypacker SP (1991b) Influence of grower activity on concentrations of airborne conidia of Botrytis cinerea among geranium cuttings. Plant Dis 75:1236–1243 Hoggan IA (1933) Some factors involved in aphid transmission of the Cucumber-mosaic virus to tobacco. J Agric Res 47:689–704 Hollings M, Stone OM (1965) Studies of pelargonium leaf curl virus. Ann Appl Biol 56:87–98 Hollings M, Stone OM (1974) CMI/AAB Descr Pl Viruses No 130, 4 pp Hong CX, Moorman GW, Wohanka W, Büttner C (eds) (2014) Biology, detection, and management of plant pathogens in irrigation water. APS Press, St. Paul

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Makkouk KM, Koenig R, Lesemann DE (1981) Characterization of a tombusvirus isolated from eggplant. Phytopathology 71:572–577 Manning WJ, Vardaro PM, Cox EA (1973) Root and stem rot of geranium cuttings caused by Rhizoctonia and Fusarium (Pelargonium hortorum). Plant Dis Rep 57:177–179 Martelli GP, Cirulli M (1969) Le virosi delle piante ortensi in Puglia. Una maculatura gialla del pomodoro causata dal virus della necrosi perinervale del tabacco (Tobacco streak virus). Phytopathol Mediterr 8:154–156 (In Italian) McGuire JM (1964) Efficiency of Xiphinema americanum as a vector of tobacco ringspot virus. Phytopathology 54:799–801 Messieha M (1969) Transmission of tobacco ringspot virus by thrips. Phytopathology 59:943–945 Moorman GW, Kim SH (2004) Species of Pythium from greenhouses in Pennsylvania exhibit resistance to propamocarb and mefenoxam. Plant Dis 88:630–632 Moorman GW, Lease RJ (1992) Benzimidazole- and dicarboximide-resistant Botrytis cinerea from Pennsylvania greenhouses. Plant Dis 76:477–480 Moorman GW, Kang S, Geiser DM, Kim SH (2002) Identification and characterization of Pythium species associated with greenhouse floral crops in Pennsylvania. Plant Dis 86:1227–1231 Munnecke DE (1954) Bacterial stem rot and leaf spot of Pelargonium. Phytopathology 44:627–632 Munnecke DE (1956) Alternaria leaf spot of geranium. Plant Dis Rep 40:452–454 Murant AF, Lister RM (1967) Seed-transmission III: The ecology of nematode-borne viruses. Ann Appl Biol 59:63–76 Nameth ST, Daughtrey ML, Moorman GW, Sulzinski MA (1999) Bacterial blight of geranium: A history of diagnostic challenges. Plant Dis 83:204–212 Neergaard P (1977) Seed-borne viruses. Chapter 3. In: Seed pathology, Vol I. MacMillan Press, London/Madras, pp. 839 Nester E, Gordon MP, Kerr A (eds) (2005) Agrobacterium tumefaciens: from plant pathology to biotechnology. APS Press, St. Paul Noordam D (1956) Waardplanten en toestplanten van het ratelvirus van de tabak. Tijdschr Plantenziekten 62:219–225 (In Dutch) Oglevee-O’Donovan W (1993) Clean stock production: culture-indexing for vascular wilts and viruses. In: White JW (ed) Geraniums IV. Ball Publishing, Geneva, pp 277–286 Palukaitis P, García-Arenal F (2003) Cucumoviruses. Adv Virus Res 62:241–323 Powell CC (1993) Seedling diseases. In: White JW (ed) Geraniums IV. Ball Publishing, Geneva, pp 215–219 Price WC (1936) Specificity of acquired immunity from tobacco-ringspot diseases. Phytopathology 26:665–675 Price WC (1940) Comparative host ranges of six plant viruses. Am J Bot 27:530–541 Quacquarelli A, Gallitelli D (1979) Three viruses of Pelargonium zonale in Apulia. Phytopathol Mediterr 18:61–70 Rana GL, Roca F (1976) Trasmissione con nematodi del virus “latente Italiano del carciofo” (AILV). In: Atti del 2 Congresso Internazionale di Studi sul Carciofo, Bari 1973, pp. 855–858 (In Italian) Rasoulpour R, Izadpanah K (2011) Isolation and partial characterization of Pelargonium leaf curl virus, Moroccan pepper virus and Eggplant mottled crinkle virus from plant and soil in Iran. J Phytopathol 159:802–804 Rockey W, Potnis N, Timilsina S, Hong JC, Vallad GE, Jones JB, Norman DJ (2015) Multilocus sequence analysis reveals genetic diversity in xanthomonads associated with poinsettia production. Plant Dis 99:874–882 Samuel G, Bald JG, Pittman HA (1930) Investigations on ‘spotted wilt’ of tomatoes. Aust Counc Sci Ind Res Bull 44:8–11 Schmelzer K (1957) Untersuchungen über den Wirtspflanzenkreis des Tabakmauche-Virus. Phytopathol Z 30:281–314 (In German) Schuster MF (1963) Flea beetle transmission of Tobacco ringspot virus in the Lower Rio Grande Valley. Plant Dis Rep 47:510–511

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Shew HD, Lucas GB (1991) Compendium of tobacco diseases. APS Press, St. Paul Siegel A, Wildman SG (1954) Some natural relationships among strains of tobacco mosaic virus. Phytopathology 44:277–282 Simons JN (1955) Some plant-vector-virus relationships of southern cucumber mosaic virus. Phytopathology 45:217–219 Smith KM (1935) A new virus disease of the tomato. Ann Appl Biol 22:731–741 Smith KM, Bald JG (1935) A description of a necrotic virus disease affecting tobacco and other plants. Parasitology 27:231–245 Smith KM, Markham R (1944) Two new viruses affecting tobacco and other plants. Phytopathology 34:324–329 Stahl CF, Carsner E (1923) A discussion of Eutettix tenellus Baker as a carrier of Curly top of sugar beets. J Econ Ent 16:476–479 Stone OM, Hollings M (1973) Some properties of pelargonium flower-break virus. Ann Appl Biol 75:15–23 Strider DL (1985) Geranium. In: Strider DL (ed) Diseases of floral crops, vol 2. Praeger Scientific, New York, pp 111–187 Sulzinski MA, Moorman GW, Schlagnhaufer B, Romaine CP (1996) Characteristics of a PCR-based assay for in planta detection of Xanthomonas campestris pv pelargonii. J Phytopathol 144:393–398 Sulzinski MA, Moorman GW, Schlagnhaufer B, Romaine CP (1997) A simple DNA extraction method for PCR-based detection of Xanthomonas campestris pv. pelargonii in geraniums. J Phytopathol 145:213–215 Taylor CE, Brown DJF (1997) Nematode vectors of plant viruses. CAB International, Wallingford Teakle DS (1962) Transmission of tobacco necrosis virus by a fungus, Olpidium brassicae. Virology 18:224–231 Teakle DS, Gold AH (1963) Further studies of Olpidium as a vector of tobacco necrosis virus. Virology 19:310–315 Téliz D, Grogan RG, Lownsbery BF (1966) Transmission of tomato ringspot, peach yellow bud mosaic, and grape yellow vein diseases by Xiphinema americanum. Phytopathology 56:658–663 Thomas C (1969) Transmission of tobacco ringspot virus by Tetranychus sp. Phytopathology 59:633–636 Ullman DE, German TL, Sherwood JL, Westcot DM, Cantone FA (1993) Tospovirus replication in insect vector cells: immunocytochemical evidence that the nonstructural protein encoded by the S RNA of tomato spotted wilt tospovirus is present in thrips vector cells. Phytopathology 83:456–463 Uschdraweit HA, Valentin H (1956) Das Tabakmauchevirus an Zierpflanzen. Nachrichtenbl Dtsch Pflanzenschutzdienst 8:132–133 (In German) Vovlas C (1974) Le malformazioni fogliari, una nuova virosi del geranio. Phytopathol Mediterr 13:139–142 (In Italian) Vovlas C, Martelli GP, Quacquarelli A (1971) Le virosi delle piante ortensi in Puglia. Il complesso delle maculature anulari della cicoria. Phytopathol Mediterr 10:244–254 (In Italian) Watson MA, Roberts FM (1939) A comparative study of the transmission of Hyoscyamus virus 3, potato virus Y and cucumber virus 1 by the vector Myzus persicae (Sulz.), M. circumflexus (Buckton) and Macrosiphum gei (Koch). Proc R Soc Ser B 127:543–576 Wijkamp I, van Lent J, Kormelink R, Goldbach R, Peters D (1993) Multiplication of tomato spotted wilt virus in its insect vector, Frankliniela occidentalis. J Gen Virology 74:341–349 Wildy P (1971) Classification and Nomenclature of Viruses. First Report of the International Committee on Nomenclature of Viruses. Monographs in Virology no. 5. Basel: Karger. Basel, pp. 33–34 Williamson L, Nakaho K, Hudelson B, Allen C (2002) Ralstonia solanacearum race 3, biovar 2 strains isolated from geranium are pathogenic on potato. Plant Dis 86(9):987–991

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Wintermantel WM, Hladky LL (2013) Complete genome sequence and biological characterization of Moroccan pepper virus (MPV) and reclassification of Lettuce necrotic stunt virus as MPV. Phytopathology 103(5):501–8 Zhao G, Liu W, Brown JM, Knowles CO (1995) Insecticide resistance in field and laboratory strains of western flower thrips (Thysanoptera: Thripidae). J Econ Entomol 88:1164–1170 Zitter TA, Murphy JF (2009) Cucumber mosaic. The Plant Health Instructor. doi:10.1094/PHI-I2009-0518-01

Diseases of Hydrangea Yonghao Li, Margaret T. Mmbaga, Boru Zhou, Jacqueline Joshua, Emily Rotich, and Lipi Parikh

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Powdery Mildew (Golovinomyces orontii DC) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Cercospora Leaf Spot (Cercospora hydrangea L.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Botrytis Blight (Botrytis cinerea Pers.: Fr.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Rust (Pucciniastrum hydrangeae (B. & C.) Arth.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Anthracnose (Colletotrichum gloeosporioides (Penz.) Penz. and Sacc.) . . . . . . . . . . . . . 2.6 Corynespora Leaf Spot (Corynespora cassiicola (Berk. & M.A. Curtis) C.T. Wei) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7 Phoma Leaf Spot (Phoma spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.8 Myrothecium Leaf Spot and Blight (Myrothecium roridum Tode Ex Fr.) . . . . . . . . . . . . 2.9 Alternaria Leaf Spot (Alternaria spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Bacterial Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Bacterial Leaf Spot (Xanthomonas campestris) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Viral Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Hydrangea Mosaic Virus (HdMV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Hydrangea Ringspot Virus (HRSV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Tomato Ringspot Virus (ToRSV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4 Tobacco Ringspot Virus (TRSV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Y. Li (*) Department of Plant Pathology and Ecology, The Connecticut Agricultural Experiment Station, New Haven, CT, USA e-mail: [email protected] M.T. Mmbaga • J. Joshua • E. Rotich • L. Parikh Department of Agricultural and Environmental Sciences, Tennessee State University, Nashville, TN, USA e-mail: [email protected]; [email protected]; [email protected]; lp. [email protected] B. Zhou Department of Forest Protection, Northeast Forestry University, Harbin, China, e-mail: [email protected] # Springer International Publishing AG (outside the USA) 2016 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florists' Crops Diseases, Handbook of Plant Disease Management, DOI 10.1007/978-3-319-32374-9_36-1

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4.5 Tomato Spotted Wilt Virus (TSWV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6 Cherry Leaf Roll Virus (CLRV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7 Alfalfa Mosaic Virus (AMV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract

Hydrangea, a native to Asia and America, is a popular ornamental plant due to its interesting large, showy flowers with different colors and inflorescence forms. Many hydrangea species are often used to design landscapes and gardens. In greenhouses, nurseries, and landscapes, hydrangea production and values are affected by fungal, bacterial, and viral diseases. In this chapter, the biology of pathogens, development of diseases, and strategies for their control of common diseases of hydrangea were described. Keywords

Hydrangea • Powdery mildew • Leaf spot • Rust • Anthracnose • Bacterial • Virus

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Introduction

Hydrangeas are among the most loved flowering shrubs and vines in landscapes. Many species in the genus Hydrangea vary in shrub size, texture, flower shape, and color, and their aesthetic values include popular floral arrangements and crafts (Mmbaga et al. 2008). The most common hydrangea is the bigleaf/French/garden hydrangea (H. macrophylla (Thunb.) Ser.) that thrives in hardiness US zones 6–9 (Dirr 2004). Although more than 700 cultivars have been described in H. macrophylla, only about 25% of these are available in the US trade (Dirr 2004; Van Gelderen and Van Gelderen 2004). Several other common species in this genus are climbing hydrangea (H. anomala), oakleaf hydrangea (H. quercifolia), smooth hydrangea (H. arborescens), and Pee Gee hydrangea (H. paniculata) (Dirr 2004).

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Fungal Diseases

2.1

Powdery Mildew (Golovinomyces orontii DC)

Geographical occurrence and impact. Powdery mildew of hydrangea has been reported in the Unites States and Korea (Park et al. 2012). The disease is a common problem in bigleaf hydrangea (H. macrophylla) in greenhouses, nurseries, and landscapes when environment conditions are warm and humid. The disease also affects H. serrata as well as smooth (H. arborescens), bigleaf (H. macrophylla), and panicle (H. paniculata) hydrangeas (Hagan and Mullen 2001). Symptoms/signs. Early signs of powdery mildew on hydrangea consist of circular light gray patches composed of mycelia and spores on both sides of the

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leaf surface. These pustules of fungal tissue eventually turn into yellow or purple blotches on the leaf (Fig. 1). As the disease progresses, disfiguring white mycelium develops on leaf surfaces along with extensive chlorosis or yellowing of the leaves (Fig. 2). Premature defoliation may occur on severely infected plants. Growth of the plant may be impacted with the reduction in active leaf area and shoot elongation directly reducing the production of photosynthesis (Hagan et al. 2004; Sinclair and Lyon 2005). Powdery mildew is rarely fatal to plants in landscape settings. Its major damage is the reduction in aesthetic quality and plant vigor. In addition economic values of infected plants are lowered and may even render plants unmarketable. Biology and epidemiology. The bigleaf hydrangea powdery mildew is caused by G. orontii (formerly Erysiphe polygoni DC.) and E. poeltii (Pscheidt 2014). Powdery mildew caused by Oidium hortensiae on H. macrophylla was first reported in Korea in 2012 (Park et al. 2012). The pathogen is an obligate parasitic fungus that occupies the leaf surface of the plant (Li et al. 2009a). The disease is favored by dry conditions, high humidity, and warm day/cool night temperatures. The pathogen obtains its nutrition from epidermal cells that are parasitized by root-like structures called haustoria (Li et al. 2008). The fungus overwinters as hyphae or spores in buds of previously infected plants in the landscape and living plants in greenhouses. Management • Cultural practices – Propagation should be done using cuttings from healthy stock plants. Efforts to reduce the relative humidity in the greenhouse and increasing plant spacing to improve air circulation will suppress powdery mildew. Placing plants in sunny locations and avoiding overhead sprinkler irrigation will also help to reduce the incidence of disease. Growers should remove infected plants as well as remove debris from previous crops as soon as possible to reduce the amount of inoculum and infection source. • Chemical – Fungicides labeled for use against powdery mildew on hydrangea are available. Applications should be initiated when initial symptoms of the disease are observed or conditions are favorable for disease development. Fungicide applications should be repeated at 7–14-day intervals or by following label recommendations to provide effective control. Some labeled fungicides reported to control powdery mildew on hydrangea include azoxystrobin, fenarimol, paraffinic oil, and thiophanate-methyl (Hagan et al. 2004; Hagan and Mullen 2001). Fungicide rotations should be practiced to minimize the development of fungicide resistance. Rotations should include fungicides from different fungicide groups (FRAC groups) that operate using different modes of action. • Biorationals – Applications of biorational products are also effective in controlling powdery mildew (Mmbaga and Sauvé 2004). However products have to be applied preventatively when initial symptoms of powdery mildew are observed. Thorough coverage of the tops and undersides of leaves is recommended. Sodium bicarbonate (baking soda) marketed as Armicarb™ (potassium bicarbonate (MilStop)) and fatty acids from neem seed oil marketed as Triact 701 are labeled for powdery mildew control (Hagan et al. 2005; Horst et al. 1992; Mmbaga and Oliver 2007).

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Fig. 1 The early symptom of powdery mildew of hydrangea with white mildew and brown patches (Photo by Yonghao Li)

Fig. 2 Symptoms of hydrangea powdery mildew with infected leaves covered by heavy white mildew (Photo by Yonghao Li)

• Biocontrols –Bacillus subtilis strain QST 713 (Hagan et al. 2004) is labeled as a biological control agent against powdery mildew in hydrangea. • Resistance – The best way to manage powdery mildew is by growing diseaseresistant selections. Previous reports indicate that cultivars belonging to H. macrophylla ssp. serrata are more resistant than H. macrophylla ssp. macrophylla cultivars (Dirr 2004). Out of 90 H. macrophylla cultivars evaluated for resistance to powdery mildew over a 3-year period, three cultivars, “Amagi Amacha,” “Shirofuji,” and “Veitchii,” were among the most powdery mildewresistant cultivars each year (Windham et al. 2011). Other cultivars, namely, “Diadem,” “Komachi,” and “Omacha,” were highly resistant in 2006 and 2008, but only moderately resistant in 2007 (Windham et al. 2011). Bigleaf hydrangea “Veitchii” also showed higher resistance to powdery mildew in vitro tests (Li et al. 2009b).

2.2

Cercospora Leaf Spot (Cercospora hydrangea L.)

Geographic occurrence and impact. This disease generally affects bigleaf and smooth types of hydrangea as well as oakleaf hydrangea in both landscape and

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nursery settings (Vann 2010; Mmbaga et al. 2012, 2015). The disease is particularly important in warmer regions of southeastern United States including Arkansas, Alabama, Georgia, and Tennessee (Vann 2010; Hagan and Mullen 2001). This leafspot disease hardly ever kills the plant, but it can cause significant premature defoliation and reduce the plant vigor and flower bud set (Vann 2010; Hagan and Mullen 2001). Symptoms/signs. Leaf spot symptoms differ based on the type of hydrangea infected. Older leaves at the bottom of the plant usually exhibit the first visible symptoms and then spread upward toward the top of the plant (Vann 2010; Hagan and Mullen 2001). Early spots on the leaves are purple and small with a circular shape (Fig. 3). This is followed by enlargement of the spot that turns angular or irregular in shape and acquires a tan or gray center enclosed by a purple or brown border commonly referred to as frogeye leaf spot pattern in bigleaf hydrangea (Vann 2010; Hagan and Mullen 2001). Alternatively, leaf spots on oakleaf hydrangea are angular shaped and dark brown to purple and may become yellow green and defoliate. Leaf spotting often commences in the midsummer and becomes visible by early fall. Biology and epidemiology. The primary source of inoculum is the conidia of C. hydrangeae that overwinter in the leaf debris. The spores are disseminated to healthy lower foliage via splashing rain or overhead irrigation as well as wind (Hagan and Mullen 2001; Vann 2010). Recurring summer rain showers can significantly accelerate the rate of disease spread, level of spotting, and defoliation. On the other hand, suppression of the disease development and spread can be prolonged by drought periods and prolonged exposure to full sun (Hagan and Mullen 2001; Li et al. 2008; Mmbaga et al. 2012). Management • Cultural – Application of sufficient nitrogen will help ensure good growth of the plants (Hagan and Mullen 2001). Drip irrigation or using soaker hoses and wide plant spacing that allow air movement and reduce leaf wetness have been reported to reduce Cercospora leaf spot incidences in landscape plantings. Proper site selection is imperative in order to avoid known problematic areas. Choice of site by a grower may not always be feasible, but the use of well-drained areas and partial shade with direct exposure to morning sunlight promotes better growth of the hydrangea and reduces foliage disease incidence (Li et al. 2008). A good sanitation strategy requires the removal and destruction diseased leaf debris that act as the source of infection for new plants. This will help to minimize infection level and disease severity. • Chemical – Using protective fungicides is advocated for high-value landscape plants that annually show noticeable damages. The fungicide application should commence when first leaf spot symptoms are observed and continue as needed. Products containing chlorothalonil, myclobutanil, or thiophanate-methyl are highly effective if applied at the onset of the disease. The use of protective chlorothalonil or mancozeb programs was found to significantly reduce Cercospora leaf spot disease severity in bigleaf hydrangea (Morrison 1980).

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Fig. 3 Small circular brown necrotic lesions with ash-colored centers on a hydrangea leaf infected by Cercospora sp. (Photo by Margarete Mmbaga)

Since protection by these products is not guaranteed for new growth that is not covered by the fungicide, multiple fungicide applications at 10–14-day intervals are necessary to ensure continuous leaf protection and good disease control. • Resistant varieties – Selection of resistant and disease-free cultivars for new planting is important to prevent the introduction of the disease to new areas. Mmbaga et al. (2012) reported ten cultivars of H. macrophylla, “Ami Pasquier,” “Ayesha,” “Blue Bird,” “Forever Pink,” “Fuji Waterfall” (“Fujinotaki”), “Miyama-yae-Murasaki,” “Seafoam,” “Taube,” “Tricolor,” and “Veitchii,” for expressing resistance or moderate resistance to multiple pathogens including Cercospora leaf spot in full sun, full shade, and partial shade environments (Li et al. 2008; Mmbaga et al. 2012).

2.3

Botrytis Blight (Botrytis cinerea Pers.: Fr.)

Geographic occurrence and impact. Botrytis blight, also called gray mold, is a serious disease problem in the production and postharvest of floral crops. B. cinerea has a wide host range including many herbaceous annual and perennial plants (Plant Disease Diagnostic Clinic 2015). The fungus is considered an opportunistic pathogen that infects weakened or injured leaves and flowers especially in cloudy, humid, and rainy weather conditions. All species of hydrangea are susceptible to this disease, but bigleaf hydrangea is most susceptible (Hagan and Mullen 2001). Symptoms/signs. When flowers are infected, the initial symptoms are watersoaked spots on petals, which develop reddish-brown irregular blotches that may, in time, cover the whole flower later. Affected flowers quickly turn brown and wither. The fuzzy gray growth (spores and conidiophores of the fungal pathogen), a characteristic symptom of Botrytis blight, is usually noticed on diseased flowers when environment conditions are wet or highly humid (Hagan and Mullen 2001). Leaves can be infected and show irregular brown necrotic lesions on leaves when diseased petals or other infected debris are fallen on them (Fig. 4). Biology and epidemiology. B. cinerea can survive in plant debris as a saprophyte or as a pathogen on a broad range of host plants (Daughtrey et al. 2000). The fungus also overwinters as black resting structures (sclerotia) on dead plant tissues or in soil

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Fig. 4 Irregular necrotic lesions on hydrangea leaves affected by Botrytis blight (Photo by Margarete Mmbaga)

(Hagan and Mullen 2001; Plant Disease Diagnostic Clinic 2015). The pathogen can be transmitted by wind, water splash, insects, and human activities from affected plants. Spore germination needs high relative humidity or water films on plant surfaces. A 5- to 8-h wet period is required for infection of B. cinerea (Jewett and Jarvis 2001). Germinated spores directly penetrate healthy blossoms and leaves and also enter through natural openings and wounds. The favorable conditions for the disease development are unbalanced nutrition, low light intensity, cool temperature, and high humidity, especially in the condition of several consecutive days of cloudy, humid, rainy weather (Sinclair and Lyon 2005). The optimal temperature for the onset of Botrytis blight in the greenhouse is approximately 15  C. All species of hydrangea are susceptible to this disease, but damage is noted most often on the bigleaf hydrangea (Hagan and Mullen 2001). Management • Cultural practices – Venting and concurrently heating greenhouses at dusk can expunge humidity. Avoiding overhead irrigation will help to keep leaf canopies dry and reduce relative humidity which can suppress the disease (Daughtrey et al. 2000). Increasing plant spacing and pruning weakened leaves from the lower part of plants to improve air circulation and encourage rapid evaporation of moist from plants and soil surfaces, avoiding wounds of leaves or shoots which can promote further spread of the disease. In addition, watering plants in the morning to allow rapid evaporation of excess moisture from plant surfaces is recommended. • Sanitation – Proper sanitation is crucial in controlling this disease. But, sanitation should not be conducted when plants are wet with dew or rain because it will spread fungal spores into the air. Removing infected leaves and damaged flower heads from plants and destroying them to reduce the inoculum in greenhouses and landscapes. In the greenhouse, clearing of benches and beds should be done before bringing in fresh plant materials of next crops. This can be followed by

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treatment of wooden surfaces in propagation and production areas with 2% disinfectant (Hagan and Mullen 2001). • Fungicide – Protection of plants against infection can be easily managed using fungicides. The first fungicide application should be made in the spring when continuously cool and wet weather is predicted or where Botrytis blight has been a problem in the previous year. Choice of fungicides is based on the site and type of plant(s) to be treated. Some effective fungicides recommended include iprodione, mancozeb, and thiophanate-methyl (Hagan and Mullen 2001). To prevent fungicide resistance development, always alternate fungicide applications between chemicals with different modes of actions. Resistance to thiophanatemethyl and iprodione has been reported in Botrytis populations. • Biological – Some biofungicides, such as Mycostop (Streptomyces griseoviridis Strain K61), actinovate (Streptomyces lydicus), are effective to control Botrytis blight. • Resistant varieties – The use of resistant cultivars is the most effective; however, no major resistance gene to Botrytis has been identified in hydrangeas.

