Effectors in Plant-Microbe Interactions E-Book

• Herausgeber: John Wiley & Sons
• Kategorie: Wissenschaft und neue Technologien
• Sprache: Englisch

Seitenzahl: 840

Veröffentlichungsjahr: 2011

Beschreibung

Plants and microbes interact in a complex relationship that can have both harmful and beneficial impacts on both plant and microbial communities. Effectors, secreted microbial molecules that alter plant processes and facilitate colonization, are central to understanding the complicated interplay between plants and microbes. Effectors in Plant-Microbe Interactions unlocks the molecular basis of this important class of microbial molecules and describes their diverse and complex interactions with host plants. Effectors in Plant Microbe Interactions is divided into five sections that take stock of the current knowledge on effectors of plant-associated organisms. Coverage ranges from the impact of bacterial, fungal and oomycete effectors on plant immunity and high-throughput genomic analysis of effectors to the function and trafficking of these microbial molecules. The final section looks at effectors secreted by other eukaryotic microbes that are the focus of current and future research efforts. Written by leading international experts in plant-microbe interactions, Effectors in Plant Microbe Interactions, will be an essential volume for plant biologists, microbiologists, pathologists, and geneticists.

Leseprobe

Contents

Cover

Title Page

Copyright

Contributors

Foreword

References

Preface

Section 1: Plant Immune Response Pathways

1: Innate Immunity: Pattern Recognition in Plants

1.1 Pattern Recognition through MAMPs (Microbe-Associated Molecular Patterns)

1.2 Some Classical MAMP-Receptor Pairs

1.3 Physiological Responses and Signaling Events Induced by Elicitors

1.4 The Biological Relevance of PTI

References

2: Microbial Effectors and Their Role in Plant Defense Suppression

2.1 The Gene-for-Gene Concept and the Emergence of Effectors

2.2 Diversity of Effectors

2.3 Effector Targets

2.4 Models to Explain Recognition of Effectors by R-gene Products

2.5 Synthesis and Discussion

References

Section 2: Genome-Wide Analyses of Microbial Effectors and Effector Evolution

3: Comparative Genomics and Evolution of Bacterial Type III Effectors

3.1 Introduction

3.2 Effector Structure

3.3 Effector Acquisition

3.4 Effector Change and Loss

3.5 Effector Repertoire Evolution

3.6 Future Prospects

References

4: The Effectors of Smut Fungi

4.1 Introduction

4.2 Plant Responses to U. maydis

4.3 The effectors of U. maydis

4.4 Regulation of U. maydis Effector Genes

4.5 Stage and Organ Specificity of U. maydis Effectors

4.6 The Effectors of Smut Fungi Related to U. maydis

4.7 Outlook

4.8 Acknowledgements

References

5: Evolutionary and Functional Dynamics of Oomycete Effector Genes

5.1 Introduction

5.2 Oomycete Effectors Target Different Sites in Host Plant Tissue

5.3 Oomycete Effectors have a Modular Architecture

5.4 Oomycete Effector Genes Show Distinct Patterns of Expression During Plant Colonization

5.5 Effector Genes Populate Plastic Regions of Oomycete Genomes

5.6 Evolution of P. infestans Genome and Effector Genes Following Host Jumps

5.7 Several Oomycete Effectors Suppress Plant Immunity

5.8 Effectors Are Useful in Breeding and Deployment of Disease Resistance

5.9 Outlook

References

Section 3: Microbial Effector Functions: Virulence and Avirulence

6: Suppression and Activation of the Plant Immune System by Pseudomonas syringae Effectors AvrPto and AvrPtoB

6.1 Pseudomonas syringae pv. tomato Interactions with Plants

6.2 AvrPto and AvrPtoB Have Both Redundant and Unique Activities in Plants

6.3 AvrPto is a Small Effector with Two PTI-Suppressing Domains Both of Which Can Activate ETI in Certain Solanaceous Plants

6.4 AvrPtoB is a Large Modular Effector with Domains that Suppress PTI and ETI but Which Also Activate ETI in Certain Tomato Genotypes

6.5 AvrPtoB Virulence Activity

6.6 An Evolutionary Model of the Tomato–Pseudomonas Interaction

6.7 Summary

6.8 Acknowledgments

References

7: Rust Effectors

7.1 General Introduction to Rusts

7.2 Identification of Effectors in Bean Rust and Flax Rust as Haustorial Secreted Proteins

7.3 Genome-Wide Effector Prediction in the Poplar Rust and Wheat Stem Rust Genomes

7.4 Comparative Genomics of Effectors

7.5 Function of Rust Effectors

7.6 Conclusions

References

8: Dothideomycete Effectors Facilitating Biotrophic and Necrotrophic Lifestyles

8.1 Introduction to Dothideomycetes

8.2 Pregenome Identification of Avirulence and Effector Genes in Dothideomycetes

8.3 Pregenomic Identification of Host-Selective Proteinaceous Toxins of Dothideomycetes

8.4 Whole-Genome Searches for Effectors

8.5 Translocation of Fungal Effectors

8.6 Effector Diversification and Avoidance of R Protein-Mediated Resistance

8.7 Concluding Remarks

8.8 Acknowledgements

References

Section 4: Effector Trafficking: Processing/Uptake by Plants and Secretion/Delivery by Microbes

9: Effector Translocation and Delivery by the Rice Blast Fungus Magnaporthe oryzae