2.4

Rust (Pucciniastrum hydrangeae (B. & C.) Arth.)

Geographic occurrence and impact. In nurseries, gardens, and landscapes, hydrangea rust is most commonly found on smooth hydrangea, a native shrub in the eastern North America. The pathogen can also affect hemlock, but it is more commonly found on hydrangeas. The disease is reported from New York to Illinois, Arkansas, Tennessee, and Georgia (Sinclair and Lyon 2005; Li et al. 2010). Symptoms/signs. On hydrangea leaves, symptoms initiate as small yellow or orange spots, which then develops to necrotic angular lesions because fungal growth is normally restricted by major veins (Fig. 5). An orange dust (urediniospores) forms on the lower surface of infected leaves when the fungal fruiting bodies (uredinia) are mature (Fig. 6). As the disease develops, symptoms are noticeable on the whole plant (Fig. 7). Biology and epidemiology. To complete its life cycle, the fungus, P. hydrangea, needs two genetically distinctive hosts, hemlock and hydrangea, which includes Tsuga canadensis (eastern hemlock), T. caroliniana (Carolina hemlock), H. arborescens (smooth hydrangea), and H. paniculata (panicle hydrangea). Urediniospores formed on hydrangea leaves repeat several infection cycles on hydrangeas during a season. Flat, reddish-brown telia develop within the epidermal cells of both sides of leaf surfaces of hydrangea in late summer and fall. The teliospores germinate and produce basidiospores that infect hemlock needles in the spring (Sinclair and Lyon 2005). In late spring to early summer, cylindric aecia with light cream color are formed on the lower surface of hemlock needles. When aecia are mature, orange-yellow aeciospores are released and infect hydrangea leaves in the spring.

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Fig. 5 Yellow spots on a smooth hydrangea leaf infected with hydrangea rust (Photo by Yonghao Li)

Fig. 6 Yellow to orange pustules on the lower surface of a hydrangea leaf infected with hydrangea rust (Photo by Yonghao Li)

Management • Cultural practices – Reduction of relative humidity and the duration of plant tissue wetness are critical for the disease control. • Fungicide – Chlorothalonil 720 SC and Daconil Weatherstik fungicides are labeled for hydrangea rust. The first fungicide application should be made in the spring to protect leaves. Fungicide doses and replications need to follow the fungicide labels. • Resistance – Significant differences in resistance to hydrangea rust were reported between cultivars of smooth hydrangeas (Li et al. 2010). Among seven tested varieties, “Frosty” was highly resistant, “Green Dragon” was highly susceptible, and the other five varieties, “Ryan Gainey,” “Pink Pincushion,” “White Dome,” “Mayes Starbust,” and “Annabelle,” were intermediate (Li et al. 2010).

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Fig. 7 A hydrangea plant with rust-infected leaves (Photo by Yonghao Li)

2.5

Anthracnose (Colletotrichum gloeosporioides (Penz.) Penz. and Sacc.)

Geographic occurrence and impact. Anthracnose of hydrangea is caused by C. gloeosporioides and found periodically in landscape and field plantings of bigleaf hydrangea. This disease affects both leaves and blossoms of hydrangea plants. Symptoms/signs. Initially, circular or slightly irregular brown spots are formed on hydrangea leaves with the center of the spots measuring up to 2.5 cm or more in diameter and turning light brown to tan in color (Hagan and Mullen 2001). This is followed by alternating dark and slightly lighter rings of dead tissue forming a bull’s eye or target-spot appearance (Fig. 8). Large, dark brown, irregular blotches may extend across the flower petals and leaves when environmental conditions are favorable for the disease development. Biology and epidemiology. C. gloeosporioides has a very broad host range in woody shrubs and trees. The pathogen overwinters in diseased leaf debris of hydrangea and other plant debris. The disease is favored by hot, wet weather conditions. Masses of spores ooze from fruiting bodies (acervuli) embedded in leaf debris. In the presence of continuous leaf wetness, spores spread to other leaves and flower petals by splashing water (Hagan and Mullen 2001). The pathogen is able to quickly penetrate and colonize the host tissues at warm temperatures of 24–32  C. The rate of disease infection and symptom appearance are promoted by prolonged periods of heavy fog, frequent showers, and dew. Hydrangeas that are extensively fertilized may be more sensitive to infection by C. gloeosporioides (Hagan and Mullen 2001). Management • Cultural – The use of disease-/symptom-free plant cuttings can significantly minimize the chances of spreading the disease. Removing and destroying diseased leaf debris or blighted blooms eliminate and reduce the inoculum source. • Chemicals – The use of protective fungicide treatments is a viable option to protect susceptible hydrangea from anthracnose. The application should be done

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Fig. 8 Circular to irregularshaped brown spots with concentric rings and ash-gray center on hydrangea leaves infected by Colletotrichum gloeosporioides (Photo by Margarete Mmbaga)

Fig. 9 Various sized brown spots on hydrangea leaves infected with Corynespora cassiicola (Photo by Margarete Mmbaga)

at 10–14-day intervals during the summer for good efficacy. Some of the recommended fungicides include chlorothalonil and thiophanate-methyl, and they should be administered as per manufacturer’s instructions (Hagan and Mullen 2001).

2.6

Corynespora Leaf Spot (Corynespora cassiicola (Berk. & M.A. Curtis) C.T. Wei)

Geographic occurrence and impact. This disease infects leaves, stems, and flower petals and causes irregularly shaped brown necrotic lesions. The disease may spread rapidly during hot and wet weather and has the potential to kill small plants and severely reduce the aesthetic value of the plants. Symptoms/signs. Symptoms of this disease are characterized as various sized brown spots on hydrangea leaves (Fig. 9) that are similar to those of Cercospora leaf spot. Like Cercospora leaf spot, Corynespora leaf spot is favored by the similar environments and is often isolated as part of a disease complex (Hagan et al. 2004; Zaher et al. 2005; Smith and Schlub 2007).

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Biology and epidemiology. Although C. cassiicola has been reported on over 300 plant species, a high degree of host specificity was reported in the fungal populations (Pereira et al. 2003). The pathogen survives in infested plant debris and on stems of previously infected plants. Management • Cultural – Removal and destruction of diseased leaf debris that act as the source of infection for new plants are important. Avoid overhead irrigation and plant crowding. • Chemicals – Both Cercospora and Corynespora leaf spot diseases tend to have annual outbreaks when the pathogens have become established in an area (Hagan and Mullen 2001; Hagan et al. 2004; Zaher et al. 2005; Smith and Schlub 2007; Mmbaga et al. 2012). Thus, protective fungicide applications are advocated for high-value landscape plants that annually show noticeable damages. The fungicide application should commence when first leaf spot symptoms are observed and continue as needed. • Resistant varieties – Selection of resistant and disease-free cultivars for new planting is important to prevent the introduction of the disease to new areas. Cultivars “Ami Pasquier,” “Ayesha,” “Blue Bird,” “Forever Pink,” “Fuji Waterfall” (“Fujinotaki”), “Miyama-yae-Murasaki,” “Seafoam,” “Taube,” “Tricolor,” and “Veitchii” were rated resistant or moderate resistant to multiple pathogens including Corynespora leaf spot in full sun (Mmbaga et al. 2012).

2.7

Phoma Leaf Spot (Phoma spp.)

Geographic occurrence and impact. This disease can occasionally cause leaf spots in hydrangea. Phoma exigua has been reported to cause leafspot disease in hydrangea in different parts of the United States and central Italy (Garibaldi et al. 2006; Mmbaga et al. 2009). Symptoms/signs. Phoma leaf spot is characterized by small necrotic spots surrounded by chlorotic haloes on the upper side of infected leaves (Garibaldi et al. 2006). The color of the leaf spots may differ slightly based on cultivars as observed by Mmbaga et al. (2010) where H. macrophylla cultivar “Lady in Red” developed small reddish-brown lesions and cultivar “Seafoam” had large lighter brown lesions during pathogenicity tests with P. exigua (Fig. 10). Biology and epidemiology. Phoma leaf spot is favored by relatively low temperatures (24  3  C) for symptom development (Mmbaga et al. 2010). Severely infected leaves become chlorotic and abscised leading to premature defoliation. Infected plants rarely die, but the presence of lesions on mature plants decreases aesthetic quality and subsequent market values. Management • Cultural practices – Sanitation and removal of diseased leaf debris help to eliminate source of infection.

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Fig. 10 Brown spots on a hydrangea leaf infected by Phoma exigua (Photo by Margarete Mmbaga)

2.8

Myrothecium Leaf Spot and Blight (Myrothecium roridum Tode Ex Fr.)

Geographic occurrence and impact. The disease was first reported in Tennessee in 2010 (Mmbaga et al. 2010; Mmbaga 2010). Symptoms/signs. In H. macrophylla, brown necrotic lesions with concentric rings and ash-colored centers (Fig. 11) characterized this disease and are similar to symptoms from M. roridum in other ornamental plants such as salvia, gardenia, begonia, and New Guinea impatiens (Mmbaga et al. 2010; Mangandi et al. 2007). The pathogen causes blighting of the stems, mostly beginning at the soil line, tan concentric leaf spots, and yellowing and wilting of foliage. Biology and epidemiology. M. roridum resides mostly in the soil and has a broad host range consisting of several vegetable, agronomic, and ornamental crops (Ponappa 1970). Infected plant materials and infested soil disseminate spores of M. roridum, and fungal spores may remain viable for 2–3 months in field soil at 20  C (Murakami et al. 2000). Management • Cultural practices – Some of the management strategies comprise of soil disinfestation using fumigants or steam and soil solarization to kill the pathogen in the soil. Preventative measures focus on sanitation practices to reduce disease incidence. • Host resistance – When available, disease resistance is the best method for controlling hydrangea leaf spot diseases. Out of 88 cultivars of H. macrophylla exposed to natural airborne leaf spot pathogens in McMinnville, Tennessee, USA, ten cultivars, “Ami Pasquier,” “Ayesha,” “Blue Bird,” “Forever Pink,” “Fuji Waterfall” (“Fujinotaki”), “Miyama-yae-Murasaki,” “Seafoam,” “Taube,” “Tricolor,” and “Veitchii,” were rated resistant or moderately resistant in full shade, full sun, and partial shade environments (Mmbaga et al. 2012). These cultivars may be considered to have multiple resistance to the pathogens including Cercospora spp., C. cassicola, C. gloeosporioides, M. roridum, and P. exigua (Mmbaga et al. 2012, 2015).

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Fig. 11 Concentric ring spots on a bigleaf hydrangea leaf infected by Myrothecium roridum (Photo by Margarete Mmbaga)

2.9

Alternaria Leaf Spot (Alternaria spp.)

Geographic occurrence and impact. Alternaria leaf spot of hydrangea can be caused by several Alternaria species that has been reported in Italy and the United States (Daughtrey 1995; Garibaldi et al. 2006, 2008). Infected plants rarely die, but the presence of lesions reduces the aesthetic quality and subsequently the commercial value. Symptoms/signs. Initial symptoms of Alternaria leaf spot are small brown spots with yellow halos that appear on the upper surface of the infected leaves. And then, lesions become dark brown or black and often coalesced into large necrotic areas. Symptoms are commonly observed on the leaf margins and near the petioles. Severely affected plants are defoliated (Garibaldi et al. 2006, 2008). Biology and epidemiology. Two species, A. alternate and A. compacta, have been identified on bigleaf hydrangea and climbing hydrangea, respectively, in Italy (Garibaldi et al. 2006, 2008). The pathogens of Alternaria leaf spot can survive as mycelium or spores in plant debris. In moist and wet conditions, spores of the pathogen are spread by wind, rain, or irrigation, which causes disease epidemics in landscapes and nurseries. Consequently, periods of moist conditions from rain and overhead irrigation are favorable for the disease development. Management • Cultural practices – Spacing plants to minimize dense canopy formation and improve air circulation. Avoiding overhead irrigation, watering plants in the morning to allow water on leaf surfaces to be evaporated quickly. Appropriately heating and ventilating greenhouses to reduce humidity. • Sanitation – Removing infected leaves from infected plants and destroying them or putting them in trash. • Fungicides – Fungicide application is a part of integrated disease management programs of the disease. Since most fungicides are preventative, the first application should be started when the disease is detected in nurseries or landscapes. The interval of applications needs to follow label recommendations.

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Bacterial Diseases

3.1

Bacterial Leaf Spot (Xanthomonas campestris)

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Geographic occurrence and impact. Bacterial leaf spot caused by a pathovar of X. campestris is a common problem on oakleaf hydrangea, smooth hydrangea, and bigleaf hydrangea. The disease has been reported in the United States (Uddin et al. 1996). Symptoms/signs. On oakleaf hydrangea, the early symptoms of the disease initiate as water-soaked spots on the lower surface of leaves and then develop to angular purple lesions because lesions are usually restricted by leaf veins (Fig. 12). Normally, the disease starts on the lower leaves and spreads upward on the plant. On bigleaf hydrangea and smooth hydrangea cultivars, symptoms are small purple angular spots (Fig. 13). At the late stage of lesion development, the necrosis develops in the center of lesions which dries, turns tan to brown, and can frequently fall out giving a shot-hole appearance. Biology and epidemiology. Humid and moderately wet conditions are favorable for the disease epidemic in greenhouses, nurseries, and landscapes. Infection begins on the lower leaves and then spreads to upper leaves and adjacent plants as the pathogen is transported by splashing water. Small water-soaked spots develop on leaves within 3–5 days after inoculation, with typical purple spots developing within 7 days (Uddin et al. 1996). Management • Cultural practices – Avoid overhead irrigation to prevent the spread of the inoculum through splashing water. Space plants adequately to improve air circulation. Water plants in the morning and avoid handling plant when they are wet. • Sanitation – Promptly remove and discard severely infected leaves during the growing season. At the end of a season, remove and destroy fallen leaves. • Fungicide and biological control – Kaolin clay (Surround) and neem oil (Neem Gold) were reported to reduce disease severities of bacterial leaf spot in the field experiments (Mmbaga and Oliver 2007). Copper-containing fungicides may provide some control when applied in late spring, but will not be effective in plants that are grown under conditions favoring the disease.

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Viral Diseases

4.1

Hydrangea Mosaic Virus (HdMV)

The typical symptom of HdMV on bigleaf hydrangea is chlorotic mosaic on leaves. The virus can be transmitted mechanically by grafting, by leaf contact, or by tools, but not by aphids or through seed. Control of HdMV includes using virus-free transplants and making cuttings from healthy plants (Thomas 1983).

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Fig. 12 Angular brown lesions of bacterial leaf spot on an oakleaf hydrangea leaf (Photo by Yonghao Li)

Fig. 13 Angular brown lesions of bacterial leaf spot on a bigleaf hydrangea leaf (Photo by Yonghao Li)

4.2

Hydrangea Ringspot Virus (HRSV)

The early symptoms of HRSV on hydrangea leaves are brown spots or rings, and then plants show leaf distortion, leaf roll, and stunted growth. The virus can be transmitted mechanically through saps, but aphids or seed does not transmit HRSV (Daughtrey et al. 1995; Williams-Woodward and Daughtrey 2001). Cultural control of HRSV includes buying clean stock and sanitizing pruning tools.

4.3

Tomato Ringspot Virus (ToRSV)

ToRSV has a wide host range including woody and herbaceous plants. On hydrangea, the virus causes leaf distortion and chlorosis and stunted plant growth. The virus can be transmitted by nematodes, but not by pruning tools. Control strategies for ToRSV are weed management and the use of nematode-free soil mix for containergrown plants (Williams-Woodward and Daughtrey 2001; Brierley 1954; Daughtrey et al. 1995).

Diseases of Hydrangea

4.4

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Tobacco Ringspot Virus (TRSV)

TRSV is widespread in woody and herbaceous plants in North America and also occurs in Europe, Asia, and Australia. Various symptoms can be found on infected plants, which include leaf distortion, leaf chlorosis, and stunted plant growth. Nematodes in the genus Xiphinema are major vectors of the virus (Williams-Woodward and Daughtrey 2001). Strategies for TRSV control include planting clean stocks in containers in soilless media and controlling weeds that may be reservoirs the virus and nematode vectors.

4.5

Tomato Spotted Wilt Virus (TSWV)

TSWV infects over 1,000 plant species. The virus is transmitted by thrips. Prevention is the first step of the disease control, which can be achieved by purchasing virus-free plant materials. In greenhouses, control of TSWV can be achieved by destroying infected plants and thrip management (Williams-Woodward and Daughtrey 2001).

4.6

Cherry Leaf Roll Virus (CLRV)

CLRV is widespread in Eurasia and North America and also occurs in Australia and New Zealand. Symptoms of CLRVon hydrangea are leaf deformation, chlorosis, and necrotic spots. This virus can be transmitted through seed and pollens and also by grafting (Veerakone et al. 2012). Since CLRV is a relatively new disease on hydrangea and no effective control methods have been reported, using clean and virus-free plant materials is effective to prevent the introduction of the virus to nurseries and landscapes.

4.7

Alfalfa Mosaic Virus (AMV)

AMV has a wider host range including 150 plant species and is widely distributed in the world (Jaspars and Bos 1980). This virus was first reported on Hydrangea spp. in British Columbia (Chiko et al. 1986). In the United States, occurrence of AMV in hydrangea was first reported in 2013 (Lockhart et al. 2013). A typical symptom of AMV on hydrangea is yellow leaf blotching. AMV can be transmitted by various aphid species. Control of AMV is achieved by weed management and aphid control (Daughtrey et al. 1995).

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References Brierley P (1954) Symptoms in the florists’ hydrangea caused by tomato ringspot virus and an unidentified sap-transmissible virus. Phytopathology 44:696–699 Chiko AW, Godkin SE (1986) Occurrence of alfalfa mosaic, hydrangea ringspot, and tobacco ringspot viruses in Hydrangea spp. in British Columbia. Plant Dis 70:541–544 Daughtrey ML, Wick RL, Peterson JL (1995) Compendium of flowering potted plant diseases. American Phytopathological Society, St. Paul, 90 pp Daughtrey ML, Wick RL, Peterson JL (2000) Botrytis blight of flowering potted plants. Plant Health Prog. doi:10.1094/PHP-2000-0605-01-HM Dirr MA (2004) Hydrangeas for American Gardens. Timber Press, Portland 236pp Garibaldi A, Bertetti D, Gullino ML (2008) First report of a leaf spot caused by Alternaria compacta on Hydrangea anomala subsp. petiolaris in Italy. Plant Dis 92:173 Garibaldi A, Gilardi G, Minerdi D, Gullino ML (2006) First report of leaf spot caused by Phoma exigua on Hydrangea macrophylla in Italy. Plant Dis 90:1113 Hagan AK, Mullen JM (2001) Diseases of hydrangea. ANR-1212. http://www.aces.edu/pubs/docs/ A/ANR-1212/ANR-1212.pdf. Verified 17 Jan 2017. Hagan AK, Olive JW, Stephenson J, Rivas-Davila ME (2004) Impact of application rate and interval on the control of powdery mildew and Cercospora leaf spot on big leaf hydrangea with azoxystrobin. J Environ Hortic 22:58–62 Hagan AK, Olive JW, Stephenson J, Rivas-Davila ME (2005) Control of powdery mildew and Cercospora leaf spot on bigleaf hydrangea with heritage and milstop fungicides. https://aurora. auburn.edu/bitstream/handle/11200/4098/BULL0658.pdf?sequence=1 Verified 17 Jan 2017. Horst RK, Kawamoto SO, Porter LL (1992) Effect of sodium bicarbonate and oils on the control of powdery mildew and black spot on roses. Plant Dis 76:247–251 Jaspars EM, Bos L (1980) Alfalfa mosaic virus. No 229. In: Descriptions of plant viruses. Commonwealth Mycology Institute/Association of Applied Biologists, Kew, Surrey, England Jewett T, Jarvis W (2001) Management of the greenhouse microclimate in relation to disease control: a review. Agronomie 21:351–366 Li YH, Windham MT, Trigiano RN, Reed SM, Spiers JM, Rinehart TA (2008) Effects of shading on Cercospora leaf spot in bigleaf hydrangea. Proc South Nurse Assoc Res Conf 53:379–380 Li YH, Windham MT, Trigiano RN, Reed SM, Spiers JM, Rinehart TA (2009a) Bright-field and fluorescence microscopic study of development of Erysiphe polygoni in susceptible and resistant bigleaf hydrangea. Plant Dis 93:130–134 Li YH, Windham MT, Trigiano RN, Reed SM, Rinehart TA, Spiers JM (2009b) Assessment of resistance components of bigleaf hydrangeas (Hydrangea macrophylla) to Erysiphe polygoni in vitro. Can J Plant Pathol 31:348–355 Li YH, Windham MT, Trigiano RN, Windham A, Reed SM, Rinehart TA, Spiers JM (2010) Evaluation of resistance to Pucciniastrum hydrangeae in Hydrangea arborescens. Phytopathology 100:S71 Lockhart B, Mollov D, Doughtrey M (2013) First report of Alfalfa mosaic virus occurrence in hydrangea in the United States. Plant Dis 97:1258 Mmbaga MT, Sauvé R (2004) Management of powdery mildew in flowering dogwood with biorational products and fungicides. Can J Plant Sci 84:837–844 Mmbaga MT, Oliver JB (2007) Effect of biopesticides on foliar diseases and Japanese beetle (Popillia japonica) adults in roses (Rosa spp.), oakleaf hydrangea (Hydrangea quercifolia) and crape myrtle (Lagerstroemia indica). Arboricult Urban For 33(3):1–10 Mmbaga MT, Reed SM, Windham MT, Li YH, Rinehart TA (2008) Disease resistance in commercial cultivars of Hydrangea macrophylla. Phytopathology 98:S108 Mmbaga MT, Windham MT, Li YH, Sauve’ RJ (2009) Fungi associated with naturally occurring leaf pots and leaf blights in Hydrangea macrophylla. Proc South Nurse Assoc Res Conf 54:49 Mmbaga MT (2010) Myrothecium roridum in garden Hydrangea. Proc South Nurse Assoc Res Conf 55:25–28

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Mmbaga MT, Li YH, Reed SM, Trigiano RN, Windham MT (2010) Phoma leaf spot in bigleaf hydrangea. Proc South Nurse Assoc Res Conf 55:21–25 Mmbaga MT, Kim MS, Mackasmiel L, Li YH (2012) Evaluation of Hydrangea macrophylla for resistance to leaf-spot diseases. J Phytopathol 160:88–97 Mmbaga MT, Kim MS, Mackasmiel L, Klopfenstein NB (2015) Differentiation of Corynespora cassiicola and Cercospora sp. in leaf-spot diseases of Hydrangea macrophylla using a PCR-mediated method. Can J Plant Sci 95:711–717. doi:10.4141/CJPS-2014-354 Morrison LS (1980) Control of leaf spot with foliar sprays (1979). Fungicide Nematicide Tests 35:119 Murakami R, Shirata A, Inoue H (2000) Survival and fluctuation in density of Myrothecium roridum in mulberry field soil. J Gen Plant Pathol 66:299–302 Park MJ, Cho SE, Park JH (2012) First report of powdery mildew caused by Oidium hortensiae on mophead hydrangea in Korea. Plant Dis 96:1072 Pereira JM, Barreto RW, Ellison CA, Maffia LA (2003) Corynespora cassiicola f. sp. lantanae: a potential biocontrol agent from Brazil for Lantana camara. Biol Control 26:21–31 Plant Disease Diagnostic Clinic (2015) Botrytis blight. Botrytis cinerea; Botrytis spp. http:// plantclinic.cornell.edu/factsheets/botrytisblight.pdf. Verified on 18 Jan 2017. Ponappa KM (1970) On the pathogenicity of Myrothecium roridum-Eichhornia crassipes isolate. Hyacinth Control J 8(1):18–20 Pscheidt JW (2014) Hydrangea powdery mildew. PNW plant disease management. https:// pnwhandbooks.org/plantdisease/host-disease/hydrangea-powdery-mildew. Verified on 18 Jan 2017 Rivera MC, Morisigue DE, Lopez SE (2004) Hydrangea macrophylla flower spot caused by Botrytis cinerea in Buenos Aires. Plant Dis 88:1160 Sinclair WA, Lyon HH (2005) Diseases of trees and shrubs, Second edn. Cornell University Press, Ithaca 660 pp Smith, LJ, Schlub, RL (2007) Diagnostic features of Corynespora cassiicola and its associated diseases. NPDN National Meeting Abstract 39. http://www.plantmanagementnetwork.org/ proceedings/npdn/2007/posters/39.asp. Verified on 17 Jan 2017 Thomas BJ (1983) Hydrangea mosaic virus, a new ilarvirus from Hydrangea macrophylla (Saxifragaceae). Ann Appl Biol 103:161–270 Uddin W, McCarter SM, Gitaitis RD (1996) First report of oakleaf hydrangea bacterial leaf spot caused by a pathovar of Xanthomonas campestris. Plant Dis 80:599 Van Gelderen CJ, Van Gelderen DM (2004) Encyclopedia of hydrangeas. Timber Press, Portland 279pp Vann S (2010) Cercospora leaf spot of hydrangea. University of Arkansas, FSA7570-PD-11-09N. https://www.uaex.edu/publications/pdf/FSA-7570.pdf. Verified on 17 Jan 2017 Veerakone S, Liefting LW, Lebas BSM, Ward L (2012) First report of cherry leaf roll virus in Hydrangea macrophylla. Plant Dis 96:463 Williams-Woodward JL, Daughtrey ML (2001) Hydrangea diseases. In: Jones RK, Benson DM (eds) Diseases of woody ornamentals and trees in nurseries, Second edn. APS Press, St Paul, pp 191–194 Windham MT, Reed SM, Mmbaga MT, Windham AS, Li YH, Rinehart TA (2011) Evaluation of powdery mildew resistance in Hydrangea macrophylla. J Environ Hort 29:60–64 Zaher EA, Hilal AA, Ibrahim I AM, Mohamed NT (2005) Leaf spots of ornamental foliage plants in Egypt with special reference to Corynespora cassiicola [(Berk. & Curt.) Wei] as a new causal agent. Egypt J Phytopathol 33:87–103

Diseases of Kalanchoe Robert L. Wick

Abstract

Kalanchoe blossfeldiana. Keywords

Erysiphe sedi • Golovinomyces orontii • Pseudoidium kalanchoes • Aecidium kalanchoe • Stemphylium solani • Stemphylium xanthosomatis • Stemphylium bolickii • Dickeya chrysanthemi • Pectobacterium carotovorum • Phytophthora niederhauserii • Phytophthora cactorum • Phytophthora nicotianae • Pythium

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungal-like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Cercospora Leaf Spot (Cercospora sp.). . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Leaf Spot and Scorch (Stemphylium bolickii Sobers & C.P. Seym., S. floridanum C.I. Hannon & G.F. Weber, S. solani G. F. Weber, and S. xanthosomatis B. Huguenin) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Phytophthora Root and Stem Rot [Phytophthora cactorum (Lebert & Cohn) J. Schrot], P. nicotianae (Breda de Haan), and P. niederhauserii Z.G. Abad et J. A. Abad . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Powdery Mildews [Podosphaera fuliginea (Schltdl.: Fr.) U. Braun & S. Takam.], (Erysiphe sedi U. Braun), [Pseudoidium kalanchoes (Lustner ex U. Braun) U. Braun & R.T.A. Cook, comb. nov.], and [Golovinomyces orontii (Castagne) Heluta, Ukrayins’k]) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Pythium Root and Stem Rot . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 Rust (Aecidium kalanchoe J. R. Hern., A. umbilici Trott.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7 Other Minor Diseases Caused by Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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R.L. Wick (*) Stockbridge School of Agriculture, University of Massachusetts, Amherst, MA, USA e-mail: [email protected] # Springer International Publishing AG (outside the USA) 2017 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florist's Crops Diseases, Handbook of Plant Disease Management, DOI 10.1007/978-3-319-32374-9_37-2

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3 Bacterial Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Bacterial Soft Rot and Wilt [Dickeya chrysanthemi (Burkholder et al.) Samson et al., Pectobacterium carotovorum subsp. carotovorum (Jones) Hauben et al. emend. Gardan et al.)]. Dickeya chrysanthemi and Pectobacterium carotovorum subsp. carotovorum formerly Erwinia chrysanthemi and Erwinia carotovora subsp. carotovora, Respectively . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Virus Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Kalanchoe Latent Virus (KLV) (Genus Carlavirus; Family Betaflexiviridae) . . . . . . . . 4.2 Kalanchoe Mosaic Virus (KMV) (Genus Potyvirus; Family Potyviridae Also Known as Green Island Mosaic) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Kalanchoe Top-Spotting Virus (KTSV) (Genus Badnavirus; Family Caulimoviridae) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4 Tomato Spotted Wilt Virus/Impatiens Necrotic Spot Virus (TSWV/INSV) (Genus Tospovirus; Family Bunyaviridae) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Introduction

The florists’ Kalanchoe, Kalanchoe blossfeldiana Poelln., is in the family Crassulaceae. A succulent perennial herb from Madagascar, it was first introduced to the floriculture trade in Germany in the 1930s. It is now widely propagated in Europe and North America and sold as a potted house plant. The genus also serves as the common name, but it also has many other common names.

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Fungal and Fungal-like Diseases

2.1

Cercospora Leaf Spot (Cercospora sp.).

Geographic occurrence and impact. Cercospora leaf spot was described in 1973 in Mississippi (Spencer 1973). It has been noted in Louisiana and California. Symptoms/signs. The single published description of Cercospora on Kalanchoe describes it as a severe purple-brown leaf spot, sunken when young and enlarging up to 3 mm. Older lesions become tan with dark margins and may have chlorotic halos; abscission may occur. Biology and Epidemiology. Cercospora spp. are known to survive well on plant debris between crops. They produce spores which are easily disseminated by wind currents. The Cercospora sp. from Kalanchoe grew best at 28 C/82 F at a pH of 5.5 in a liquid medium. Management. • Cultural Practices – Diseased plants and plant debris should be removed from the presence of healthy plants. Once the crop is finished, all crop debris should be removed. Avoid overhead watering and reduce leaf wetness duration as much as possible.

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• Chemical Control – The protective fungicide chlorothalonil and the single-site mode of action triazoles and strobilurins give very good protection against Cercospora diseases in field crops.

2.2

Leaf Spot and Scorch (Stemphylium bolickii Sobers & C.P. Seym., S. floridanum C.I. Hannon & G.F. Weber, S. solani G. F. Weber, and S. xanthosomatis B. Huguenin)

Geographic occurrence and impact. Stemphylium bolickii (originally spelled as bolicki) leaf spot was first described on Kalanchoe, Sedum, and Echeveria in 1963, USA; this was also the first description of the species epithet. Kalanchoe was described as a host of Stemphylium floridanum, 1965, USA. Stemphylium solani was first reported in Taiwan in 2011 and S. xanthosomatis in Korea in 2012. Stemphylium solani has been reported all over the world on a number of vegetable crops and ornamentals. However, S. solani isolated from tomato in Florida failed to infect Kalanchoe (Sobers and Seymour 1963). All four Stemphylium species are reported to be fairly destructive to Kalanchoe. Symptoms/signs. Leaf spots caused by Stemphylium bolickii and S. floridanum were compared and described by Sobers (Sobers and Seymour 1963; Sobers 1965). With S. floridanum, lesions enlarge rapidly and are circular to irregular, black, depressed, and up to 30 mm in diameter. S. bolickii causes lesions that are similar in shape, depressed at first but then become raised and scabby, rarely measuring more than 5 mm. Lesions caused by S. solani described in Taiwan measured up to 2 mm in diameter with a chlorotic halo (Shen et al. 2011). Stemphylium xanthosomatis is described as causing leaf scorch in Korea (Kwon 2012). Browning of the leaf margins is followed by interveinal chlorosis or darkening; finally leaves become desiccated and develop a velvety black olive growth of conidiophores and conidia. Defoliation also occurs. Biology and Epidemiology. Stemphylium survives in plant debris as mycelium and its thick-walled, darkly pigmented spores likely provide protection against UV light and desiccation. Optimum temperature for growth of S. floridanum isolated from Kalanchoe was 24–28 C/75–82 F, while optimum temperature for growth of S. bolickii was 28–32 C/82–90 F (Sobers 1965). Optimum temperature for growth of S. solani from Kalanchoe was reported to be 24–28 C (Shen et al. 2011). Conidia are produced on the surface of leaves and spread easily by air currents. Free moisture from dew, fog, rain, or irrigation is necessary for spore germination. Management. • Cultural Practices – Diseased plants and plant debris should be removed from the presence of healthy plants. Once the crop is finished, all crop debris should be removed. Avoid overhead watering and reduce leaf wetness duration as much as possible.

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• Chemical Control – Protective fungicides such as chlorothalonil and ethylene bisdithiocarbamates (EBDCs) have been shown to protect plants from infection from some species of Stemphylium.

2.3

Phytophthora Root and Stem Rot [Phytophthora cactorum (Lebert & Cohn) J. Schrot], P. nicotianae (Breda de Haan), and P. niederhauserii Z.G. Abad et J. A. Abad

Geographic occurrence and impact. Phytophthora causes many important diseases of floriculture crops, vegetables, and trees worldwide. All three of the above species are widely distributed. On Kalanchoe, P. cactorum was reported in Germany (Farr and Rossman 2015), P. nicotianae was reported in Korea (Farr and Rossman 2015), and P. niederhauserii was reported in Norway (Abad et al. 2014). Symptoms/signs. Root rot symptoms are indistinguishable from those caused by Pythium. Phytophthora is more likely to cause crown and stem rot than Pythium and causes a stem rot of Kalanchoe. Infected plants collapse and may have lesions moving up the stem. Biology and Epidemiology. Phytophthora is not as common in greenhouses as Pythium, but it is much more aggressive. There are more than 80 species of Phytophthora and most have wide host ranges, especially the three species reported on Kalanchoe. Like Pythium, Phytophthora is an oomycete and is most active when water is plentiful in the growing medium. Dispersal of inoculum is dependent on movement of contaminated water, soil, or plant material. Nearly all species produce oospores, but some need opposite mating types to do so, and some produce chlamydospores. Phytophthora is a soil-borne organism and can survive for many years in the absence of a host. Oospores are stimulated to germinate when root exudates are present, and germination often results in the formation of sporangia with swimming zoospores. Temperatures of 28 C/82 F and above favor P. nicotianae disease development (Daughtrey et al. 1995). Optimum temperature for growth of P. niederhauserii is 30 C/86 F (Abad et al. 2014). Optimum temperature for P. cactorum is 25 C/77 F and the maximum is 31 C/88 F (Erwin and Ribeiro 1991). Management. • Cultural Practices – When growing plants in a soil-less medium, take care not to introduce field soil by hands, tools, and hose ends dropped on the floor. Use a well-drained medium and avoid overwatering and overfertilization. • Chemical Control - Fungicides specific for Phytophthora include mefenoxam, etridiazole, and fluopicolide. See comments listed under chemical control of Pythium.

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Fig. 1. Powdery mildew of Kalanchoe blossfeldiana (Photo courtesy of L. Pundt)

2.4

Powdery Mildews [Podosphaera fuliginea (Schltdl.: Fr.) U. Braun & S. Takam.], (Erysiphe sedi U. Braun), [Pseudoidium kalanchoes (Lustner ex U. Braun) U. Braun & R.T.A. Cook, comb. nov.], and [Golovinomyces orontii (Castagne) Heluta, Ukrayins’k])

Geographic occurrence and impact. The first report of powdery mildew on Kalanchoe was in Germany in 1935 followed by England in 1945 (Wheeler 1978). Powdery mildew of Kalanchoe has been reported from the USA, Sweden, Switzerland, England, France, Korea, Germany, Hungary, Norway, Bulgaria, and Venezuela. Symptoms/signs. Symptoms can be nearly undetectable on Kalanchoe, but typically powdery colonies with robust mycelial growth can occur and the entire plant may become covered with mildew. When colonies are not obvious, damage to the epidermis can still be evident as a corky or pitted surface (Fig. 1). Biology and Epidemiology. Powdery mildews are obligate parasites of plants and are unique from other plant pathogenic fungi in that the spores do not need free moisture to germinate and infect, and can be more destructive at a relatively low (50–70%) relative humidity. Germination and infection can be inhibited by free moisture (Agrios 2005). Powdery mildews can survive in the absence of the host via chasmothecia. Many powdery mildews have restricted host ranges. Podosphaera fuliginea has been reported on the Plantaginaceae and also on Kalanchoe by the synonyms Sphaerotheca fuliginea and S. humuli var. fuliginea (Braun and Cook). Pseudoidium kalanchoes is reported in the Crassulaceae and probably has a wider

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host range (Braun and Cook 2012). Golovinomyces orontii has been reported on about 30 plant families. Management • Cultural Practices – As noted earlier, reduction of humidity is not a practical method to control powder mildews. • Chemical Control – Many fungicides will control powdery mildew but nearly all the modern chemistries have a single-site mode of action with some systemic or redistribution potential. Demethylation inhibitors and related chemistry, quinone outside inhibitors, and benzimidazoles all have a single-site mode of action and also select for resistant powdery mildews (Hollomon and Wheeler 2002). Therefore, rotation of fungicides with different modes of action is important.

2.5

Pythium Root and Stem Rot

Geographic occurrence and impact. Pythium root and stem rot have been reported in Denmark and Florida, USA; however, it is presumed to occur worldwide. Pythium is one of the most common causes of root rot in greenhouses (Daughtrey et al. 1995) Symptoms/signs. Pythium causes feeder roots to become brown and soft rotted. Usually the cortex easily separates from the vascular system. Root rot causes wilting of the foliage. Occasionally a crown rot occurs, and Pythium can move up the stem for some distance. Biology and Epidemiology. Pythium is a widespread and common genus of the Oomycota (Daughtrey et al. 1995); nearly every handful of soil contains one or several species. Pythium is not usually an aggressive plant pathogen unless plants are growing in a soil-less medium, soluble salts are high, and moisture is abundant. There are more than 200 species of Pythium, which generally have a wide host range and vary in their pathogenicity and distribution. Pythium species are not strong soil competitors but can survive in soil for long periods of time via oospores. When soil moisture is abundant, root exudates cause oospores to germinate and produce a sporangium where 10–30 zoospores develop in an extra-sporangial vesicle. The zoospores find roots by chemotaxis, and they have flagella that propel them through the water. Once on the root surface, they infect, colonize, and kill the root cells and produce oospores. Young roots and seedlings are particularly susceptible to infection. Management. • Cultural Practices – Remove diseased plants from the greenhouse being careful not to spill the soil from the pots. When growing plants in soil-less media, take care not to introduce field soil by hands, tools, and hose ends dropped on the floor. Use a well-drained growing medium and avoid overwatering and overfertilization. • Chemical Control – Many modern fungicides have a single-site mode of action and readily select for resistance. To avoid resistance development, rotate between

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different FRAC groups (FRAC Code List 2015); do not use one fungicide repeatedly. Note that some isolates of Pythium and Phytophthora may already be resistant to some of these fungicides. Many greenhouse isolates of Pythium in the USA have developed resistance to mefenoxam. The fungicides listed here are followed by the FRAC code in parentheses followed by the risk of selecting for resistance: mefenoxam (4), high; etridiazole (14), low to medium; azoxystrobin (11), high; fluopicolide (43), unknown; fosetyl-al (33), low; fenamidone (11), high; and cyazofamid (21) medium to high.

2.6

Rust (Aecidium kalanchoe J. R. Hern., A. umbilici Trott.)

Geographic occurrence and impact. Aecidium kalanchoe has only been reported in the USA (Idaho) and Colombia. The fungus was reported to cause defoliation in Kalanchoe blossfeldiana cvs. Sofie and Colbuco (Hernadez et al. 2004). Symptoms/signs. Chlorotic lesions composed of spermagonia surrounded by aecia, 1–1.5 cm in diameter form on the leaves (Fig. 2). Defoliation may occur. Biology and Epidemiology. This rust produces only spermagonia with spermatia and aecia with aeciospores; uredinia and telia are unknown. An alternate host is unknown or may not occur. Management. • Cultural Practices – Diseased plants and plant debris should be removed from the presence of healthy plants. Reduce humidity and leaf wetness duration. • Chemical Control – Ethylene bisdithiocarbamates and chlorothalonil provide broad-spectrum protection against rust. These fungicides have a multisite mode of action and will not select for resistance. Single-site mode of action sterol inhibiting fungicides and strobilurins will provide some curative activity and have longer application intervals. Fig. 2 Aecia of Aecidium kalanchoe on Kalanchoe blossfeldiana (Photo courtesy of J. Hernandez)

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Other Minor Diseases Caused by Fungi

Note that these diseases may be minor in importance or they are rare in occurrence. The following fungi are listed in various indices and reports as occurring on Kalanchoe blossfeldiana: Alternaria sp. Venezuela and West Indies; Botrytis cinerea, the USA, Bulgaria, and Korea; Cylindrocladium sp., Florida, USA; Fusarium herbarum, Germany; Fusarium oxysporum, Korea; Fusarium proliferatum, Korea; Glomerella cingulata, Venezuela, Lasiodiplodia theobromae, Venezuela; and Rhizoctonia solani, Florida, USA.

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Bacterial Diseases

3.1

Bacterial Soft Rot and Wilt [Dickeya chrysanthemi (Burkholder et al.) Samson et al., Pectobacterium carotovorum subsp. carotovorum (Jones) Hauben et al. emend. Gardan et al.)]. Dickeya chrysanthemi and Pectobacterium carotovorum subsp. carotovorum formerly Erwinia chrysanthemi and Erwinia carotovora subsp. carotovora, Respectively

Geographic occurrence and impact. Soft rot and wilt have been reported in Italy (Tamietti and Gullino 1988), the Netherlands (Janse and Ruissen 1988), Denmark (Dinesen 1979), and the USA (Engelhard et al. 1986); however, Dickeya and Pectobacterium have a worldwide distribution and occur on many plants and thus could occur wherever Kalanchoe is propagated. Bacterial soft rot is not a major problem, but it can be destructive and persistent if not controlled completely. Symptoms/Signs. Both Dickeya and Pectobacterium cause soft rot, stunting, and wilt of Kalanchoe blossfeldiana. Leaves may become grayish and lose their luster. Browning and soft rot of the pith and basal rot may be evident. Biology and Epidemiology. Dickeya and Pectobacterium produce pectic enzymes which are responsible for soft rot. These ubiquitous bacteria can survive in soil and may contact plants from dirty hands and tools, splashing water, insects, or hose ends that have contacted the ground. The bacteria may also come in on cuttings from infected mother plants. Bacteria need wounds or natural openings to gain entry into the host. Plants propagated vegetatively are very vulnerable to infection by soft rot bacteria, especially at high temperatures which are often provided by heating mats to encourage rooting. Pectobacterium carotovorum subsp. carotovorum has an optimum temperature for growth of 28–30 C/82–86 F but can grow at up to 37–42 C/98–108 F; optimum growth temperature of Dickeya chrysanthemi is 34–37 C/93–98 F, and some strains do grow above 40 C/104 F (Daughtrey et al. 1995). Management. • Cultural Practices – Do not take cuttings from plants that have evidence of bacterial infections. The cutting bench area should be clean and disinfested.

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Cuttings should be taken with a sharp knife, and knives should be disinfested frequently. Stick cuttings in growth medium right away or keep plants covered in a disinfested container for a minimum amount of time. If soft rotting occurs in the propagation area, discard all plant materials and thoroughly disinfest all surfaces with a quaternary ammonium compound or similar product (Daughtrey et al. 1995). • Chemical Control – Various formulations of copper can afford some protection against bacteria, but are not likely to prevent basal rots and wilts that initiate below the surface of the growing medium.

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Virus Diseases

4.1

Kalanchoe Latent Virus (KLV) (Genus Carlavirus; Family Betaflexiviridae)

Geographic occurrence and impact. USA and Denmark but likely to occur wherever Kalanchoe is propagated. Symptoms/Signs. This virus is symptomless in Kalanchoe. Biology and Epidemiology. Kalanchoe latent virus is a single-stranded RNA Carlavirus (Hearon 1982). Management. Meristem-tip culture can eliminate the virus.

4.2

Kalanchoe Mosaic Virus (KMV) (Genus Potyvirus; Family Potyviridae Also Known as Green Island Mosaic)

Geographic occurrence and impact. USA and Denmark but likely to occur wherever Kalanchoe is propagated. Symptoms/Signs. Mosaic and “green island” mosaic or a diffuse chlorotic mottle. Biology and Epidemiology. Pinwheel inclusions typical of Potyvirus were observed as is the case for other Potyviruses. KMV can be transmitted by aphids (Myzus persicae) in a non-propagative (nonpersistent) fashion. Symptoms occur 2–3 weeks. after aphid feeding (Husted et al. 1994). Management. Do not propagate from symptomatic plants. ELISA can be used to detect this virus, and meristem-tip culture can free plants of the virus (Bech and Husted 1996).

4.3

Kalanchoe Top-Spotting Virus (KTSV) (Genus Badnavirus; Family Caulimoviridae)

Geographic occurrence and impact. USA and Europe but probably occurs wherever Kalanchoe is propagated.

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Fig. 3 Top-spotting virus on Kalanchoe blossfeldiana (Photo courtesy of B. Lockhart)

Symptoms/Signs. Newly developing leaves show chlorotic spots which may become depressed as the symptoms fade (Fig. 3). Biology and Epidemiology. Ultrastructural studies showed bacilliform virus particles associated with top-spotting of Kalanchoe (Hearon 1981), and subsequent research confirmed the presence of a Badnavirus (Hearon and Locke 1984; Lockhart and Ferji 1988). The virus can be transmitted by vegetative cuttings, grafting, pollen, and seed (Hearon and Locke 1984). The virus can also be vectored by the mealy bug Planococcus citri (Brunt et al. 1996). The host range is considered to be very narrow, probably restricted to Kalanchoe. Management. Vegetative propagation is the most likely way this virus is spread. Do not propagate from infected plants. Also, K. blossfeldiana can be an asymptomatic carrier. PCR can detect the virus in plants, but unfortunately false positives can occur due to putative integrated pararetroviral elements of KTSV that are non-infective (Yang et al. 2005).

4.4

Tomato Spotted Wilt Virus/Impatiens Necrotic Spot Virus (TSWV/INSV) (Genus Tospovirus; Family Bunyaviridae)

Geographic occurrence and impact. The Tospoviruses TSWV and INSV have become distributed worldwide; the viruses are known to occur in the USA, South America, Canada, Europe, India, China, and Japan. Reports on Kalanchoe include USA, Canada (Tehrani et al. 1990), Finland, and Europe, but it is probably more widespread. TSWV and INSV are not destructive viruses on Kalanchoe but they are of economic importance as infected plants would be discarded. Symptoms/Signs. TSWV/INSV can cause distortion of new growth (Fig. 4); ring patterns may or may not occur (Fig. 5). Biology and Epidemiology. TSWV/INSV are Tospoviruses in the family Bunyaviridae. Prior to 1990, TSWV was considered to have two strains, TSWV I (impatiens strain), and TSWV L (lettuce strain) (German et al. 1992). In 1990, INSV

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Fig. 4 Distortion on Kalanchoe blossfeldiana foliage by a Tospovirus (Photo courtesy of R. L. Wick)

Fig. 5 Ringspots on Kalanchoe blossfeldiana foliage by a Tospovirus (Photo courtesy R. L. Wick)

was described as distinct from TSWV; however, prior to this time, plants showing symptoms of TSWV may in fact have been infected by INSV. At least in the USA, ornamental plants are far more likely to be infected by INSV than TSWV, and the symptoms can be similar (Daughtrey et al. 1997). Antisera are available to distinguish the two viruses. Tospoviruses are vectored exclusively by at least seven species of thrips (Thysanoptera: Thripidae). Only the first and early second stage larvae can acquire the virus, and both the larvae and adults can transmit the virus. The virus/thrip relationship is propagative (persistent) and INSV and TSWV can replicate in the vector. Management. • Cultural Practices – Inspect plants for viruses before introducing them into the greenhouse. Control weeds on the greenhouse floor and exterior. Monitor for thrips with yellow sticky cards. Screening can be used to cover greenhouse vents, but the material should have apertures of 135 μ or less; this will exclude thrips but

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can greatly reduce air flow into the greenhouse. Avoid carrying over crop material from season to season. • Chemical Control – Insecticides should be used to control thrips; however, many thrips have developed resistance to one or more insecticides. More information can be found in the introductory chapter “Insect Management for Disease Control in Florists’ Crops.”

References Abad ZG et al (2014) Phytophthora niederhauserii sp. nov. a polyphagous species associated with ornamentals, fruit trees and native plants in 13 countries. Mycologia 106:431–447 Agrios GN (2005) Plant pathology, 5th edn. New York, Elsevier Academic Press Bech K, Husted K (1996) Verification of ELISA as a reliable test method for Kalanchoe mosaic potyvirus (KMV) and establishment of virus-free Kalanchoe blossfeldiana by meristem-tip culture. Acta Hortic 432:298–305 Braun U, Cook RTA (2012) Taxonomic manual of the Erysiphales (powdery mildews). CBS-KNAW Fungal Biodiversity Centre, Utrecht Brunt AA, Crabtree K, Dallwitz MJ, Gibbs AJ, Watson L, Zurcher EJ (eds.) (1996 onwards) Plant viruses online: descriptions and lists from the VIDE database. Version: 20 Aug 1996 Daughtrey ML, Wick RL, Peterson JL (1995) Compendium of flowering potted plant diseases. APS Press, St. Paul Daughtrey ML, Jones RK, Moyer JW, Daub ME, Baker JR (1997) Tospoviruses strike the greenhouse industry; INSV has become a major pathogen in flower crops. Plant Dis 81:1220–1230 Dinesen IG (1979) A disease of Kalanchoe blossfeldiana caused by Erwinia chrysanthemi. Phytopathol Z 95:59–64 Engelhard AW, McGuire RG, Jones JB (1986) Erwinia carotovora pv. carotovora, a pathogen of Kalanchoe blossfeldiana. Plant Dis 70:575 Erwin DC, Ribeiro OK (1991) Phytophthora diseases worldwide. APS Press, St. Paul Farr DF, Rossman AY (n.d.) Fungal databases, systematic mycology and microbiology laboratory, ARS, USDA. Retrieved 18 Dec 2015, from http://nt.ars-grin.gov/fungaldatabases/ FRAC Code List# (2015) Fungicides sorted by mode of action. http://www.frac.info/docs/defaultsource/publications/frac-code-list/frac-code-list-2015-finalC2AD7AA36764.pdf?sfvrsn=4 German TL, Ullman DE, Moyer JW (1992) Tospoviruses: diagnosis, molecular biology, phylogeny, and vector relationships. Annu Rev Phytopathol 30:315–348 Hearon SS (1981) Detection of viruses in ultrastructural studies of naturally infected Kalanchoe blossfeldiana. (abstr.) Phytopathology 71:767 Hearon SS (1982) A carlavirus from Kalanchoe blossfeldiana. Phytopathology 72:838–844 Hearon SS, Locke JC (1984) Graft pollen and seed transmission of an agent associated with top spotting in Kalanchoe blossfeldiana. Plant Dis 68:347–350 Hernadez JR, Aime MC, Newbry B (2004) Aecidium kalanchoe sp. nov., a new rust on Kalanchoe blossfeldiana (Crassulaceae). Mycol Res 108:846–848 Hollomon DW, Wheeler I (2002) In: Belanger RR, Bushnell WR, Dik AJ, Carver LW (eds) The powdery mildews, a comprehensive treatise. APS Press, St. Paul, pp 249–255 Husted K, Bech K, Albrechtsen M, Borkhardt B (1994) Identification, partial sequencing, and detection of a potyvirus from Kalanchoe blossfeldiana. Phytopathology 84:161–166 Janse JD, Ruissen MA (1988) Characterization and classification of Erwinia chrysanthemi strains from several hosts in The Netherlands. Phytopathology 78:800–808 Kwon JH (2012) First report of Kalanchoe leaf scorch caused by Stemphylium xanthosomatis in Korea. Plant Dis 96:292

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Lockhart BEL, Ferji Z (1988) Purification and mechanical transmission of Kalanchoe top-spottingassociated virus. Acta Hortic 234:73–78 Shen YM, Yang YC, Fu YJ, Hung TH (2011) First report of Stemphylium solani causing leaf spot of Kalanchoe blossfeldiana in Taiwan. New Dis Rep 25:10. doi:10.5197/j.20440588.2012.025.010 Sobers EK (1965) A form of Stemphylium floridanum pathogenic to species of Kalanchoe. Phytopathology 55:1313–1316 Sobers EK, Seymour DP (1963) Stemphylium leafspot of Echeveria, Kalanchoe and Sedum. Phytopathology 53:1443–1446 Spencer JA (1973) A new Cercospora leaf spot of Kalanchoe. Phytopathol Abst 63:449 Tamietti G, Gullino G (1988) Una tracheobatteriosi della Kalanchoe causata da Erwinia chrysanthemi. Informatore Fitopatologico 3(88):73–75. (in Italian) Tehrani B, Allen WR, Matteoni JA (1990) Update on the incidence of tomato spotted wilt virus in greenhouses. Can Plant Dis Surv 70:102–103 Wheeler BEJ (1978) Powdery mildew of ornamentals. In: Spencer DM (ed) The powdery mildews. Academic, New York, pp 411–445 Yang Z, Nicolaisen M, Olszewski NE (2005) Sequencing, improved detection and a novel form of Kalanchoe top-spotting virus. Plant Dis 89:298–302

Diseases of Poinsettia Margery L. Daughtrey and A. R. Chase

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-Like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Alternaria Leaf Spot [Alternaria euphorbiicola E. Simmons and Engelhard, Alternaria euphorbiae (Barth) Aragaki and Uchida (comb. nov.)] . . . . . . . . . . . . . . . . . . 2.2 Amphobotrys Blight [Amphobotrys ricini (Buchw.) Hennebert (syn. Botryotinia ricini (Godfrey) Whetzel)] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Anthracnose [Colletotrichum truncatum (Schwein.) Andrus and W. D. Moore (syn. C. capsici (Sydow) E. J. Butler and Bisby)] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Botrytis Blight or Gray Mold [Botrytis cinerea (syn. Botryotinia fuckeliana (de Bary) Whetz)] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Choanephora Wet Rot [Choanephora cucurbitarum (Berk. and Rav.) Thaxter] . . . 2.6 Corynespora Leaf and Bract Spot [C. cassiicola (Berk. and M. A. Curtis) C. T. Wei (syn. Helminthosporium cassiicola Berk. and M. A. Curtis)] . . . . . . . . . . . . 2.7 Fusarium Stem Rot [Fusarium solani (Mart.) Sacco] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.8 Powdery Mildews: Leveillula clavata Nour, L. taurica (Lev.) G. Arnaud, Pseudoidium poinsettiae (U. Braun, Minnis, and Yañez-Morales), Phyllactinia poinsettiae, and Ovulariopsis erysiphoides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.9 Pythium Root Rot (Pythium spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.10 Phytophthora Crown and Root Rot (Phytophthora drechsleri, P. nicotianae, P. cryptogea) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.11 Rhizoctonia Cutting Rot, Crown Rot, and Root Rot [Rhizoctonia solani Kuhn (syn. Thanatephorus cucumeris (A. B. Frank) Donk)] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.12 Rhizopus Blight [R. stolonifer (Ehrenb.:Fr.) Vuill. (syn. Rhizopus nigricans Ehrenb.)] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.13 Scab (Spot Anthracnose) (Sphaceloma poinsettiae Jenk. and Ruehle) . . . . . . . . . . . . . .

2 3 3 5 6 6 10 11 12

12 17 22 23 26 28

M.L. Daughtrey (*) Section of Plant Pathology and Plant-Microbe Biology, Cornell University, Long Island Horticultural Research & Extension Center, Riverhead, NY, USA e-mail: [email protected] A.R. Chase Chase Agricultural Consulting LLC, Cottonwood, AZ, USA e-mail: [email protected] # Springer International Publishing AG 2016 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florist's Crops Diseases, Handbook of Plant Disease Management, DOI 10.1007/978-3-319-32374-9_39-1

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Thielaviopsis Black Root Rot: Thielaviopsis basicola (Berk. and Broome) Ferraris (syn. Chalara elegans) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31 3 Bacterial Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 35 3.1 Bacterial Canker [Curtobacterium flaccumfaciens pv. poinsettiae (Starr and Pirone) Collins and Jones (Previously Corynebacterium flaccumfaciens subsp. poinsettiae)] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 35 3.2 Greasy Canker [Pseudomonas viridiflava (Burkholder) Dowson] . . . . . . . . . . . . . . . . . . . . 36 3.3 Soft Rot and Stem Rot [Dickeya chrysanthemi (syn. Pectobacterium chrysanthemi) and Pectobacterium carotovorum (syn. Erwinia carotovora (Jones)) Holland.] . . . . . 37 3.4 Xanthomonas Leaf Spot [X. axonopodis pv. poinsettiicola (Patel et al. 1951) Vauterin, Hoste, Kersters, and Swings 1995 (syn. X. campestris pv. poinsettiicola)] . . . . . . . . . . . 38 3.5 Poinsettia Branch-Inducing Phytoplasma (Candidatus Phytoplasma sp.) . . . . . . . . . . . . 40 4 Viruses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 42 4.1 Poinsettia Mosaic (Poinsettia mosaic virus PnMV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 42 4.2 Poinsettia Latent Virus (PnLV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 43 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 43

Abstract

Poinsettia, Euphorbia pulcherrima Willd. ex. Klotsch, is a major flowering potted plant for winter holidays. Because it is vegetatively propagated and is so widely grown, poinsettia diseases are relatively well known and well studied. Being a woody plant, poinsettias are not prone to tospoviruses, but they are susceptible to a wealth of foliar problems (Botrytis blight, powdery mildew, scab, Alternaria leaf spot, Xanthomonas leaf spot, and anthracnose) as well as many common root and stem diseases (Pythium and Phytophthora root rots, Thielaviopsis root rot) as well as occasional Fusarium or Rhizoctonia stem problems. Poinsettia mosaic virus (PMV) has been associated with minor problems, and a phytoplasmal infection has contributed free branching for more attractive plants. Poinsettia diseases may be managed through clean stock production coupled with integrated pest management strategies in greenhouses where the crops are propagated and finished for sale. Keywords

Poinsettia • Euphorbia pulcherrima • IPM • Botrytis • Powdery mildew • Scab • Xanthomonas • Pythium • Poinsettia mosaic • Branch-inducing phytoplasma

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Introduction

Poinsettia, Euphorbia pulcherrima Willd. ex. Klotsch, is a woody plant used as an ornamental, especially as a potted plant for the winter holidays. The poinsettia is native to an area near Taxco, Mexico, where the Aztecs valued it for its red bract color and made use of it as a dye and for medicines (Ecke et al. 2004). Poinsettias were brought to the United States in 1825 by Joel Roberts Poinsett, the first US ambassador to Mexico. The poinsettias sold from the 1920s to 1960s were largely selections or sports from a single seedling called ‘oak leaf’ grown in New Jersey in

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the 1920s, many of them developed by Paul Ecke Sr. in Encinitas, California (Ecke et al. 2004). The plant was developed as an ornamental crop in the United States and Europe in the latter half of the twentieth century, through commercial, university, and federal breeding programs. Their success led to a diversity of bract colors and forms and attractive dark green foliage and branching habit (Ecke et al. 2004). Today the poinsettia is the leading potted flowering plant for indoor use, with a wholesale value of $141 million in 2014 (USDA 2015). Poinsettias are also used as landscape ornamentals in subtropical areas, including the southern United States. Poinsettias for holiday potted plants are produced from cuttings under protective glass or plastic structures in temperate climates during the summer and fall for late fall sales. Poinsettias are one of the most important flower crops, thus their diseases have received more attention than many. For some of the less common diseases, our knowledge extends only to identification, while for others research has been conducted on poinsettias or other hosts to unravel the secrets of their epidemiology and management (Benson et al. 2001; Strider and Jones 1985). Throughout this chapter, the numeric groupings assigned by the Fungicide Resistance Action Committee (FRAC) will be used to designate mode of action for fungicide active ingredients.

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Fungal and Fungus-Like Diseases

2.1

Alternaria Leaf Spot [Alternaria euphorbiicola E. Simmons and Engelhard, Alternaria euphorbiae (Barth) Aragaki and Uchida (comb. nov.)]

2.1.1 Geographic Occurrence and Impact Species of Alternaria have been reported worldwide on poinsettia (Table 1). The appearance of the disease is inconsistent, however, and its impact on poinsettia production is not high in most greenhouses. 2.1.2 Symptoms/Signs Alternaria leaf spot causes symptoms that are sometimes similar to other foliar diseases of poinsettia, including Phytophthora blight, bacterial canker, and scab. Lesions form on bracts, leaves, petioles, and stems. Purplish-black spots (initially 0.5 mm in diameter) grow to an elliptical shape (2 4  4 7 mm); eventually they become irregularly shaped and turn brown, reaching up to 20 mm in diameter (Fig. 1). Chlorotic halos often surround lesions (Fig. 2), with leaf abscission common in severe infections. Lesions on leaf veins lead to distortion. Tan to brown stem lesions (3  8 mm) are elongated and sunken. Infection of cyathia has been reported (Yoshimura et al. 1986). 2.1.3 Biology and Epidemiology In Florida and other subtropical regions that experience significant rainfall, the disease is most severe on poinsettia crops produced in shade houses or in the

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Table 1 Alternaria species reported on poinsettia worldwide (Farr and Rossman 2015) Alternaria species Alternaria alternata (syn. A. tenuis) Alternaria biproliformis (syn. A. heveae) Alternaria euphorbiae Alternaria euphoribiicola Alternaria obtecta Alternaria pipionipisi, A. rostellata A. pseudorostrata A. protenta (syn. A. pulcherrimae) Alternaria sp. Alternaria tenuissima

Fig. 1 Alternaria spots may occur on bracts as well as leaves

Fig. 2 Alternaria leaf spots often show yellow haloes

Location Japan, New Caledonia China Hawaii Florida, Hawaii, Hong Kong California, Louisiana California California, United States Australia Michigan, Tanzania, Venezuela Hong Kong

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landscape. The disease is rarely a problem in greenhouses, where the foliage can be kept dry through careful environmental regulation, especially with the advent of ebband-flood or flood-floor production, common for this crop today in many countries.

2.1.4 Management Cultural practices that eliminate or minimize leaf wetness help limit development of Alternaria leaf spot. Diseased plants and fallen leaf debris should be removed frequently from the growing area. Thiophanate-methyl should not be used for this disease as it has been shown to increase severity of Alternaria leaf spot on some floricultural crops. Iprodione (FRAC 2), triflumizole (FRAC 3), chlorothalonil (FRAC M5), and trifloxystrobin (FRAC 11) provided about 70 % control in Florida trials, but the best control was seen with fludioxonil (FRAC 12), azoxystrobin (FRAC 11), and mancozeb (FRAC M3) (McGovern 1999; McGovern and Wilfret 1998b; McGovern et al. 2001). Poinsettia cultivars that are relatively resistant to Alternaria leaf spot should be grown. Susceptibility varies among poinsettia cultivars; the only available reports are from cultivars available in the 1980s–1990s. Poinsettia cultivars that are relatively resistant to Alternaria leaf spot should be grown. The most susceptible cultivars included some V-14 cultivars (Glory, White, and Jingle Bells) and Eckespoint C-1 Red. V-10 Amy had intermediate susceptibility. Annette Hegg (Dark Red, Top White, Brilliant Diamond, and Hot Pink) developed only tiny leaf spots with tan centers in resistance trials (Engelhard and Schubert 1985). McGovern and Wilfret (1998a, 1999) reported that the poinsettia cvs. consistently least susceptible to Alternaria leaf spot included Freedom White, Pearl, Petoy, Sonora Pink, Spotlight Dark Red, and Subjibi.

2.2

Amphobotrys Blight [Amphobotrys ricini (Buchw.) Hennebert (syn. Botryotinia ricini (Godfrey) Whetzel)]

2.2.1 Geographic Occurrence and Impact Amphobotrys blight and stem rot on poinsettia were first reported from Louisiana and Florida in 1988 and were noted in Bermuda in 1995 (McGovern, personal communication). In 2000, the same pathogen was reported from Florida attacking another species of Euphorbia (E. milii) causing a flower blight (McMillan et al. 2000). This disease appears to be regional and very rare.

2.2.2 Symptoms/Signs A foliar blight and shoot blight occur; mycelial webbing forms rapidly. Basal stem rot characterized by a soft, watery rot starting at the soil line has also been reported. The rot results in wilting without extensive stem or root rot (Holcomb and Brown 1990). Small, black sclerotia have been found in infected poinsettia stems.

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2.2.3 Biology and Epidemiology Several species in the genus Euphorbia, as well as Ricinus communis (castor bean), are susceptible to this pathogen: E. supina Raf. ex Boiss., E. heterophylla L., and E. hirta L. as well as E. pulcherrima. 2.2.4 Management Shoot tip infections were controlled by used of a thermal fog formulation of chlorothalonil (FRAC M5) but this did not control rot at the stem base (Holcomb and Brown 1990).

2.3

Anthracnose [Colletotrichum truncatum (Schwein.) Andrus and W. D. Moore (syn. C. capsici (Sydow) E. J. Butler and Bisby)]

2.3.1 Geographic Occurrence and Impact Anthracnose on poinsettia caused by C. capsici (syn. C. truncatum) was reported from Florida and Japan (Sato et al. 2015). Additionally, C. gloeosporioides has been reported from Mexico, Florida, and Texas. Anthracnose on poinsettia is not common in the United States. 2.3.2 Symptoms/Signs Leaf and stem spot and blight on poinsettia cultivar Angelika were found in a greenhouse in Japan in 2004. Brown spots 3–5 mm in diameter appeared at first on lower leaves and then enlarged along veins or leaf margins. The spots developed pale brown centers with dark brown borders and dead areas eventually dropped off. Stems became brittle as elongated lesions developed on them. Black acervuli with setae were produced on the lesions under high humidity. Wounded and non-wounded poinsettia plants (cv. Angelika) were inoculated, but symptoms developed only on the wounded plants, 11–21 days after inoculation (Sato et al. 2008). 2.3.3 Management Utilize environmental control to minimize periods of wetness on leaves and bracts of poinsettias. Avoid wounding foliage or bracts, which might provide entry points for the fungus.

2.4

Botrytis Blight or Gray Mold [Botrytis cinerea (syn. Botryotinia fuckeliana (de Bary) Whetz)]

2.4.1 Geographic Occurrence and Impact Botrytis cinerea is a ubiquitous fungus. Botrytis blight occurs wherever poinsettias are grown and causes losses in propagation, finishing, and post-harvest stages.

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Fig. 3 Botrytis leaf spots often form wedges at the edge of the leaf

Fig. 4 Closeup of Botrytis lesion at leaf tip

2.4.2 Symptoms/Signs Botrytis blight is particularly feared as a disease of bracts, but may attack all aboveground portions of poinsettia plants: stems, leaves, bracts, and flowers. Leaf lesions are typically large, light to medium brown, and enlarge further with each period of high humidity, sometimes creating zonate lesions. Infections often start as small light brown lesions at the edge of bracts and grow into large wedges, having begun as infections at the perimeter of the leaf or bract (Figs. 3 and 4). Stem cankers are usually light brown to tan and may girdle main stems or side branches, causing wilt (Fig. 5). Flower parts may also be infected. The long brown conidiophores topped by clumps of hyaline spores form conspicuous masses on dead plant tissue to give the disease its common name: gray mold. 2.4.3 Biology and Epidemiology There are hundreds of host plants for Botrytis cinerea, but poinsettia is one of the most susceptible flower crops. The pathogen is a necrotroph, growing in moribund

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Fig. 5 Botrytis stem cankers are tan in color; affected portions of plants will wilt

tissue and secreting cell wall-degrading enzymes in advance of its hyphae. Its ability to infect plants from a morsel of infected tissue is 200 times greater than its ability to cause infection via conidia (Sirjusingh et al. 1996). Wounds in leaves are a common site of infection, especially for cuttings during shipment and propagation. Disease spread in the greenhouse is largely by dispersal of conidia by air currents (Jarvis 1980). The epidemiology of Botrytis populations has been more closely studied in crops other than poinsettia. Conidial germination is best at 20–25  C/ 68–77  F (Blakeman 1980) in the presence of free water (Strider and Jones 1985), although germination has also been noted at extremely high humidity under laboratory conditions. Germination may be delayed for at least 3 weeks (Salinas et al. 1989) when conditions are not sufficiently moist. Nutrients on the surface of plants are beneficial for infection, which may explain why wounded tissues and flower parts are more likely to be infected than intact leaves (Blakeman 1975). Applications of glucose at 0.1 M and 0.05 M KH2PO4 were found to increase germination and infection (Kulek and Floryszak-Wieczorek 2002). Older, more mature leaves are the most susceptible of the intact foliage (Elad and Evensen 1995). Latent infections by B. cinerea are one of the main reasons why the disease is so important as a post-harvest problem: symptoms often develop after poinsettias are packed and shipped (Jarvis 1977; Pritchard et al. 1996). Bract-edge burn in poinsettias, which develops as a result of calcium deficiency at the end of production in certain cultivars, can predispose bracts to Botrytis infection that destroys a plant’s salability (Barrett et al. 1995). Inverting the day/night temperature for a poinsettia crop so that night temperatures were warmer than day for the purpose of producing compact plants was shown not to affect poinsettias’ susceptibility to Botrytis blight (Pritchard et al. 1996). Higher finishing temperatures, either day or night, did increase blighting and sporulation.

2.4.4 Management Management of Botrytis blight in greenhouses requires an integrated strategy, as described by Hausbeck and Moorman (1996), comprised of sanitation,

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environmental control, and appropriate use of fungicides. The cutting wounds made on stock plants and the wounds on the cuttings themselves create critical control points for Botrytis blight management. Poinsettia susceptibility has been shown to increase with plant maturity (Pritchard et al. 1996), which increases the need for careful management just when growers are concerned with shipping. Injury to bracts quickly destroys crop value. Cultural and environmental controls are extremely important to avoid Botrytis losses in poinsettia culture. Organic debris removal from the greenhouse eliminates a source of conidial inoculum, as B. cinerea often sporulates profusely when living as a saprophyte on dead plant material. Poinsettias should be grown with adequate nitrogen and light to prevent senescence of lower leaves that could provide a substrate for Botrytis. Avoiding the presence of free moisture is largely accomplished by irrigation early in the day and nighttime humidity control. To prevent condensation on plant surfaces, ventilation to remove warm humid air and adequate spacing between plants is necessary. Air movement up between plants is facilitated by a wiremesh bench, while, in a solid-bench or tray system, horizontal air flow may be employed to provide the necessary air movement (Augsburger and Powell 1986). When considering nutrient effects on Botrytis blight, it is critical to avoid calcium deficiency. Calcium can reduce the effects of B. cinerea on roses (Volpin and Elad 1991). On poinsettias, calcium is an essential nutrient to prevent bract-edge burn, which provides a pathway for the initiation of Botrytis lesions on bracts. See chapter “▶ Nutritional Disorders of Florists’ Crops,” for additional information. Botrytis cinerea is notorious for developing resistance to systemic fungicides. Resistance to benomyl and thiophanate-methyl has been known in greenhouse populations of B. cinerea for some time (Maude 1980; Moorman and Lease 1992). Resistance to dicarboximides is known (Pappas 1982; Yourman and Jeffers 1999), and fenhexamid resistance has also been noted in a US greenhouse (Moorman et al. 2012). A low-residue treatment safe for use on bracts is desired by poinsettia growers, but many fungicides specifically prohibit use on poinsettias after bract development. Treatments with thermal dusts of the fungicide chlorothalonil have been used in the past to protect poinsettias at the end of a crop – but with the negative effect of bract fading noted in some cultivars (Powell 1977). In one study, injury to bracts with a thermal dust treatment of chlorothalonil was found on more cultivars grown with a 24  C/75  F night temperature than with at 21  C/70  F night temperature (Carlson and Emino 1969). Current labels in the United States indicate that a chlorothalonil thermal dust is to be used on poinsettias prior to bract formation only. The greatest reduction in disease severity was seen with fenhexamid and pyraclostrobin + boscalid treatment in a trial on poinsettias in bract (Leonberger and Beckerman 2009). Prior to bract coloration, a large number of active ingredients are available for use against Botrytis blight on poinsettias in the United States: chlorothalonil (FRAC M5), mancozeb (M3), coppers (FRAC M1), fludioxonil (FRAC 12), cyprodinil + fludioxonil (FRAC 9 + FRAC 12), fenhexamid (FRAC 17), iprodione (FRAC 2), polyoxin D (FRAC 19), pyraclostrobin + boscalid (FRAC 11 + 7), and strobilurins (FRAC 11). Biofungicides labeled in the United States for Botrytis management

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include Bacillus subtilis (FRAC NC), Streptomyces spp. (FRAC NC), and Reynoutria sachalinensis extract (FRAC P). Studies have shown the effectiveness of newer fungicides (Hausbeck and Harlan 2012), including fenhexamid (Benson and Parker 1999) and strobilurin materials (Benson and Parker 2000a, 2000b), on poinsettia against Botrytis blight. Genetic control is also relevant for Botrytis blight on poinsettias. Cultivars grown in the 1970s varied in susceptibility to the disease (Shanks 1972; Witte and Miller 1976; Manning et al. 1972). Since cultivars grown today are different from those in the 1970s, more recent cultivar comparisons would have more practical value. Elad and Evensen (1995) discussed the various processes associated with senescence that could inform the approach to breeding for resistance to B. cinerea in its many hosts. They suggested various techniques for thwarting senescence effects such as increased membrane permeability, including increasing the antioxidant systems and reducing the ethylene response. Physical barriers could also be optimized by developing a thicker, more hydrophobic cuticle, or providing inhibitors of cutinase. Giving poinsettias new defenses against cell wall-degrading enzymes or increasing the wound response rate would be additional strategies. Keeping latent infections quiescent longer and increasing the expression of various chemical defenses are additional goals that would result in improved poinsettia resistance to Botrytis blight. Learning how to stimulate natural plant resistance mechanisms may provide new avenues of management for B. cinerea on poinsettias in the future. Systemic acquired resistance effects were obtained in poinsettia with benzothiadiazole (BTH): treatments at 0.3 mM decreased the susceptibility of ‘Coco White’ and ‘Malibu Red’ poinsettia cultivars in a leaf-disc assay (Kulek and FloryszakWieczorek 2002).

2.5

Choanephora Wet Rot [Choanephora cucurbitarum (Berk. and Rav.) Thaxter]

2.5.1 Geographic Occurrence and Impact Choanephora cucurbitarum causes a soft, wet rot of poinsettias. The disease usually occurs at the hottest times (August through October) and may affect plants at all stages of production. 2.5.2 Symptoms/Signs Symptoms of Choanephora wet rot resemble those caused by Rhizopus stolonifer. A soft, mushy decay can develop on infected leaves and petioles. Infected stems wilt prior to collapsing. Young plants are destroyed under humid conditions, while older plants may develop only a few branches with symptoms. C. cucurbitarum produces characteristic whisker-like sporangiophores on diseased tissues (Engelhard 1987). 2.5.3 Biology and Epidemiology C. cucurbitarum also causes rot of fruits and vegetables as well as blighting of flowers and, in some cases, immature stems of Hibiscus rosa-sinensis and Petunia  hybrida.

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Disease develops very rapidly under warm to hot, humid to wet conditions. On bell pepper and swamp Hibiscus, the fungus ceases sporulation at temperatures above 14  C/57  F.

2.5.4 Management Minimizing plant stress and spacing to promote air circulation and minimizing leaf wetness have been suggested for management of this disease. Excellent control of wet rot on poinsettia has been reported with azoxystrobin (FRAC 11), copper pentahydrate (FRAC M1), and chlorothalonil (FRAC M5) (McGovern 1999).

2.6

Corynespora Leaf and Bract Spot [C. cassiicola (Berk. and M. A. Curtis) C. T. Wei (syn. Helminthosporium cassiicola Berk. and M. A. Curtis)]

2.6.1 Geographic Occurrence and Impact Corynespora leaf and bract spot on poinsettia was first reported from Florida (Chase and Simone 1986) and Louisiana (Holcomb and Fuller 1993). The disease has appeared only rarely in the past 20 years. 2.6.2 Symptoms On poinsettia, large, irregularly shaped, brown to black lesions form on bracts and leaves and strongly resemble those caused by Botrytis cinerea infections (Fig. 6). They occur primarily at the tips and margins of leaves and may be as large as 3 cm in diameter. In some cases disease is more prevalent on immature plants. 2.6.3 Biology and Epidemiology Wounding is not usually necessary for infection on poinsettia. C. cassiicola is spread by airborne spores. High moisture and humidity are conducive to infection. The fungus has been observed to survive on plant debris for 2 years. Lesions develop within 10 days of infection. Most isolates show no host specialization. A variety of plants are hosts for C. cassiicola, including Aeschynanthus pulcher (lipstick vine), Aphelandra squarrosa (zebra plant), Catharanthus roseus (annual vinca), Hydrangea macrophylla, Saintpaulia ionantha (African violet), and Salvia splendens (scarlet sage), as well as vegetable and field crops. 2.6.4 Management Reduction of humidity and leaf wetness duration are important for preventing infection. Fungicide trials on hydrangeas indicate excellent control with azoxystrobin (FRAC 11), triforine (FRAC 3), copper products (FRAC M1), and a premix of propiconazole and chlorothalonil (FRAC 3 and M5).

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Fig. 6 Corynespora leaf and bract spot on transitional bracts

2.7

Fusarium Stem Rot [Fusarium solani (Mart.) Sacco]

2.7.1 Geographic Occurrence and Impact Fusarium stem rot was reported from Wisconsin in 1987 (Heimann and Worf 1987). No additional research and few reports are available: the disease is apparently rare. 2.7.2 Symptoms Basal cankers, usually about 2 cm above the soil line and 3–6 cm long, are formed (Fig. 7). Lesions noted in Wisconsin were brown to black, somewhat shrunken, and had distinct necrotic margins. All stem tissues except the pith were affected and sometimes the bark tissue appeared shredded. Although cankers became obvious after potting, infected plants developed normally. Stress in the retail environment did result in wilting on retail shelves. 2.7.3 Management Symptom similarity with Rhizoctonia stem rot may cause misidentification by plant producers. Fungicides for prevention of Rhizoctonia stem rot fortunately could be expected to prevent Fusarium stem rot as well.

2.8

Powdery Mildews: Leveillula clavata Nour, L. taurica (Lev.) G. Arnaud, Pseudoidium poinsettiae (U. Braun, Minnis, and Yañez-Morales), Phyllactinia poinsettiae, and Ovulariopsis erysiphoides

2.8.1 Geographic Occurrence and Impact See Table 2 for reports of occurrence. The greatest impact of a powdery mildew on poinsettia has been seen from Pseudoidium poinsettiae (sometimes referred to as Oidium poinsettiae) affecting plants in the United States, Mexico, Puerto Rico, and Europe. The disease was first seen in the United States in 1990 in Pennsylvania and the Pacific Northwest

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Fig. 7 Fusarium cankers are brown and form at the soil line

Table 2 Powdery mildew reports on poinsettia from around the world (Farr and Rossman 2015; Siahaan et al. 2015; Daughtrey and Hall 1992; Anonymous 2002) Powdery mildew species Leveillula clavata Leveillula taurica syn. Oidiopsis taurica Leveillula sp. Oidium sp. Ovulariopsis erysiphoides Pseudoidium poinsettiae (syn. Oidium poinsettiae), Erysiphe poinsettiae Phyllactinia poinsettiae

Location Canary Islands, Italy, India, Indonesia, Kenya, Tanzania Canary Islands, Tanzania, India, Indonesia, Kenya Israel El Salvador, Ethiopia, Guatemala, Mexico, United States Venezuela Mexico, Puerto Rico, Denmark, Germany, Sweden, United States Indonesia

(Daughtrey and Hall 1992; Kim and Olson 1994); crops in Mexico and Puerto Rico were affected by the disease at about the same time (Daughtrey and Hall 1992). The disease appeared in numerous other states in subsequent years. The mildew seen in the United States has also been seen in the United Kingdom and Europe (Denmark, Germany, and Sweden) starting with the appearance of the Pseudoidium in Denmark in 2005 (Anonymous 2002). The impact of this powdery mildew comes not from its effect on plant health, but from the visibility of the fungus: the aesthetic quality of the crop plummets immediately, and infected plants are not saleable. In addition, if poinsettias are sold that have unnoticed infections, the disease will continue to develop in homes and interiorscapes, resulting in obvious white spotting on colored bracts that leads to immense customer dissatisfaction. After the initial few years in which the disease was seen in the poinsettia production industry, when many growers experienced total crop failures, incidence has been sporadic and economic losses have been much less severe. This is due to vigilant scouting and judicious

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Fig. 8 Powdery mildew causes chlorotic spots on the upper leaf surface, which mark the location of colonies on the undersurface

Fig. 9 Powdery mildew colonies appear first on the leaf undersurface

applications of protective fungicide at all levels of production. This powdery mildew continues to be found in a few greenhouses in North America almost every year.

2.8.2 Symptoms/Signs Powdery mildew caused by Pseudoidium poinsettiae can be difficult to scout for during the spring and summer, as there are no white colonies then on the upper leaf surface: the only symptoms may be a few round, diffuse, chlorotic spots on the leaves (Fig. 8). Beneath these pale areas, on the abaxial side of the leaf, there are sparse, often indistinct, powdery mildew colonies (Fig. 9). In the fall, when greenhouse temperatures cool, conspicuous white colonies develop on the upper surface of leaves and bracts (Fig. 10). When oval to cylindrical conidia are produced (singly, but forming false chains) from the surface mycelium, the colonies take on a more granulated (“sugar-coated”) appearance. If the growth of the powdery mildew is unchecked by fungicide application, colonies will grow until they merge and cover the surface of leaves or bracts.

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Fig. 10 Bracts as well as leaves are susceptible to powdery mildew

Powdery mildew symptoms caused by Leveillula clavata (syn. Oidiopsis taurica) as described from a 2005 appearance in Italy (Garibaldi et al. 2006) are similar to those in early (summer) infections of poinsettia by Pseudoidium poinsettiae in the United States – both fungi cause yellow spots on adaxial leaf surfaces. In other respects the disease caused by L. clavata is quite different: bract infections have not been reported with L. clavata. Club-shaped conidia are borne singly on conidiophores that emerge through stomates on the abaxial leaf surface, where a rose to reddish mycelium also appears. Chasmothecia with a hemispherical, lenticular or pezizoid shape and simple appendages, containing many asci each with two ascospores, were described on poinsettias in Kenya (Nour 1957). A similar powdery mildew fungus has also been reported on poinsettia from Japan (Horie et al. 2006). A third genus of powdery mildew, Phyllactinia poinsettiae (syn. Ovulariopsis poinsettiae), was recently reported from Indonesia (Siahaan et al. 2015). This mildew forms subevanescent colonies on the abaxial side of leaves. Club-shaped conidia with papillate tips are produced singly on conidiophores arising from hyphae.

2.8.3 Biology and Epidemiology Studies of powdery mildew management on poinsettia have largely been confined to Pseudoidium poinsettiae (syn. Erysiphe poinsettiae). Studies at Michigan State University on this fungus showed conidial germ tubes appearing on poinsettia ‘Freedom Red’ at 2 h post-inoculation (Celio and Hausbeck 1998); germination reached a peak at 36 h. Appressoria were slightly lobed to lobed. Over half of the germinated conidia had established a globose haustorium within 48 h. Basal cells of conidiophores were arced distinctively. Early steps in colony establishment (percentage of conidial germination, secondary germ tube formation, and haustorium formation) were limited at 30  C/86  F as compared to 20  C/68  F. The powdery mildew can be maintained on symptomless plants over the summer (Kim et al. 1995a), thus growers should not count on disease eradication through exposure to typical summer greenhouse temperatures. The suppressive effect of high temperatures on powdery mildew of poinsettia is, however, reflected in field observations

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that the disease does not progress during the summer months in the northeastern United States (Kim et al. 1995a). The optimum for germination of conidia was found to be 25  C/77  F and 85 % relative humidity (RH) in greenhouse studies (Hausbeck and Kalishek 1994), conditions which are typically available in greenhouses in temperate climates in the fall. Peak concentrations of conidia in the greenhouse were associated with dips in RH or changes in temperature, which often occurred simultaneously (Shaw and Hausbeck 1995; Byrne et al. 2000). Symptoms were more severe and the conidial concentration was higher (1,221 conidia/m3 air/h) in one greenhouse with an average RH of 64 %, while the conidial concentration in a similar greenhouse with an average RH of 53 % was only 317 conidia/m3 air/h. Studies in Germany indicated that the powdery mildew from poinsettia (referred to as Oidium sp. but presumably the Pseudoidium seen in the United States at about the same time) could be transferred to other Euphorbia spp. as well E. exigua, E. heterophylla, E. helioscopia, and E. marginata (Anonymous 2004). These plants or wild E. pulcherrima could provide a reservoir of inoculum near poinsettia cutting production facilities.

2.8.4 Management • Cultural practices – Keeping humidity down (30  C/86  F) has been suggested as a possible technique for specialist propagators to use prior to shipping cuttings (Celio and Hausbeck 1998), but the plant safety and effectiveness of this procedure have not yet been tested. • Sanitation – Fortunately P. poinsettiae has not been found causing disease on any other hosts in northern temperate climates, so removing all poinsettias from the greenhouse at the end of the growing season is likely to provide complete eradication of inoculum. Inadvertent reintroduction on cuttings the next season is the grower’s only concern unless plants are being grown in subtropical areas where poinsettias grow year-round. Removing infected leaves as soon as they are detected is not often a practical solution, but reduces inoculum in the greenhouse and at high greenhouse temperatures could be eradicative early in the growing season (Hausbeck et al. 1994). Early detection of powdery mildew is important, as it alerts the grower to the need for an appropriate fungicide program for that year’s crop. Leaf removal must be combined with fungicide treatment to avoid extensive crop loss in the fall, when greenhouse conditions become more conducive to disease development and spread. Chasmothecia have not been found for P. poinsettiae, thus there is no persistent inoculum on leaf debris in the greenhouse after plants are sold. • Fungicides and biocontrols – Fungicides have been very effective against the Pseudoidium on poinsettia, perhaps in part because the fungus was introduced to horticultural productions systems only recently. Both protectant and systemic

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materials have been very effective when used preventively. Sprays for powdery mildew control need not be applied unless the pathogen is present, so a careful scouting program should be in place in order to detect the disease before it is widespread. Many fungicide labels restrict use on poinsettias to the time before bracts begin to color, so chemical control must be established before the finishing stages. Weekly or biweekly treatments of a wide range of materials including potassium bicarbonate (FRAC NC), copper pentahydrate (FRAC M1), thiophanate-methyl (FRAC 1), trifloxystrobin (FRAC 11), kresoxim-methyl (FRAC 11), polyoxin D (FRAC 19), pyraclostrobin (FRAC 11), iprodione (FRAC 2), and demethylation inhibitor materials (triflumizole, triadimefon, and myclobutanil) (FRAC 3) have provided significant protection (Daughtrey and Macksel 1995; Daughtrey and Tobiasz 1999; Gould and Bergen 1995a, b; Hausbeck and Kusnier 1995a, b, c, 1996; Hausbeck et al. 2002a, b; Daughtrey and Tobiasz 2009). Rotation of active ingredients with different modes of action is essential for powdery mildew management, so it is fortunate that there are many fungicide options for this purpose during all but the finishing stages of production. Biological controls are desirable for powdery mildew control in poinsettias, particularly because growers are not comfortable applying fungicides to colored bracts and because many fungicide labels prohibit such use. A program alternating a biological control (Bacillus subtilis) with a biorational material (potassium bicarbonate) reduced powdery mildew significantly in an inoculated trial (Daughtrey and Tobiasz 2010). • Resistance – There is no known resistance to powdery mildew (Pseudoidium poinsettiae) in poinsettia, but the level of susceptibility varies among cultivars. Celio and Hausbeck (1997) tested 11 cultivars for susceptibility, and all developed powdery mildew colonies within 31 days. Three red cultivars tested had significantly more powdery mildew than other cultivars in two trials. Kim et al. (1995b) reported higher susceptibility in leaves and bracts of the cultivar ‘Freedom Red’ as compared to the older cultivars ‘Dark Red Hegg’ and ‘V-17 Angelika White’ in a greenhouse trial. If the most popular red cultivars are prone to powdery mildew, growers may not have the option of growing less susceptible types. Prioritized scouting of the more susceptible red cultivars has been suggested (Celio and Hausbeck 1997) to improve scouting efficiency. In a trial in Germany, Euphorbia fulgens and E. milii were not infected with the pathogen although four other Euphorbia species were shown to be hosts (Anonymous 2004). Plant breeders will ideally expose prospective new lines to P. poinsettiae to select for poinsettias with low susceptibility to this disease.

2.9

Pythium Root Rot (Pythium spp.)

2.9.1 Geographic Occurrence and Impact Pythium species are water molds belonging to the Oomycetes and not true fungi. The species reported to occur on poinsettia are P. aphanidermatum, P. cryptoirregulare (syn. Globisporangium cryptoirregulare), P. debaryanum (syn. Globisporangium

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Fig. 11 Pythium root rot (at right) in contrast to a healthy root system

debaryanum), P. irregulare (syn. Globisporangium irregulare), P. perniciosum, and P. ultimum (Farr and Rossman 2015) as well as P. myriotylum and P. helicoides (Miyake et al. 2014). Strider and Jones (1985) mention P. ultimum as the primary problem in poinsettia production up until that time, but P. aphanidermatum has caused the most dramatic losses in recent years (Moorman et al. 2002). Pythium root rot is widely distributed, and no doubt occurs wherever this crop is grown. Formal reports of Pythium root rot on poinsettia caused by one or more species have come from South America, the United States, Japan, New Zealand, and Brazil (Farr and Rossman 2015). The disease can have imperceptible effects on plant quality or can cause extensive mortality of poinsettias, especially in irrigation systems that allow recirculation of water. The particular Pythium species that are important today are different from those that were important in the first half of the twentieth century, largely due to a shift to production systems using soilless media and recirculating subirrigation. Changes in the cultivars grown may also have affected which root diseases are most important, and production of cuttings in new geographic areas around the world may have led to the introduction of new pathogen species or strains.

2.9.2 Symptoms/Signs There is a soft brown or gray-brown rot of the cortex of the fibrous roots, which contrasts sharply with the white, vigorous growth of a healthy root system (Fig. 11). Severe aboveground symptoms of Pythium root rot include stunting of the whole plant and even wilting and death. Leaves yellow, starting with the lower leaves, and may curl upward as they wilt. In advanced infections, the cortex will slough off the hard xylem core as the plant is pulled from the pot, leaving behind thin white strings of xylem (Fig. 12). Symptoms appear on the roots on the outside of a rooting cube or on roots adjacent to pot sidewalls; they are generally most pronounced in the bottom of the pot. Because Thielaviopsis basicola and Rhizoctonia solani both cause a drier looking rot, Pythium root rot is most easily confused with Phytophthora or Fusarium

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Fig. 12 Pythium root rot (showing xylem remaining after cortex has sloughed off)

root rots. Fungus gnat feeding is often coincident with Pythium root rot, but either the insect or the water mold acting alone can cause serious root injury. See chapter “▶ Insect Management for Disease Control in Florists’ Crops,” for additional information.

2.9.3 Biology and Epidemiology The biology of the different Pythium species varies somewhat, but there are characteristics that they share. They are fungus-like in their appearance, but molecular studies have verified the longtime suspicion that they are members of a unique group, more closely related to algae than to fungi. Pythium spp. possess coenocytic hyphae with cell walls composed of cellulose rather than the chitin of a true fungus. In the wet environments that are favorable for infection, most Pythium species accomplish asexual reproduction by forming sporangia (in some cases these are merely swollen hyphae). Motile zoospores are released from thin-walled vesicles produced by the sporangium of a Pythium sp., rather than being formed within the sporangium in the manner of Phytophthora. Often chlamydospores and/or hyphal swellings may be formed by the hyphae as well. The sexual structures are termed oogonia and antheridia, with these structures sometimes coming from the same individual and sometimes requiring an opposite mating type. The result of fertilization of an oogonium by one or more antheridia results in the formation of an oospore. The thick walls of the oospore make it very resistant to environmental stresses and to other microorganisms, so it is a long-term survival structure. Oospores in organic debris are the main source of contamination in a greenhouse operation. Oospores are sometimes present in the peat moss itself or sometimes linger in recycled pots or trays that have not been thoroughly cleaned and disinfested. Morphological differences were originally used to identify Pythium species, but DNA analysis has led to many changes in Pythium taxonomy and nomenclature in recent years, including the creation of new genera that were once considered to be within the genus Pythium (Uzuhashi et al. 2010). Because different Pythium species have different temperature optima, problems with particular species may appear at different times of year. The most commonly

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encountered Pythium in earlier years, when poinsettias were grown in soil-based media, was P. ultimum; root rot from this pathogen was shown to be most severe at 17  C/62  F and to cause little effect at temperatures at or above 26  C/79  F (Bateman and Dimock 1959). Pythium root rot was at that time seen commonly at the end of the production season, when temperatures were lowered to slow plant development prior to sale. Today, P. ultimum is occasionally encountered, but the most serious Pythium pathogen on poinsettia in North America today is Pythium aphanidermatum, first seen in 1977 as a pathogen on poinsettia in Canada on plants grown from cuttings propagated in the United States (Moorman et al. 2002; Komorowska-Jedrys et al. 2008). This organism causes disease at higher temperatures than P. ultimum (Hendrix and Campbell 1973) and produces prolific numbers of zoospores that make it ideally suited to thrive in recirculating irrigation systems (Sanogo and Moorman 1993). Temperatures are favorable to P. aphanidermatum in summer, when most of the vegetative growth of poinsettias takes place. This species was reported causing cutting rot in soilless production systems in Brazil (Palmucci and Grijalba 2007). In addition, Miyake et al. (2014) reported sudden outbreaks of root rot in ebb-and-flood production of poinsettias in Japan, with P. myriotylum and P. helicoides discovered to be the pathogens. Their subsequent research found that isolates of all three high-temperature-loving Pythium species – P. aphanidermatum, P. myriotylum, and P. helicoides – gave the greatest incidence and severity of root rot at 35  C/95  C when inoculated to poinsettia. Although favored by hot environments, all three were able to cause disease at 20  C/68  F at higher inoculum (zoospore) levels. Bateman (1961) found that overwatering (keeping soil at 70 % moisture holding capacity or above) promoted Pythium root rot in ‘Barbara Ecke’ poinsettias. He also showed that neutral to alkaline soil was favorable to root rot caused by P. ultimum (Bateman 1962). A study by Moorman (1986) showed that growing poinsettias with higher fertility increased Pythium root rot caused by P. ultimum. Fertilization rates tested ranged from 100 to 600 μg N/g. Higher soluble salt levels were correlated with increased mortality in poinsettias inoculated with P. ultimum early in production in three different soilless mixes as well as in a soil-containing mix.

2.9.4 Management Identification of the species of Pythium present is important to designing a management program. Both P. irregulare and P. cryptoirregulare, indistinguishable morphologically, occur on poinsettias in greenhouses (Garzon et al. 2007) and are rarely problematic unless cultural controls are mismanaged. P. aphanidermatum, P. helicoides, and P. myriotylum are all relatively aggressive pathogens and are all favored by high temperature. Globalization of the ornamental plant trade is no doubt having an effect on the species and genotypes of Pythium and other pathogens that are encountered in greenhouses. An analysis of P. aphanidermatum found on poinsettia in Pennsylvania greenhouses noted genotypes from around the world (Lee et al. 2010). Variation in susceptibility to P. aphanidermatum among poinsettia

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cultivars has been noted (Daughtrey and Hyatt 2012); growers experiencing extensive losses may find it worthwhile to trial different cultivars. Poinsettia growers are generally fearful of Pythium root rot losses and use very careful sanitation procedures and treatments with biological and chemical fungicides to ensure crop safety. Sanitation efforts are directed against the introduction of oospores in field soil, a particular threat in flood-floor operations. Whereas earlier production methods included steam pasteurization to ensure the safety of soilcontaining growing media, today’s poinsettia growers use soilless growing mixes based largely on sphagnum peat moss. Cuttings are often rooted in artificial foam materials – a practice which eliminates peat moss as a possible source (Graham 1994). Clean growing surfaces and clean irrigation and handling practices using soilless media provide a measure of security that was not available to a poinsettia grower 50 years ago. Sanitation between crops will be especially important in operations with recirculating irrigation systems, which favor dissemination and infection of Pythium spp. Miyake et al. (2014) noted that poinsettias were first seen with root rot caused by P. helicoides in Japan in ebb-and-flow production following rose and begonia crops that were known hosts of that species. Although monitoring for Pythium root rot is usually a matter of watching for symptoms, bentgrass traps have been used to monitor for populations of high-temperature Pythium species in ebb-and-flood systems in Japan. These traps successfully detected the pathogens 30 days before symptom development was evident from infections of P. helicoides in potted miniature roses (Watanabe et al. 2008). Boehm and Hoitink (1992) found that higher microbial activity in a light-colored, less-decomposed sphagnum peat was associated with short-term suppression of Pythium root rot. Treatments with bioantagonists against P. ultimum var. ultimum in subirrigated poinsettias were trialed in Canada (Little et al. 2003). Quality was similar when inoculated plants were protected with metalaxyl or with treatments of Streptomyces griseoviridis strain K61 or S. lydicus strain WYEC 108. Other bioantagonist products including Trichoderma spp. and Bacillus subtilis active ingredients are also labeled for use against Pythium spp. in greenhouses. There are few fungicides with strong effectiveness against Pythium. Trials have shown the relative effectiveness of some of the oomycete fungicides (Parker and Benson 2011, 2012, 2013; Benson and Parker 2001b; Hausbeck and Harlan 2007). Fungicides that have been used for Pythium management include metalaxyl/ mefenoxam (FRAC 4), the effectiveness of which has been reduced in the greenhouse arena due to the development of resistance in some Pythium populations (Moorman et al. 2002). The effectiveness of etridiazole (FRAC 14), sold singly and also in combination with thiophanate-methyl (FRAC 1) for broad-spectrum root rot management, continues: there has not been any resistance to this active ingredient in greenhouse Pythium populations. Fosetyl-Al and other phosphorous acid materials (FRAC 33) can reduce Pythium losses to some degree. More recently, cyazofamid (FRAC 21) and fluopicolide (FRAC 43) have been introduced for control of Pythium root rot in greenhouses. Strobilurins (FRAC 11) give partial but not strong control of Pythium.

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M.L. Daughtrey and A.R. Chase

Phytophthora Crown and Root Rot (Phytophthora drechsleri, P. nicotianae, P. cryptogea)

2.10.1 Geographic Occurrence and Impact Three species of Phytophthora have been reported from poinsettia: P. drechsleri (Korea and the United States) and P. nicotianae (United States, Korea, Brazil, Puerto Rico, Japan) have been known to be poinsettia pathogens for some time (Farr and Rossman 2015; Kanto et al. 2007; Estevez de Jensen et al. 2006). In Taiwan, P. cryptogea as well as P. nicotianae occurred on rotted poinsettia roots (Ann 1992), and Orlikowski also described P. cryptogea on poinsettia in Poland (Orlikowski and Ptaszek 2013). Some additional species have been seen on Euphorbia relatives of poinsettia, including P. cactorum, P. citricola, and P. palmivora. The disease is lethal to infected plants. Its impact depends upon the production system: in greenhouses with recirculating irrigation systems, the potential for extensive crop loss is high. Poinsettia cultivars vary in their susceptibility to Phytophthora root rot (Woodworth and Hausbeck 2003). 2.10.2 Symptoms/Signs The part of the poinsettia plant affected by Phytophthora varies from outbreak to outbreak: whereas Engelhard and Ploetz (1979) described P. nicotianae causing a crown and stem rot, in Hawaii a short time later, P. nicotianae and P. drechsleri were found causing symptoms on leaves, cyathia, and bracts (Yoshimura et al. 1985). In the United States, P. drechsleri has been a significant problem in cutting production in ebb-and-flood benches, causing root rot and stem rot (Lamour et al. 2003) (Fig. 13). P. cryptogea caused extensive stem base rot as well as root rot and wilt on stunted poinsettias in an outbreak in Poland (Orlikowski and Ptaszek 2013). 2.10.3 Biology and Epidemiology The pathogen is thought to be moved from greenhouse to greenhouse on cuttings (Lamour et al. 2003). Infected stock plants may lead to extensive loss of cuttings during propagation. Because of the production of sporangia and zoospores, dispersal Fig. 13 Root rot caused by Phytophthora drechsleri

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of Phytophthora inoculum during production is especially likely in plants grown on a subirrigated recirculating irrigation system (Strong et al. 1997). In a US study, the populations of P. drechsleri affecting two poinsettia greenhouses were unique to the two locations and showed no genetic diversity within a greenhouse (Lamour et al. 2003), indicating asexual reproduction within the two facilities.

2.10.4 Management Removing the obviously diseased plants from a recirculating irrigation system is not sufficient to remove all of the inoculum when attempting to halt an epidemic. A system for filtering beneath each pot would be ideal (Van der Gaag et al. 2001), or alternatively treating water to be toxic to zoospores would be useful (von Broembsen and Deacon 1997). Trials have shown the benefit of certain fungicide treatments to prevent Phytophthora disease on poinsettias (Daughtrey and Tobiasz 2003; Hausbeck and Harlan 2009). Current methods in the production industry rely heavily upon chemical control in response to an outbreak, with fungicides registered for this use in the United States including cyazofamid (FRAC 21), dimethomorph and mandipropamid (FRAC 40), etridiazole (FRAC 14), fluopicolide (FRAC 43), fosetyl-Al and other phosphorous acid compounds (FRAC 33), mefenoxam (FRAC 4), and strobilurins (FRAC 11) (Daughtrey et al. 2015). There is little published data on the use of biopesticides for Phytophthora disease suppression in poinsettias (Hausbeck et al. 2004a).

2.11

Rhizoctonia Cutting Rot, Crown Rot, and Root Rot [Rhizoctonia solani Kuhn (syn. Thanatephorus cucumeris (A. B. Frank) Donk)]

2.11.1 Geographic Occurrence and Impact Rhizoctonia solani is widely distributed, has a wide host range, and is very common in propagation of poinsettias. 2.11.2 Symptoms/Signs When R. solani causes cutting rot, stems become darkened and mushy at the soil line (Fig. 14). Under the warm, humid conditions typical of propagation, the fungus will grow over the rooting cube as well as the entire cutting (Fig. 15). When this occurs, a light brown mycelium, closely appressed to the plant tissues and soil, is sometimes visible. The fungus grows out more or less radially from the point of contamination, giving the damage a circular or arc-shaped pattern. The mycelium can also cause a web blight and appears as a brown, cobweb-like growth on the leaves. Leaves in contact with the growing medium can develop roughly circular lesions (Fig. 16). Root rot caused by R. solani usually shows discrete, brown lesions and rotting of the cortical tissues, leading to decreased plant development. R. solani causes cutting rot more often than root rot, although poinsettia is quite susceptible to Rhizoctonia root rot.

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Fig. 14 Rhizoctonia stem rot

Fig. 15 Root rot caused by Rhizoctonia solani, with visible mycelium on the surface of rooting cubes

Crown rot commonly occurs in the absence of root rot and first appears as a brown canker at the root crown. Longitudinal cracking and a dry appearance of the rotted crown tissues often develop on older plants. Other aboveground symptoms include chlorosis, wilting, loss of lower leaves, and in severe cases stunting and plant death.

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Fig. 16 Mycelium of Rhizoctonia solani on a blighted leaf

2.11.3 Biology and Epidemiology Poinsettias are most susceptible just before or soon after rooting and just prior to plant maturity. R. solani generally grows best in potting media that are evenly moist and warm: Rhizoctonia root rot increases at soil temperatures between 17 and 26  C (62 and 79  F) (Bateman and Dimock 1959) with a moisture holding capacity below 40 %. R. solani also grows best in soils with high oxygen and low carbon dioxide levels (Bateman 1961). 2.11.4 Management Sanitation practices to guard against soilborne fungi are applicable to R. solani. Methods for keeping field soil out of contact with a soilless growing medium are effective at preventing most Rhizoctonia disease problems for poinsettias. • Cultural methods – Composts from municipal sewage sludge have been explored for use in ornamental production. Compost cured for at least 4 months with an internal temperature 93% relative humidity) prevail, making it a common occurrence in the greenhouse. The fungus, Botrytis cinerea, is ubiquitous and infects most aboveground plant tissue. It infects almost all florist crops (Daughtrey et al. 1995). On cyclamen, it is considered one of the most important diseases of the crop (Kessel et al. 1999) and can reduce profitability directly by destroying the flowers and making them unmarketable. Symptoms/signs. Symptoms on petals begin as small, bleached spots (Fig. 4) that enlarge, coalesce, and eventually cause the petals to blight. Similarly, the fungus can infect petioles causing them to collapse. B. cinerea can often be observed in the lower canopy sporulating on dead and senescing tissue (Fig. 5). These sites become

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Fig. 5 Botrytis cinerea sporulating on cyclamen stems (Courtesy of Diffusion Morel Co.)

major focal points for the production of inoculum. The symptoms of Botrytis blight can be diverse, manifesting as blight or as distinct lesions. Leaves tend to be less susceptible than the flowers. Biology and epidemiology. Survival of the pathogen occurs as mycelium on plant debris (Yunis and Elad 1989) and as sclerotia (Araújo et al. 2005). The pathogen produces spores, borne on stalks, which are released with air currents and water splashing. A factor that is crucial to Botrytis infection is high humidity (>93%), and this is commonly achieved in microclimates within the canopy in a greenhouse. Initial infections begin following a buildup of inoculum on dead and senescing leaves within the canopy. Once sufficient inoculum has been released, healthy leaves and flowers are infected and the epidemic begins, provided susceptible tissue is available and high humidity prevails. Management. Successful management of Botrytis blight on cyclamen must include cultural practices and sanitation along with chemical and/or biological practices to suppress inoculum buildup. • Cultural practices – Given that Botrytis blight occurs when the relative humidity reaches above 93%, one major component of disease management is to reduce the humidity. Proper spacing and ventilation can help eliminate the humid microclimates within the canopy where infection can occur. Heating and venting greenhouses at dusk can expunge large amounts of humid air and suppress infection (Daughtrey et al. 1995). Any efforts to keep the foliage dry by using subirrigation systems can also reduce humidity in the canopy. When subirrigation is not available, overhead watering should be done in the morning to allow time for the foliage and flowers to dry. Proper spacing of plants favors rapid drying of leaves and flowers. When possible, computer-controlled environmental systems can be used to increase ventilation and raise temperature to avoid high humidity. Postharvest conditions can also affect Botrytis blight development in cyclamen. On C. persicum, the presence of ethylene increased disease, while applying the ethylene antagonist 1-methylcyclopropene extended the aesthetic life of inoculated flowers by 25 d (Seglie et al. 2009).

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• Sanitation – Although the fungus is ubiquitous, any effort to prevent inoculum from increasing in and around the production facility will result in fewer infections. Efforts should be made to remove dead tissue on the bench or greenhouse floor, which the fungus will use as a substrate to sporulate. Efforts should be made to remove all pruned flowers as they, along with defoliating leaves, can land on healthy tissue and quickly lead to new infections. Fungicides and biocontrols – A considerable amount of research has been done on chemical control of Botrytis blight, in part due to the propensity of the pathogen to develop resistance (Leroux 2007; Moorman and Lease 1992). There are many products in many different families of fungicides that can provide excellent suppression providing tolerant strains have not been allowed to proliferate. The fungicides commonly used today include chlorothalonil, singly and in combinations, along with iprodione, fludioxonil, fenhexamid, strobilurins, and triflumizole. Additional combination products including cyprodinil + fludioxonil and pyraclostrobin + boscalid are also employed in Botrytis blight management. Fungicide resistance is common for thiophanate-methyl and iprodione due to the extensive movement of plant materials in the horticultural trade. Once resistant strains are present in an operation, alternate classes of fungicides need to be used. More environmentally safe active ingredients are also labeled for control of Botrytis blight, such as bicarbonates, Bacillus subtilis, and reduced risk materials like polyoxin D. There has been other work using the biological control Ulocladium atrum, which performed as well as chemical fungicides in suppressing Botrytis blight on cyclamen (Köhl et al. 2000). The chitinolytic bacterium Serratia marcescens also suppressed Botrytis blight on cyclamen petals by 60% and was equal in effect to the dicarboximide fungicide iprodione (Iyozumi et al. 1996).

2.3

Fusarium Wilt of Cyclamen (Fusarium oxysporum Schlechtend.:Fr. f. cyclaminissp. Gerlach)

Geographic occurrence and impact. Fusarium wilt was first reported in 1935 in Germany (Wollenweber and Reinking 1935). Outbreaks were subsequently reported from France (Barthelet and Gaudineau 1936), Italy (Bongini 1940), England (Moore 1947), the United States in 1949 (Tompkins and Snyder 1972), Bulgaria (Khristova 1958), Belgium (Rouxel and Grouet 1974), and Portugal (Pitta and Teranishi 1979). The fungus was described and labeled F. oxysporum f. sp. cyclaminis in 1954 by Gerlach (1954). In 1977, Fusarium wilt was reported for the first time in the Netherlands, where it was particularly destructive due to the recycling of contaminated irrigation water (Rattink 1986). No races of the pathogen exist, but some clonal lineages have been observed (Woudt et al. 1995). The disease can be extremely destructive and has resulted in losses of over 50% of the crop in greenhouses (Elmer and Daughtrey 2012). Symptoms/signs. All reports of Fusarium wilt of cyclamen have described the same symptoms regardless of locale. In young plants, the disease first appears as

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Fig. 6 Chlorosis associated with Fusarium wilt (Photo courtesy of Margery Daughtrey)

Fig. 7 Collapsed plant due to Fusarium wilt (Photo courtesy of Margery Daughtrey)

Fig. 8 Vascular discoloration in the cyclamen corm due to Fusarium wilt (Photo courtesy of Margery Daughtrey)

bright-yellow chlorosis (Fig. 6), which begins at the base of the leaf and then expands outward toward the perimeter. Alternatively, and more often in older plants, chlorotic patches may develop on the leaf blade or at the leaf edge. In both situations, the affected leaf portions become flaccid and loose turgidity (Fig. 7). Symptoms may appear at all stages of growth and can be observed on roots, corms, and aboveground parts. Infected roots exhibit vascular discoloration and may be totally discolored and darkened. Reddish-brown vascular discoloration is frequently observed in the corm (Fig. 8). Fusarium wilt can be distinguished from the bacterial soft rot diseases (caused by

Diseases of Cyclamen

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Pectobacterium spp. and Dickeya spp.; see below) largely because the corm remains firm and shows vascular discoloration in the case of Fusarium wilt. The vascular discoloration can turn to brownish black, dark red, or purplish over time. Many times, no symptoms are apparent when the plant is vegetative, but then it quickly develops wilting and chlorosis when flowering occurs. The timing is most unfortunate since growers have expended considerable resources only to lose the crop when it becomes marketable. The latent expression of symptoms (Gerlach 1954; Rouxel and Grouet 1974) has cost the industry considerable profits. Molecular assays that quickly detect the pathogen in pot leachates might allow growers to identify and discard contaminated seedlings. Recently, a set of polymerase chain reaction (PCR) primers proved effective experimental tools for identification and detection of F. oxysporum f. sp. cyclaminis, distinguishing the pathogen from nonpathogenic strains of F. oxysporum often associated with cyclamen. However, these molecular tools have not been commercialized for use. Biology and epidemiology. Infection results from invading hyphae that can originate from a microconidium, macroconidium, chlamydospore, or mycelium. Most studies on inoculum density revealed disease severity is a function of inoculum load (Elmer 2002b; Garibaldi 1988; Rattink 1986, 1990). Although seed transmission is strongly suspected, it has been very difficult to routinely detect the pathogen from seeds (Tompkins and Snyder 1972). The pathogen can be spread from plant to plant by overhead watering, contaminated tools, soil, pots, and trays. Although insect vectoring is strongly suspected, definitive experiments demonstrating transmission by fungus gnats and/or shore flies have yet to be done (Gillespie and Menzies 1993). Spread by windblown conidia may also be a means for shortdistance spread of F. oxysporum f. sp. cyclaminis, and this should be considered a possible means of dissemination in greenhouses. Short-distance transmission can also occur via irrigation water, despite reports stating F. oxysporum f. sp. cyclaminis does not spread in water as effectively as other Fusarium fungi (Krebs 1985). Ebb and flow recycling systems have been associated with major outbreaks in the Netherlands (Rattink 1986). However, disease thresholds are rarely reached if scouting and removal of symptomatic plants are done promptly. Long-distance transmission of the Fusarium pathogen on seedling plugs is a recognized problem within the cyclamen industry. Rattink (1986) found that the time of the onset of symptoms became shorter as temperatures increased from 15  C (59  F). The ideal temperature for infection was 27.5  C (81.5  F). Wide fluctuations in temperature have also been associated with increased disease, possibly by placing heat and drought stress on the host. Management • Cultural practices – Fusarium wilt of cyclamen is one of the more difficult diseases to manage (Elmer and Daughtrey 2012). All attempts to identify horticulturally acceptable resistance in the host have been unsuccessful. OrliczLuthardt (1998) and Ewald et al. (2000) have identified tolerance to Fusarium wilt in related hybrids of C. persicum Mill.  C. purpurascens Mill., but no further breeding has been advanced for disease resistance. Steam sterilization of

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the soil can greatly reduce disease. However, growers should exercise caution when using soil amendments with steam-sterilized soil. Garibaldi (1988) reported that steam-sterilized local medium made from soil, peat, beech leaves, and manure (30:40:20:10) was highly conducive to disease. Growers should always experiment with small batches before incorporating new amendments. Fertilizing with nitrates and adding lime to achieve a 6.5–7.0 pH in the rooting medium will suppress Fusarium wilt of cyclamen. Fertilizers that acidify the soil and lower pH, such as ammoniacal sources of N [NH4NO3, (NH4)2SO4], should be avoided. Amending the soil with well-rotted composts is also effective at suppressing Fusarium wilt, because the dominant form of N in these composts is nitrate. In addition, chloride salts can reduce the severity of disease, but growers should exercise caution and test on a small number of plants before applying salts (Elmer 2002b). • Fungicides – Fungicides have been useful in protecting against Fusarium wilt if applied preventively and during early onset but generally fail in providing acceptable disease control in infected plants. Soil drenches with benomyl, thiophanate-methyl, chlorothalonil, and tolylfluanid provide little or no suppression. Gullino et al. (2002) found that soil drenches with azoxystrobin were effective, whereas the other strobilurins tested, such as kresoxim-methyl and trifloxystrobin, were phytotoxic. Elmer and McGovern (2004) found that some combinations of biological fungicides with the registered fungicide fludioxonil were more effective than the fungicide alone in reducing disease progress. Disease onset was delayed 3 weeks when acibenzolar-S-methyl, a product that induces resistance to Fusarium wilt, was applied, but marked phytotoxicity was also observed (Elmer 2006). • Biological control – Migheli and Garibaldi (1995) and Minuto et al. (1995) reported that disease incidence could be reduced by supplementing the growing media with nonpathogenic antagonistic strains of Fusarium species applied alone or in mixtures. Later, Minuto et al. (2004) showed that nonpathogenic Fusarium oxysporum strains were superior to benomyl in protecting cyclamen from Fusarium wilt. Elmer (2001, 2002a) found combinations of nonpathogenic Fusarium oxysporum, Pseudomonas fluorescens, and other agents that could suppress disease, but these products have not been made commercially available. Other attempts at biological control have resulted in some success in reducing disease incidence and severity. A strain of the bacterium Serratia marcescens that has chitinolytic activity reduced Fusarium wilt for up to 3 weeks after inoculation with the pathogen; at harvest, only 20% of the plants were diseased compared to 90% incidence in the untreated control (Someya et al. 2000). Unfortunately, experimentation with biological fungicides has not yielded a commercially acceptable management strategy. Even when combinations of biological products were applied preventively, disease control in inoculated plants never reached an acceptable level, even when the plants were subsequently returned to a conventional chemical fungicide program (Elmer and McGovern 2004). Disease suppression is sometimes improved by combining biological products with fungicides (Minuto et al. 1995; Elmer and McGovern 2004; Daughtrey and

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Tobiasz 2005). Growers should read the labels and consult resources to determine if products are compatible with one another. • Integrative strategies – Until host resistance becomes available, growers must rely on a combination of strategies. The more effective practices for management of Fusarium wilt include strict sanitation, crop monitoring, and immediate fungicide application.

2.4

Phyllosticta Leaf Spot (Phyllosticta cyclaminis Brun)

Geographic occurrence and impact. The fungus was first described in 1890 by Brun, but no reports of economic damage were recorded until 1929 when a report of its occurrence was made in Italy, where the disease caused defoliation (Bongini 1930). The disease is also listed as a potential disease to manage in California, but outbreaks appear to be sporadic. Symptoms/signs. The symptoms are first observed as yellowish-brown spots that appear near the margins. Lesions later develop concentric light and dark zones approximately 3 cm in diameter. Lesions can coalesce and, when a sufficient area of the leaf has become infected, the leaf will abscise. In the center of the lesion, signs of the fungus appear as immersed pycnidia, which liberate long, dense cirrhi of hyaline, ovoid spores. Biology and epidemiology. The fungus overwinters on infected plant tissue. Splashing rain or irrigation favors spread. It is known that high humidity is necessary for disease to occur. Bongini (1930) discovered disease would not occur when minimum and maximum temperatures were 10 –11  C (50–52  F) and 18–19  C (64–66  F), respectively. Management. Management strategies for Phyllosticta leaf spot are the same as for all leaf diseases. Practice sanitation and clean up from last year’s crop; scout and apply registered fungicides when disease is suspected. Chlorothalonil and mancozeb fungicides have preventive activity.

2.5

Pythium Root Rot (Pythium spp.)

Geographic occurrence and impact. Most reports on cyclamen have identified P. irregulare as the most commonly identified species (Moorman et al. 2002), but, in Germany, P. debaryanum was reported (Reimherr 1985). A report from Ohio found that P. aphanidermatum was the cause of root rot (Kuter et al. 1988). Regardless of the species, the disease can be very destructive when wet soil conditions prevail (Reimherr 1985). Symptoms/signs. Leaves initially develop a dull green color, lose turgor and then collapse. Many times they twist and curl. The root system darkens and feeder roots collapse. Roots frequently slough off the outer cortical tissue leaving the inner vascular stele, which gives rise to a rat-tail appearance. When seedlings are attacked, the symptoms can appear as damping-off disease.

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Biology and epidemiology. Pythium spp., like Phytophthora spp., are not true fungi but oomycetes, requiring free water to release zoospores. Consequently, the disease is favored by wet conditions, and spread of the pathogen requires irrigation water (Moorman et al. 2002). Pythium spp. commonly persist on the floors of greenhouses in organic debris; their long-term survival spores called chlamydospores and sexually produced oospores can remain dormant without a susceptible host. These propagules germinate in response to plant root exudates and invade susceptible roots. Fungus gnats (Bradysia spp.) likely assist Pythium by creating infection sites through their root feeding. (See chapter “▶ Insect Management for Control of Diseases of Florists’ Crops” for additional information.) Management • Cultural practices – Pythium root rot can be suppressed effectively by improving water drainage. Providing partial saturation of the growing medium can also reduce Pythium root rot (Elmer et al. 2011). Growers should plant in pasteurized media and immediately discard infected plants. Avoiding excess nitrogen has been shown to be important for Pythium control in geraniums (Gladstone and Moorman 1989, 1990). • Fungicides – When outbreaks appear, many fungicides are effective. In California, suppression of root rot was achieved following drenches with metalaxyl, followed by metalaxyl + benomyl and fenaminosulf (Raabe and Hurlimann 1981). Spengler (1979) found that furalaxyl successfully controlled P. debaryanum on cyclamen. Growers should rotate fungicides to prevent resistant strains of Pythium pathogens from developing. Mirkova (1995) reported in Bulgaria that Pythium isolates from Cyclamen showed tolerance to metalaxyl in 1988–1990. Growers using mefenoxam (the biologically active enantiomer of metalaxyl) against Pythium diseases in potted plant production today will want to rotate with other effective fungicides such as etridiazole and cyazofamid. • Biological control – Some advances have been made with biological control of Pythium root rot of ornamentals, but information on cyclamen is not available.

2.6

Phytophthora Root Rot of Cyclamen (Phytophthora tropicalis Aragaki) & Uchida, syn. Phytophthora capsici, Leonian

Geographic occurrence and impact. The pathogen was first described as Phytophthora capsici but was redescribed as P. tropicalis (Aragaki and Uchida 2001; Donahoo and Lamour 2008; Oudemans and Coffey 1991). The disease is not commonly found in cyclamen operations but can be devastating when it appears. Outbreaks have been reported from the United States, the Netherlands, and Germany (Gerlach and Schubert 2001). The disease caused economic losses in the late 1990s in Europe when it first appeared. The pathogen presumably entered the European operations on plant material since the fungus has no reported hosts in Europe. In the United States, P. tropicalis has been reported causing root rots on greenhouse and woody ornamentals (Olsen and Benson 2010).

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Fig. 9 Phytophthora blight (Courtesy of Diffusion Morel Co.)

Symptoms/signs. When the seeds are infested or are planted into infested soils, damping-off can occur. In older plants, the first symptom noticed is a change in leaf color, which turns from a bright, dark green to an off-colored, olive green (Fig. 9). Leaves become chlorotic and wilt followed by dark browning at the leaf base. The roots and stems eventually turn black and collapse even though the corms usually do not exhibit vascular discoloration. Biology and epidemiology. Phytophthora spp., like Pythium spp., are not true fungi but are oomycetes that will release their spores (zoospores) in free water. Consequently, a major requirement for spread and disease development is wet conditions. P. tropicalis can spread very effectively in irrigation water (Hong et al. 2006). Infection occurs on young roots where the pathogen invades and grows intercellularly, collapsing the cells while it grows. Optimal temperatures for disease development are between 20  C and 25  C (68–77  F), but the disease can also develop in cool conditions (15  C/59  F). During periods of optimal environmental conditions, the pathogen remains asexual. When moisture is limited, these spores germinate to produce infecting hyphae, whereas if free water is available, then sporangia and zoospores are produced. Once the disease has killed the plant or when conditions become unfavorable, the pathogen begins producing its sexual oospores. Species of Phytophthora are not recovered on greenhouse floors as frequently as Pythium, but the pathogen can persist in infested soils and on infested tissue for extended periods of time as oospores, chlamydospores, or mycelium. In the United States, the pathogen is reportedly causing disease on other ornamentals besides cyclamen, so growers should be aware of new sources of inoculum for cyclamen that may enter their greenhouse on other plants (Hong et al. 2006; Olsen and Benson 2010). Management. Management of Phytophthora root rot is the same as for Pythium root rot. Avoid overwatering and improve soil drainage to prevent development of the disease. Inspection and frequent scouting of new material are important given that the European outbreak presumably occurred from the importation of infested material. Fungicides that are effective against Pythium root rot will also suppress Phytophthora root rot (Aragaki and Uchida 2001). Strobilurins and phosphorous acid compounds are more effective against Phytophthora than against Pythium spp.

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Fig. 10 Powdery mildew of cyclamen caused by Pseudoidium cyclaminis (Photo courtesy of Wade Elmer)

2.7

Powdery Mildew of Cyclamen (Pseudoidium cyclaminis (Wenzl) U. Braun & R.T.A. Cook)

Geographic occurrence and impact. Powdery mildew is rarely observed on cyclamen. However, breeding programs developing new hybrids could alter susceptibility to powdery mildew, so the disease may become more important in the future. Powdery mildew on cyclamen has been reported from Europe. Symptoms/signs. The disease appears as small powdery lesions that appear on petals only (Fig. 10). Infections can reduce or destroy the value of the plant. Biology and epidemiology. The spores of this fungus spread by the wind during dry periods. The infection occurs during warm, humid weather. Temperatures of 21–25  C (70–77  F) are favorable to disease development, fostering spore formation and germination. Management. This disease is rare; it can be suppressed with chemicals when encountered. Test chemicals for phytotoxicity to cyclamen before using.

2.8

Ramularia Leaf Spot, Cyclamen Stunt Disease (Ramularia cyclaminicola Trel.)

Geographic occurrence and impact. The disease was first called cyclamen stunt, but Ramularia leaf disease was later shown to be caused by the same pathogen, Ramularia cyclaminicola. The fungus was first described as Cladosporium cyclaminis but was reclassified in 1950 as Ramularia cyclaminicola (Baker et al. 1950; Davis 1950). It was first observed in Illinois in 1914 (Trelease 1916) then in New York in 1926. Since then, outbreaks were reported in California, Colorado, Massachusetts, Ohio, Pennsylvania, New Jersey, and Quebec (Massey and Tilford 1932). Before 1950, the disease was widely distributed in North America causing moderate to severe losses, but the occurrence of disease is not significant in the horticultural trade today. However, since the original source of inoculum was speculated to be a native plant in the Primulaceae, growers should be aware of the possibility of its reoccurrence.

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Symptoms/signs. Ramularia cyclaminicola is a fungus causing systemic infection. The disease can manifest as a stunt, a leaf spot, or both. Smaller plants with small chlorotic leaves, short petioles, and short peduncles that force the flower to open within the plant canopy are typical of the stunt disease. Reddish-brown necrotic areas occur in the corm tissues and can extend up into the petioles, peduncles, and down into the primary roots. The older leaves of mature plants can develop diffuse yellow lesions and eventually wilt. The leaf spot disease is usually most obvious on the lower surface of leaves, where brown spots with indefinite margins that coalesce are seen. The fungus usually sporulates on the undersurfaces of the lower leaves but can also be recovered from the vascular tissue. Biology and epidemiology. The fungus survives as sclerotia (long-term survival structures) in soil and on plant debris. The pathogen can be spread by soil on infested pots, on airborne spores, and on seedlings and seeds. Following infection, the leaves die and sclerotia form and serve as the overwintering or survival structure. They germinate when susceptible tissue is available and the infection cycle begins again. The disease is favored by hot, humid weather . Management. Destroy affected plants and use new or disinfested growing medium. The fungicides effective against Phyllosticta and Septoria leaf spots should be effective against cyclamen leaf spot/stunt.

2.9

Rhizoctonia Crown and Stem Rot [Rhizoctonia solani Kühn (Teleomorph: Thanatephorus cucumeris (A.B. Frank) Donk)]

Geographic occurrence and impact. Rhizoctonia root rot can be a serious disease, but it appears to be restricted to isolated outbreaks. The pathogen is found globally so the disease can be common in all cyclamen-producing areas. Symptoms/signs. The fungus can infect all stages of the plant, beginning with a seedling damping-off and extending to a web blight that can colonize and infect mature aboveground tissues when humidity levels are very high. The most commonly observed symptom, however, is a root and stem rot. Once the pathogen invades the tissue, reddish-brown lesions appear, which over time can expand. Infected stems and petioles frequently are girdled and fall over. Portions of root systems may also turn brown. A common appearance on foliage of plants affected by Rhizoctonia attack is the look of nitrogen deficiency (yellowing of older leaves), often accompanied by the collapse of the aboveground tissue. Biology and epidemiology. R. solani can attack many different plants, and it can be a common resident in greenhouses. It is found in field soil, where it persists for years as resting spores called sclerotia or as a saprophyte on plant debris. R. solani has strong competitive saprophytic ability, meaning that it can grow quickly through soil if plant debris is present and the environmental conditions are suitable. The fungus does not spread in irrigation water or wind unless contaminated soil particles are being distributed: It does not have spores. Insect transmission is not common. The main sources of inoculum are non-pasteurized soil and infected plants. R. solani

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infects a wide range of ornamentals so inoculum can enter the production cycle any time new plant propagules are brought into the greenhouse. Management • Cultural practices – Given the wide host range of R. solani and its propensity for survival in soil, it would be safe to assume that all unpasteurized soil is a potential source of inoculum. Sanitation should be the first line of defense. • Fungicides and biocontrols – Fungicides also remain an essential tool for managing Rhizoctonia root rot including fludioxonil, PCNB, strobilurins, and thiophanate-methyl. Of the biological controls tested against Rhizoctonia root rot, the bacterium Serratia marcescens has been shown to be effective (Someya et al. 2000). Commercial formulations of Trichoderma harzianum have also been labeled for use on cyclamen.

2.10

Septoria Leaf Spot (Septoria cyclaminis Durieu & Mont.)

Geographic occurrence and impact. The fungus was first described in 1849 and has been reported from Central and Eastern Europe and the Middle East (Bacigálová et al. 2010). Septoria leaf spot is not a disease that causes major problems in the industry, but it can occasionally become problematic in some greenhouses. It was recently observed on a perennial cyclamen, C. fatrense, in Slovakia (Bacigálová et al. 2010). Symptoms/signs. The fungus first infects the undersides of the leaves. From above, red concentric spots are observed, which turn gray with red borders. There can be a fairly large number of leaf blotches, and they may dry up the leaf. Within the centers of the lesions, there will appear small, peppery dots called pycnidia, which bear the spores. Symptoms can be confused with those caused by Phyllosticta cyclaminis, which also produces spores in pycnidia. Spore shape distinguishes the two pathogens. Biology and epidemiology. The fungus spreads similarly to Phyllosticta cyclaminis. When spores land on the leaf and germinate, hyphae invade the plant through the stomata on the undersides of the leaves. Once established, the fungus develops in the intercellular spaces. In damp weather, the fruiting bodies exude elongated, multicelled conidia, which are spread by water, on tools and clothes. This disease spreads very fast. The fungus can survive as fruiting bodies on plant debris. Following a 12-h period when free water is on the leaf, spores will germinate in about 20 h at 18  C (64  F). Disease develops quickly when humid conditions prevail. Optimal temperatures for disease occur between 20  C (68  F) and 27  C (81  F). Management. Management of Septoria leaf spot of cyclamen is the same as for Phyllosticta leaf spot. For cultural control, reduce the length of time that the leaf surface sits wet.

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Fig. 11 Symptoms of Thielaviopsis black root rot on cyclamen roots (Courtesy of Diffusion Morel Co.)

2.11

Thielaviopsis Black Root Rot [Thielaviopsis basicola (Berk. & Broome) Ferraris (1912)]

Geographic occurrence and impact. Like Rhizoctonia root rot, Thielaviopsis root rot occurs sporadically. The disease has been reported in Europe and the United States. Thielaviopsis root rot appears to be a seedling problem, and, like many soilborne problems, outbreaks may be restricted to certain batches of seedlings. When the disease is present, losses can be severe. Symptoms/signs. A distinct black discoloration occurs in the small feeder roots, with a blackened root system seen in more advanced cases (Fig. 11). Infected plants are stunted, turn yellow, wilt, and die. Yellowing of the young leaves is seen when Thielaviopsis attacks the young roots. Biology and epidemiology. The fungus infects young roots and penetrates between cells. Discontinuous regions of root discoloration can be observed on the root system. T. basicola produces black, thick-walled, resistant chlamydospores as well as endoconidia that function as propagules for dissemination. The fungus grows best between 13  C (55  F) and 18  C (64  F) and enjoys very damp soil conditions; it is inhibited by high temperatures, ammonium fertilization, and acid (pH < 5.5) soils (Harrison and Shew 2001). The fungus is worldwide in distribution and infects a broad range of cultivated plants in many unrelated families. Fungus gnats and shore flies can spread black root rot.

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Management. Cultural practices can be effective in suppressing Thielaviopsis root rot of cyclamen. Soil pH greater than 5.5 is more conducive to disease, but acid soils can cause other problems for cyclamen, so growers should not rely on pH management. Proper diagnosis should be done to ensure the pathogen is T. basicola before taking control actions. Sanitation is also critically important in eliminating black root rot following outbreaks. Disinfesting seed trays and pots and using pasteurized soil mixes should be done following an infestation. Fungicides work best when used as protectants in combination with cultural controls and when applied early in the production cycle. Fungicides that have activity against T. basicola are fludioxonil, polyoxins, strobilurins, triflumizole, and thiophanatemethyl. Thiophanate-methyl has provided the best control in many studies, but this material should be used in rotation with others to avoid the development of fungicide resistance in the T. basicola population within a greenhouse.

3

Bacterial and Phytoplasma Diseases

3.1

Bacterial Corm and Leaf Rots

Pantoea agglomerans (Ewing & Fife) Gavini, Mergaert syn. Erwinia herbicola (Gavini, Mergaert); Pectobacterium carotovorum (Jones) Waldee syn. Erwinia carotovora; Dickeya dadantii subsp. dieffenbachiae (Samson et al.; Brady et al.), syn. Erwinia chrysanthemi; and Erwinia rhapontici Geographic occurrence and impact. Erwinia corm rot is observed in most production facilities. Reports of severe losses have come from Australia (Chandrashekar and Diriwaechter 1984), Europe (Bonifacio 1960; Carta 1993; Lemattre 1973; Panagopoulos and Psallida 1970), Asia (Amani 1967), and North and South America (Beamont 1953; Romero and Rivera 2005). This disease can become widespread in a production facility and cause considerable damage if not contained. Symptoms/signs. Symptoms are first observed in the leaves, but this is usually a reflection of advanced infection in the corm tissue. Leaves wither and droop abruptly onto the pot with blackened, flaccid stems as the transport of water and nutrients is blocked (Fig. 12). The petiole can also develop oozing lesions that form at the junction of the leaf and the petiole. Eventually, the disease advances and colonizes the whole plant as it travels upward. Many times, an unpleasant fishy odor accompanies diseased plants. Biology and epidemiology. Many bacteria in the genus Erwinia have been assigned new names, but the problems they cause are the same that were encountered in the last century. In general, all of these soft rot bacteria have the ability to dissolve pectin and other structural components in the plant, so infected tissues collapse and the bacteria utilize the plant material as food. The bacteria cannot directly penetrate plant tissue so they enter through stomata, wounds, and cracks in the corm – and at the sites where leaves or buds have been removed. Once inside the plant, tissues are soft rotted as the bacteria multiply and produce enzymes to break down the plant

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Fig. 12 Bacterial corm rot of cyclamen (Photo courtesy of University of Maryland Extension Service)

material; they may also spread throughout the plant through the vascular system. The bacteria can survive in the soil, in irrigation water, and in plant debris. Cyclamens that are planted too deeply are especially prone to bacterial soft rot, because the pathogens do well in an anaerobic environment. Fungus gnat infestations have been seen to aggravate problems with bacterial soft rot, as larvae feeding on the corms make wounds that facilitate entry of the bacteria. Management. Following an outbreak, all infected tissue should be discarded. A sanitary cleanup should be undertaken, since the pathogen has the ability to persist in soil and within plant debris. Once the operation has been sanitized, close inspection of incoming seedlings should be done to minimize reentry into the establishment. The disease tends to be more severe in warm, humid weather, since the bacterium multiplies more readily at high temperatures (between 25  C (77  F) and 30  C (86  F). Overfertilizing the plant increases disease severity by producing more young, succulent, susceptible tissue. Careful roguing of infected plants is important since billions of bacteria are released into the soil from infected plants. Minuto et al. (2004) had success using copper sulfate sprays to suppress P. carotovorum.

3.2

Ralstonia Wilt (Ralstonia solanacearum (Smith 1896) Yabuuchi et al. 1996, comb. nov.)

Geographic occurrence and impact. Bacterial wilt caused by Ralstonia solanacearum (formerly known as Pseudomonas solanacearum and now considered a heterogeneous species complex) has a very wide host range including cyclamen.

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Fig. 13 Symptoms of phyllody (the abnormal development of floral parts into leafy structures and virescence (green coloration in flowers) (Photo courtesy of Diffusion Morel Co.)

Although outbreaks on cyclamen are rare, the potential exists for losses. Were this pathogen to appear in cyclamen, it could be a matter of regulatory concern, as some pathogen strains (race 3, biovar 2) also cause losses in important vegetable crops. R. solanacearum causes latent infections, which may slow its detection in new or uncommon hosts. Biology and epidemiology. Much of the information on the biology and epidemiology for bacterial corm and leaf rot diseases of cyclamen applies to the Ralstonia disease.

3.3

Phytoplasma Diseases

Geographic occurrence and impact. Phytoplasma diseases on cyclamen are rare. One report in 1990 cited an occurrence in northern Italy and Germany (Bertaccini 1990). However, it is likely that the disease has appeared many times before and merely gone unreported. Symptoms/signs. Symptoms on cyclamen were observed as phyllody (the abnormal development of floral parts into leafy structures) and virescence (green coloration in flowers) (Fig. 13). Stunting with thickened, rolled leaves has also been associated with the disease (Fig. 14). Biology and epidemiology. Phytoplasmas are prokaryotes like bacteria, but they are obligate plant parasites vectored by phloem-feeding leafhoppers. Based on molecular analysis, the cyclamen phytoplasmas belong to 16S rDNA subgroups 1B and 1C (Alma et al. 2000). According to RFLP molecular analysis, the cyclamen phytoplasmas are in the same subgroup (1B) as aster yellows phytoplasmas. The cyclamen phytoplasma has been placed in the stolbur phytoplasma group along with the phytoplasmas detected in apricot, grape, lisianthus, papaya, and pepper (Weintraub et al. 2007). The disease is only transmitted by leafhopper feeding, and there is only one report of cyclamen serving as a reservoir for inoculum. In that study only one plant out of 366 plants tested was infected by leafhoppers that had fed on infected cyclamens. The phytoplasma disease on cyclamen was considered a dead

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Fig. 14 Phytoplasma disease of cyclamen causing stunting with thickened, rolled, cup-shaped leaves (Photo courtesy of Diffusion Morel Co.)

end for the pathogen, since transmission appeared to be extremely rare (Alma et al. 2000). Management. Given the rarity of the disease, it is not likely that a major outbreak will occur. Management, however, is achieved through monitoring and the reduction of the insect vector, which would not typically be found in greenhouses. Additional information on vector management may be found in introductory chapter “▶ Insect Management for Disease Control in Florists’ Crops.” Symptoms/signs. Symptoms are similar to bacterial corm rot. Wilting and stunting are first observed in the aboveground tissue, but this reflects advanced infection in the corm tissue. Leaves wither and droop abruptly onto the pot. Management. Refer to bacterial corm and leaf rot diseases of cyclamen.

4

Viral Diseases

4.1

Tomato Spotted Wilt Virus: TSWV

Geographic occurrence and impact. Tomato spotted wilt virus (TSWV) is considered to be one of the worst viral threats on cyclamen. It belongs to the genus Tospovirus in the family Bunyaviridae and has a very wide host range. The virus was reported in Europe in 1932 but seemed to have disappeared. It reappeared in the ornamentals industry in 1986 when the thrips Frankliniella occidentalis, one of its most effective vectors, became widespread in the horticultural trade. It still constitutes one of the most prevalent viruses in the global greenhouse industry (Daughtrey et al. 1997; Kamińska 1975; Resende et al. 1996; Vozelj et al. 2003). Symptoms/signs. The level of damage can vary widely depending on the environmental temperatures and cyclamen cultivar. In general, plants are stunted with leaf mosaics, discoloration, and necrotic patches (Fig. 15). Round brown spots are typical on more mature cyclamen. Line patterns and various chlorotic and necrotic symptoms associated with veins are seen. It is impossible to distinguish TSWV from Impatiens necrotic spot virus, another tospovirus, by symptoms, so proper diagnosis needs to be done by a professional laboratory using serological or molecular assays. Biology and epidemiology. Over 500 species play host to this virus, including ornamentals, vegetables, and weeds. Contamination is only through the feeding of

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Fig. 15 Tomato spotted wilt virus on cyclamen (Courtesy of Diffusion Morel Co.)

the thrips vectors, since the virus is not seed transmitted. Only six species of thrips are known to carry the virus; of these, the Western flower thrips (Frankliniella occidentalis) is the most common one in most greenhouses. Only the first and early second larval stages are able to acquire tospoviruses, and only immature thrips that acquire these viruses or adults derived from such immatures are vectors. Adult thrips remain viruliferous for life; but tospoviruses are not transovarial. Incubation of the virus in the cyclamen plant takes around 2 months, weeks longer than for some other flower hosts. Management. Control of the thrips vector is the single most important strategy, which should be focused on while maintaining effective scouting and sanitation practices. In the most stringent management program, one should quarantine new stock and seedlings for at least 2 months, until plants appear to be clean; prevent thrips from developing on these plants while they are in quarantine. Remove old flowers as they serve as a source of pollen for thrips. Sterilize tools used in cutting old flower peduncles as plant sap could possibly transmit the virus (it is not known to be easily sap transmitted, however). All infected plants must be destroyed and removed from the greenhouse promptly. Eradicate weed hosts and any other potentially infected host crops to eliminate inoculum reservoirs. More information can be found in the introductory chapter ▶ “Insect Management for Disease Control in Florists’ Crops.”

4.2

Impatiens Necrotic Spot Virus: INSV

Geographic occurrence and impact. Impatiens necrotic spot virus (INSV) is a very common viral disease in greenhouses, with a wide host range encompassing hundreds of flower crops. It is reported from Europe (Bellardi and Vicchi 1998); Japan (Goto et al. 2001), South America (Resende et al. 1996), New Zealand (Elliott et al. 2009), and North America (Daughtrey et al. 1997). Within North America, this is the tospovirus most often encountered in greenhouses, whereas the closely related tomato spotted wilt virus (TSWV) is more often found outdoors.

Diseases of Cyclamen

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Fig. 16 Impatiens necrotic spot virus on cyclamen (Courtesy of Diffusion Morel Co.)

Symptoms/signs. Leaf symptoms on cyclamen are sometimes very distinctive, with the production of ring spots, chestnut, or yellow in color, sometimes seen at the margin of the leaf as “fingerprints” (Fig. 16). The virus can also manifest these symptoms on the flower petals. As with TSWV, infected and damaged cells do not expand and grow at the same rate as healthy cells so distortion and leaf deformity develop over time. The virus can cause wilting, stem death, stunting, yellowing, poor flowering, and sunken round, brown spots on leaves along with the characteristic ringspots. In some situations, the plant may remain asymptomatic. Positive diagnosis is achieved only after samples are assayed using diagnostic kits or serological assays in a laboratory that can specifically identify the virus. Biology and epidemiology. INSV belongs to the genus Tospovirus in the family Bunyaviridae and acquisition and transmission of the virus by thrips is the same as with TSWV. The incubation period within the plant is 2 months. Management. Controlling the thrips vector is a key management strategy. Efforts to strongly suppress the thrips population will reduce the threat of viral infection. Eradicating weed hosts and making certain that other crops are not infected will reduce inoculum reservoirs. Scout regularly and destroy unnecessary flowers, so as to limit potential refuges and pollen sources for thrips. More information can be found in the introductory chapter “▶ Insect Management for Disease Control in Florists’ Crops.”

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4.3

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Cucumber Mosaic Virus

Geographic occurrence and impact. Cucumber mosaic virus (CMV) infects a wide range of host plants including over 100 ornamentals (Korbin and Kaminńska 1998). It can cause problems in isolated greenhouses, but it is not as devastating as the tospoviruses. Symptoms/signs. Cyclamen plants infected with CMV frequently look stunted and chlorotic, with mosaic patterns of yellow and green. Leaves can also be asymmetric (Kamińska 1975; Minuto et al. 2011). Biology and epidemiology. CMV is a single-strand RNA virus that belongs to the genus Cucumovirus in the family Bromoviridae. It is transmitted by certain species of aphids in the non-circulative (nonpersistent) mode, meaning the virus does not survive for long periods of time in the insect. Once the aphid has acquired the virus, it can transmit it to any susceptible host for a limited amount of time. Seed transmission of CMV also occurs, but data on seed transmission in cyclamen is not available. Transmission of CMV through seed can be as much as 50% in some plants (Gallitelli 2000). Management. Management of viruses, including CMV and other species in the Bromoviridae, is based on monitoring and reduction of vector populations through chemical or biological measures, excluding vectors by the use of physical barriers such as nets or screens and eliminating alternate hosts – especially established, infected crops. Aphid populations are generally not tolerated during cyclamen production, so scouting to find the rare occurrences of the disease should be the cornerstone of managing this disease. Test kits (and diagnostic laboratory analysis) are available to test for the presence of CMV, allowing the grower to distinguish it from the more common thrips-vectored TSWV and INSV. Resistance to CMV has been introduced into a wide variety of vegetables through genetic modification. Additional information on integrated disease management of viruses may be found in introductory chapter “▶ Insect Management for Disease Control in Florists’ Crops.”

4.4

Tobacco Mosaic Virus (TMV)

Geographic occurrence and impact. Tobacco mosaic virus (TMV) has a global distribution and can infect a wide range of plants. This virus is a single-strand RNA virus belonging to the genus Tobamovirus and family Virgaviridae. Outbreaks are rare and localized in cyclamen. Symptoms/signs. TMV causes mosaic patterns on leaves. Biology and epidemiology. TMV is easily mechanically transmitted and can persist in plant debris for 50 years. Management. Inspect incoming stock and keep it segregated from older plants. Rogue out infected plants and practice strict sanitation. Workers should frequently wash hands and disinfest cutting tools. Powdered milk solutions may be used to inactivate TMV in some instances.

Diseases of Cyclamen

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25

Nematodes

Cyst [Heterodera radicicola (Greeff) Mull.], root knot (Meloidogyne spp.), stem and bulb (Ditylenchus spp.), lesion (Pratylenchus spp.), and foliar nematodes (Aphelenchoides spp.). Geographic occurrence and impact. Different nematodes can attack cyclamen depending on region and cultivation practices. However, the movement by the industry to use soilless potting mixes has essentially eliminated damage from soilborne nematodes (cyst nematodes, root-knot nematodes, lesion nematodes, and stem and bulb nematodes). Isolated outbreaks have been reported but seem to be associated with nonsterilized soil (Butcher 1934; Reimherr 1985). However, outbreaks of foliar nematodes (Aphelenchoides spp.) may still cause damage in cyclamen, although this has been rarely reported. Symptoms/signs. Roots infected with nematodes can exhibit a range of symptoms depending on the pathogen. Aboveground symptoms appear as stunting and yellowing and in severe cases wilt and death. Root symptoms can also vary. Rootknot nematodes cause galls, while lesion nematodes cause stunted roots that have distinct reddish-brown lesions. Stem and bulb nematodes attack the corm and can cause wilt. Foliar nematodes (Aphelenchoides spp.) can invade leaves, leaf buds, and meristems. Due to cyclamen’s thick, dense leaves, foliar nematodes do not show the clear interveinal demarcation that is commonly seen in plants with thin leaves. Instead, cyclamen infestation with Aphelenchoides shows as yellowish patches, which over time turn brown or blackish. The leaves dry up and often remain attached to the petioles. Affected plants are unmarketable. Biology and epidemiology. Parasitic nematodes are microscopic worms that possess stylets for piercing plant cells and extracting nutrients. Different types of nematode may infect the roots, stems, or leaves of their host plants. Root-knot nematodes (Meloidogyne spp.) and cyst nematodes (Heterodera spp.) infect roots and establish feeding sites (nurse cells) that develop into visible galls and cysts, respectively. Root-knot and cyst nematodes do not migrate and are considered sedentary endoparasites. Lesion nematodes invade roots but migrate within the root system feeding on cells (migratory endoparasite). Stem and bulb nematodes (Ditylenchus spp.) invade at the soil line and migrate within the tissue (migratory endoparasite). Foliar nematodes (Aphelenchoides spp.) are also migratory endoparasites. Nematodes survive as eggs that overwinter in soil or can be introduced on host material. Handling and splashing water can spread the foliar nematode and stem and bulb nematode between crops or within an infested crop. The movement of infected crops or contaminated soil is usually necessary to spread the other kinds of nematodes. Management. If unsterilized soil is used in the potting medium, the potential for root infection by nematodes is high. Growers should be vigilant in examining roots for stunting and poor growth. Although the use of soilless potting mixes will eliminate most soilborne nematode threats, foliar nematodes usually enter a production facility on other crop hosts. Careful examination of incoming plant material

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should be practiced. Common hosts of foliar nematodes are ferns and African violets (Daughtrey et al. 1995), as well as numerous herbaceous perennials.

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Diseases of Daffodil (Narcissus) Gordon R. Hanks and Gary A. Chastagner

Abstract

Daffodils (Narcissus) are popular spring flowerbulbs in Europe, North America, Australia, and New Zealand, and tazetta narcissi are widely grown in China, Israel, and Japan. Large volumes of field-grown cut-flowers are traded, along with forced flowers and pot-grown plants from greenhouses, as well as the bulbs themselves. Basal rot (due to Fusarium oxysporum f. sp. narcissi) causes substantial losses, while the stem nematode (Ditylenchus dipsaci) is capable of devastating crops and is an ever-present concern; both are controlled by a hot-water treatment incorporating biocide and/or fungicide, but husbandry and cultural factors also play a major role. Biological control is in its infancy, but some progress is being made against basal rot using soil applications of composts that increase the population of beneficial soil microorganisms. Fungal foliar diseases reduce yields and can affect flower quality – smolder (Botrytis narcissicola), white mold (Ramularia vallisumbrosae), fire (Botrytis (Botryotinia) polyblastis), and leaf scorch (Peyronellaea (Stagonospora) curtisii) – and are controlled by a fungicide spray program. Bulb rots are also caused by species of Penicillium and Rhizopus. Instances of bacterial disease are rare, but Pectobacterium (Erwinia) carotovorum subsp. carotovorum attacks bulbs of tazetta narcissi. Daffodils are widely infested with multiple aphid-borne potyviruses, in particular Narcissus Late Season Yellows Virus, Narcissus Yellow Stripe Virus, and (in tazetta narcissi) Narcissus Degeneration Virus, reducing bulb

G.R. Hanks (*) Warwick Crop Centre, Wellesbourne Campus, University of Warwick, Wellesbourne, Warwickshire, UK e-mail: [email protected] G.A. Chastagner Department of Plant Pathology, Washington State University Research and Extension Center, Puyallup, WA, USA e-mail: [email protected] # Springer International Publishing AG 2017 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florists' Crops Diseases, Handbook of Plant Disease Management, DOI 10.1007/978-3-319-32374-9_43-1

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and flower yields and flower quality. Control of viruses is largely limited to crop inspection and the removal of affected plants. Keywords

Basal rot • Biocide • Botryotinia • Botrytis • Daffodil • Ditylenchus • Fungicide • Fusarium • Hot-water treatment • Narcissus • Nematode • Penicillium • Peyronellaea • Ramularia • Rhizopus • Stagonospora • Virus

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Basal Rot (Base Rot, Fusarium Rot) [Fusarium oxysporum f. sp. narcissi Snyder & Hansen (syn. F. bulbigenum Cooke & Massee)] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Fire [Botrytis polyblastis Dowson (Teleomorph: Botryotinia polyblastis (Greg.) Buchw.) (syn. Sclerotinia polyblastis Greg.)] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Leaf Scorch [Peyronellaea curtisii (Berk.) Aveskamp, Gruyter & Verkley (syn. Stagonospora curtisii (Berk.) Sacc.)] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Neck Rot [Fusarium oxysporum f. sp. narcissi Snyder & Hansen (syn. F. bulbigenum Cooke & Massee), and Other Fungi] . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Penicillium Rot (Penicillium spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 Smolder [Botrytis narcissicola Kleb. (Teleomorph: Botryotinia narcissicola (Greg.) Buchw.; syn. Sclerotinia narcissicola Greg.)] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7 Soft Rot (Rhizopus spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.8 White Mold (Ramularia vallisumbrosae Cav.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.9 White Root Rot (Rosellinia necatrix Prill.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.10 Other Fungal Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Bacterial and Phytoplasma Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Bacterial Rot [Pectobacterium carotovorum subsp. carotovorum (Jones) Hauben et al. (syn. Erwinia carotovora subsp. carotovora (Jones) Bergey et al.)] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Bacterial Streak (Pseudomonas sp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Diseases Due to Phytoplasmas . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Virus Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Alphaflexiviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Betaflexiviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4 Bromoviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5 Bunyaviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6 Potyviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7 Secoviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.8 Tombusviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.9 Virgaviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.10 Other Viruses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Nematode Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 Stem Nematode [Ditylenchus dipsaci (Kühn) Filipjev] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2 Aphelenchoides subtenuis (Cobb) Steiner & Buhrer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3 Root Rot and Soil Sickness: Root-Lesion Nematode [Pratylenchus penetrans (Cobb) Chitwood & Oteifa] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 Physiological Disorders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1 Chocolate Spot . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2 Rust . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

3 4 4 15 17 21 25 27 32 34 36 37 39

39 39 39 40 40 46 47 47 48 49 53 57 58 60 60 60 67 68 70 71 71

Diseases of Daffodil (Narcissus) 6.3 Grassiness . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.4 Disorders of Flower Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 Hot-Water Treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2 Temperature and Duration of HWT . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.3 Additives to HWT Tanks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.4 Timing of HWT . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.5 Damage Due to HWT . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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3 72 72 77 77 79 82 83 83 84 86

Introduction

Plants of the genus Narcissus (daffodils and narcissi) were known in antiquity, mainly as medicinal plants (Dweck 2002). Although the Chinese sacred lily (Narcissus tazetta var. chinensis) was appreciated in the Far East and the Levant from much earlier times, in the sixteenth century daffodils were imported to the Netherlands from Constantinople and thence to England by horticulturally astute Huguenots (Willis 2012). A “daffodil revival” in the nineteenth century led to the start of substantial hybridization in the British Isles (with Peter Barr and the Royal Horticultural Society greatly raising the profile of the genus) and the Netherlands (notably at de Graaff’s nursery): daffodils soon became a major floricultural crop. In the USA, the Plant Quarantine Act 1919 was extended in 1926 to ban the import of daffodils, creating a pre-emptive flood of imports, further hybridization, and a new crop for the Pacific Northwest (PNW) (Willis 2012). Currently some 4,000 ha of daffodils are field-grown in England (2008 figures; Defra/National Statistics 2009), 1,500 ha in the Netherlands (Bloembollenkeuringsdienst (BKD) 2016), 600 ha in Scotland (authors’ estimate from Scottish Government (2015) and commercial information), and 370 ha in the USA (authors’ estimate from USDA NASS (2016) and commercial information). Significant areas are also grown in Australia, Canada, New Zealand, Poland, and the Republic of Ireland and of mainly tazetta daffodils in China, Israel and Japan. Much of the information here is derived from advisory publications in the Netherlands (Bergman et al. 1978; van Aartrijk et al. 1995; van der Zwet et al. 1990), UK (ADAS 1984, 1986; Hanks 2013a; Hanks and Linfield 1994; Lane 1984; Moore et al. 1979), and USA (Byther et al. 1998; Gould and Byther 1979; Pscheidt and Ocamb 2015). Further material derives from standard textbooks (Brunt 1995; Byther and Chastagner 1993; Chastagner and Byther 1985; De Hertogh and le Nard 1993; Hanks 1993a, 2002; Kamenetsky and Okubo 2013; Melville 1980; Rees 1972, 1992; Smith et al. 1988). Databases consulted include “Descriptions of Plant Viruses” http://www.dpvweb.net, “Distribution Maps of Plant Diseases” http:// www.cabi.org/dmpd/, “Fungal Databases” http://nt.ars-grin.gov/fungaldatabases/ (Farr and Rossman 2016), “Plant Viruses Online” http://srs.im.ac.cn/vide/refs.htm, and “Species Fungorum” http://www.speciesfungorum.org/Names/Names.asp. Note that the term “daffodil” is used to include all types of Narcissus: in most cases the

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reference is to the typical or mainstream cultivars of commerce, but when referring to a particular type of daffodil (e.g., tazetta species and cultivars) this is specified. Referred to by their active ingredient (a.i.) names, pesticides and other chemicals mentioned in the text are used to report the findings of research and trials – they are neither recommendations nor statements of current approval, availability, or legality.

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Fungal Diseases

2.1

Basal Rot (Base Rot, Fusarium Rot) [Fusarium oxysporum f. sp. narcissi Snyder & Hansen (syn. F. bulbigenum Cooke & Massee)]

Geographic Occurrence and Impact Basal rot is a major problem in daffodilproducing countries (Chastagner and Byther 1985; Moore et al. 1979; van Aartrijk et al. 1995) and probably wherever daffodils are grown. Basal rot has also been reported from China (Xu et al. 1987) and Fusarium oxysporum f. sp. narcissi has been reported on daffodils in Australia, Bulgaria, Canada, Germany, Mexico, New Zealand, South Africa, and Zimbabwe (Farr and Rossman 2016). In the late-nineteenth century, daffodil bulb rot was a frequent subject of the horticultural press and attracted attention in the Netherlands and UK, especially in the exceptionally hot summer of 1911 when large numbers of bulbs rotted in both countries. Moore et al. (1979) provide an interesting history of the earlier investigations and how they were beset by poor interpretation and mischance, giving full references. Confusion was rife – was the rot caused by a nematode, a fungus, or something else? By the 1920s, UK researchers concluded that Fusarium was not involved or was at most a secondary phase of a nematode disease. Starting in 1926 large numbers of daffodils imported to the UK suffered from a rot that was apparently free of nematodes and similar findings emerged from Canada and the USA. In 1927–1932 research in the Netherlands, UK and USA finally showed that a Fusarium species – F. bulbigenum, now known to be F. oxysporum f. sp. narcissi – was the sole cause of basal rot, though it and stem nematode frequently occurred together. Weiss (1929, 1932) confirmed Fusarium as the cause of basal rot, and it was shown to be widely distributed, and of economic importance, across the USA, going on to investigate its management through hot-water treatment (HWT) or separate fungicide treatments. Similar conclusions were reached by others in the USA, Netherlands, and UK. The impact of basal rot has become more severe since the 1970s, perhaps due to changes in husbandry. Many aspects of contemporary bulb-growing exacerbate damage, such as reduced sorting by hand (more infested material planted), handling in bulk bins (rather than flats or trays), higher planting densities (20 t/ha rather than 10 t/ha), longer growing cycles (for 2 or more years instead of lifting, treating, and replanting annually), early lifting (before the bulb skins have “ripened”), bulb washing (when necessary to produce soil-free bulbs for export), and early planting

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(before soil temperatures have fallen) (Hanks 2013a). Now, increasingly robust restrictions and withdrawals of fungicides and biocides limit the opportunities for chemical management. Finally, basal rot is favored by warmer temperatures: once associated with warm summers, it now appears to be a permanent problem, perhaps exacerbated by climate change. The obvious effect of basal rot is reduced bulb yield, but the health of the bulb crop naturally has consequences for cut-flower and pot-plant production. In the UK and USA, where most flowers are picked from field crops, their yield and health will be directly dependent on that of the field crop. Bulbs lifted and sold for flower forcing, pot-plant production and retail sales do not have the HWT afforded to bulbs intended for growing-on as field crops, and bulb rot sometimes becomes apparent a few weeks after dispatch, damaging the perception of the product. Gaps due to rotting bulbs will reduce the yield of forced flowers, exacerbated by greenhouse temperatures that favor the development of Fusarium. Where pot-grown daffodils are produced, the failure or poor appearance of one bulb will downgrade the whole pot. Basal rot is a great threat to daffodil production because of its ubiquity and aggression and a long-term failure to find acceptable, effective methods for its management. In the UK, 30% losses in bulb yield have occasionally been known in seriously affected bulb stocks of basal rot-susceptible cultivars. Even when husbandry has been optimized in such bulb stocks over a number of years, it is difficult to reduce infection rates to below 2% (Hanks 1996). For calculating the benefits of effective basal rot management, an increase in yield of about 10% has been considered reasonable. Symptoms/Signs After planting, depending on the extent of damage, roots may fail to emerge from the base rot or may emerge only to be affected later, becoming dark in color. In growth, affected bulbs produce short-lived, crooked shoots, or increasingly weak foliage that senesces prematurely several weeks before that of healthy plants. In the early stages, basal rot may be seen as a small dark spot in the base plate when the bulb is cut lengthways. The rot spreads through the base plate and then upwards through the bulb scales, which become characteristically moist and dark chocolate to reddish-brown in color – the color may vary with the cultivar (Fig. 1). It is important to distinguish these symptoms from those of stem nematode (brown rings when the bulb is cut through transversely) and the normal die-back of leaf bases (which does not show the typical leading edge of basal or neck rot). Once a significant portion of the bulb has been affected, its softness is apparent on handling. A pinkish-white spore and mycelium mass forms near the base plate (Fig. 2). As the bulb becomes dehydrated, perhaps aided by bulb mites moving in to feed on damaged tissue, it becomes brittle and mummified. Several weeks before bulb lifting, infected plants may already be obvious as their leaves die-down earlier than the rest. At lifting infected bulbs may have no sign of roots, and at or soon after lifting feel soft, especially around the base, and show a discolored area extending upwards if the outer scale is removed.

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Fig. 1 Typical upwards progression of daffodil bulb rot due to Fusarium oxysporum f. sp. narcissi (Published with kind permission of the University of Warwick # 2016. All Rights Reserved)

Fig. 2 Daffodil bulb with basal rot showing eruption of mold from base plate (Fusarium oxysporum f. sp. narcissi) (Published with kind permission of the University of Warwick # 2016. All Rights Reserved)

Biology and Epidemiology F. oxysporum f. sp. narcissi is found in most soils where daffodils have been grown and sometimes in soils where they have not, despite the pathogen being limited to this host (Apt 1958; Price 1975a, b). In soil the pathogen can remain viable for up to 10 years, at least under artificial conditions

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7

(CA Linfield, personal communication, 1990). Spores of both pathogenic and nonpathogenic strains have been found on healthy bulbs in storage (Price 1973). The disease can be bulb- or soil-borne and spread can occur in both stored and planted bulbs (Hawker 1940; Price 1977a). Spread of spores on air currents and via water (during bulb washing, dipping or HWT) would be expected. Variation in the pathogenicity of F. oxysporum f. sp. narcissi isolates was studied by Linfield (1997) and the cross-infectivity of its formae speciales by Rataj-Guranowska et al. (2007). Control is important during the first months after planting as the pathogen is active only above 13  C/55  F (Langerak and Haanstra-Verbeek 1977). In planted bulbs, infection occurs when the integrity of the bulb surface is compromised: when new roots are emerging, when old roots are dying-down, or when there is mechanical damage (Gregory 1932; Hawker 1935; Langerak and Haanstra-Verbeek 1977; Price 1975a, b, 1977a). Root infection and disease development are closely related to soil temperature above 13  C/55  F, reaching a maximum at 29  C/84  F (McClellan 1952), explaining the finding that the base plate can be penetrated via the senescing roots in summer if the soil is wet (Hawker 1935, 1943a). Root senescence starts in late-May and lasts until September, corresponding to the time when temperatures favor the pathogen’s active growth (Hawker 1943a; Price 1977a). Soil-borne infection is encouraged by high temperatures (Hawker 1935) and can occur late in the season when moribund roots are present (Hawker 1943a; Price 1977a), though it has also been reported in temperatures as low as 5  C/41  F (Price 1982). Little infection was recorded when inoculum was placed 30 cm below bulbs (Price 1975c). When healthy bulbs were placed 0, 10, or 20 cm laterally from infected bulbs, 60, 27, and 6% of bulbs, respectively, were affected after one growing season (Linfield 1987). It has been suggested that because of the relatively sparse spread of daffodil roots and the high concentrations of chlamydospores needed to produce infection, bulb-borne inoculum is more important than soilborne inoculum, especially with high planting rates, reduced sorting by hand, and 2-year-down growing (Price 1977a, b). While the pathogen can move only a little through soil, it can move along or in roots for some distance to infect bulbs (Price 1975a, b). The pathogen advances intercellularly before invading damaged cells (Bald et al. 1971). The nutritional state of the soil influences infection. Using excessive N fertilizer increased the incidence of basal rots, splits, and bruises in daffodil bulbs (Biekart 1930; McClellan and Stuart 1947). Applying no more than about 120 kgN/ha was considered satisfactory (Rikhter 1976). In trials, bulbs of ‘Golden Harvest’ (basal rot-susceptible) and ‘St Keverne’ (resistant) were planted in soil amended with the equivalent of up to 300 kgN/ha, together with no, low or high rates of basal rot inoculum. Using a split-dose of 150 kgN/ha at planting and 150 kgN/ha top-dressed in April increased basal rot in ‘Golden Harvest’ and, in combination with the higher rate of inoculum, in ‘St Keverne’ also (Hanks et al. 1998). The incidence of rotting bulbs, and the numbers of pathogen propagules isolated from the base rots of healthy bulbs, increased as storage temperature was raised from 15 to 24  C/59 to 75  F and then declined somewhat at 30  C/86  F (Price 1975a, b). When apparently healthy bulbs were taken from storage and grown for a year, the

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percentage rotting was similar in all treatments. Pathogenic isolates were found on the base rots of 21% of samples. Because of these effects of temperature, it is not surprising that storage of bulbs at 15  C/59  F after lifting prevents the development of many fungal lesions. The overall optimal temperature for bulb storage is 17  C/ 62  F; storage at 20 to 30  C/68 to 86  F should be avoided. Management Extension workers have long stressed that the available fungicide treatments alone are insufficient for managing basal rot and must be combined with good bulb handling and husbandry which exert a substantial effect in reducing the incidence of basal rot if used consistently (e.g., Hanks 2002, 2013a). The “physical” management of basal rot through correct bulb drying, storage, and HWT, cultural management through using appropriate planting sites, depth and timings, and the limited chemical controls available make an interesting example of integrated crop management (ICM), but one that needs to be extended by the development of biological controls and using the potential of plant breeding. HWT, a key element in basal rot management, is even more important for the control of stem nematode (Ditylenchus dipsaci), such that it is routine to treat all daffodil stocks in the run-up to planting. Bulbs are immersed in a tank containing a biocide and/or fungicide for, typically, 3 h at 44.4  C/111.9  F. Because HWT is a key to managing stem nematode as well as basal rot, the general features of HWT (such as the temperature, duration and timing of treatment) are discussed in Sect. 7 to save repetition, and only information specific to managing basal rot is given here. HWT also contributes incidentally to the management of other daffodil pathogens (e.g., leaf scorch) and pests (e.g., narcissus flies (Merodon equestris and Eumerus spp.), bulb mites (Rhizoglyphus and Histiostoma spp.), and bulb-scale mites (Steneotarsonemus laticeps). Colloquially, HWT is referred to as “sterilizing,” “bulb-dipping” or even “boiling,” terms avoided in this chapter! In disease management short, bulb-dip treatments in water at ambient temperatures may also be used, and to distinguish such treatments from HWT, they are referred to as “cold dips.” • Cultural – Alongside their work on bulb-dipping, both Gregory (1932) and Hawker (1940) demonstrated the importance of appropriate bulb storage as a means of alleviating basal rot. Both recommended that bulbs should be stored at temperatures