9.1 Introduction

9.2 The Fungus Magnaporthe oryzae

9.3 Hyphal Tip Secretion in Filamentous Fungi

9.4 Identification of Magnaporthe oryzae Effectors

9.5 To BIC or Not to BIC—That Is the Question

9.6 Effector Translocation into Host Rice Cells by M. oryzae

9.7 Concluding Remarks

References

10: Entry of Oomycete and Fungal Effectors into Host Cells

10.1 Effector Entry into Host Cells

10.2 Mechanisms of Entry

10.3 Summary, Perspective, and Conclusions

10.4 Acknowledgments

Note

References

Section 5: Emerging Effectors–Symbionts, Nematodes, Insects, Metabolites

11: Roles of Effector Proteins in the Legume–Rhizobia Symbiosis

11.1 Introduction to the Legume–Rhizobia Symbiosis

11.2 Nodule Formation

11.3 Rhizobial Molecular Signals Required for Nodule Formation

11.4 Regulation of Rhizobial T3SS and T4SS

11.5 Effects of T3SS and T4SS on Nodulation

11.6 Secretion System Substrates—Rhizobial Effectors

11.7 Conclusions

References

12: Mutualistic Effectors: Architects of Symbiosis

12.1 The Concept of Mutualism

12.2 Restructuring of Plant Signaling Pathways

12.3 Restructuring of Plant and Fungal Cell Wall Properties

12.4 Fungal Effectors Divert and Reprogram Plant Defenses

12.5 Fungal Effectors Restructure Nutrient Fluxes

12.6 Concluding Thoughts and Future Directions

References

13: Nematode Effector Proteins: Targets and Functions in Plant Parasitism

13.1 Introduction

13.2 Cell Wall-Modifying Effectors

13.3 Nematode Effectors Manipulating Plant Cell Biology

13.4 Nematode Effectors Manipulating Plant Defenses

13.5 Perspectives

References

14: Effectors in Plant–Insect Interactions

14.1 Introduction

14.2 Herbivore Behavior and Feeding Styles

14.3 Similarities between Plant–Microbe and Plant–Insect Interactions

14.4 Evidence for Effectors That Elicit Plant Defenses in Plant–Insect Interactions

14.5 Evidence for Effectors That Promote Plant–Insect Interactions

14.6 Functional Genomics Approaches for Identification of Insect Effectors

14.7 Future Perspectives

Acknowledgments

References

15: Fungal Secondary Metabolites: Ancient Toxins and Novel Effectors in Plant–Microbe Interactions

15.1 Introduction to Fungal Secondary Metabolism

15.2 Fungal SMs as Effectors

15.3 Fungal Secondary Metabolism: A Genomic Perspective

15.4 Fungal SMs as Effectors Involved in Plant Infection

15.5 Next Challenges about Effector SMs

References

Index

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Effectors in plant-microbe interactions / edited by Francis Martin, Sophien Kamoun. p. cm. Includes bibliographical references and index. ISBN 978-0-4709-5822-3 (hard cover : alk. paper) 1. Plant-microbe relationships – Molecular aspects. I. Martin, Francis, 1954–II. Kamoun, Sophien. QR351.E34 2012 579′.178–dc23 2011028322

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Contributors

Pierre Abad INRA UMR 1301, CNRS UMR 6243 UNSA 400 route des Chappes F-06903 Sophia-Antipolis France

Silvia Ardissone Laboratoire de Biologie Moléculaire des Plantes Supérieures Université de Genève 30 Quai Ernest-Ansermet Sciences III 1211 Genève 4 Switzerland

Thomas J. Baum Department of Plant Pathology and Microbiology Iowa State University Ames, IA 50011 USA

Thomas Boller Botanisches Institut Universität Basel Hebelstrasse 1 4056 Basel Switzerland

Jorunn I.B. Bos Cell and Molecular Sciences James Hutton Institute Invergowrie Dundee, DD2 5DA UK

Liliana M. Cano The Sainsbury Laboratory Norwich, NR4 7UH UK

Delphine Chinchilla Botanisches Institut Universität Basel Hebelstrasse 1 4056 Basel Switzerland

Jérôme Collemare Wageningen University Laboratory of Phytopathology Droevendaalsesteeg 1 6708 PB Wageningen The Netherlands

Mireille van Damme The Sainsbury Laboratory Norwich, NR4 7UH UK Present address: Wageningen University Laboratory of Phytopathology Droevendaalsesteeg 1 6708 PB Wageningen The Netherlands

Eric L. Davis Department of Plant Pathology North Carolina State University Raleigh, NC 27607 USA

William James Deakin Laboratoire de Biologie Moléculaire des Plantes Supérieures Université de Genève 30 Quai Ernest-Ansermet Sciences III 1211 Genève 4 Switzerland

Peter N. Dodds CSIRO Plant Industry GPO Box 1600 Canberra, ACT 2601 Australia

Sébastien Duplessis UMR INRA-UHP 1136 Interactions Arbres/Micro-organismes Centre INRA de Nancy 54280 Champenoux France

Gunther Doehlemann Max Planck Institute for Terrestrial Microbiology Department of Organismic Interactions Karl-von-Frisch Strasse 10 D-35043 Marburg Germany

Dagmar Hann Botanisches Institut Universität Basel Hebelstrasse 1 4056 Basel Switzerland

Saskia A. Hogenhout Department of Disease and Stress Biology John Innes Centre Norwich Research Park Norwich, NR4 7UH UK

Richard S. Hussey Department of Plant Pathology University of Georgia Athens, GA 30602 USA

David L. Joly Agriculture and Agri-Food Canada Pacific Agri-Food Research Centre Summerland, BC V0H 1Z0 Canada

Regine Kahmann Max Planck Institute for Terrestrial Microbiology Dept. Organismic Interactions Karl-von-Frisch-Strasse 10 D-35043 Marburg Germany

Sophien Kamoun The Sainsbury Laboratory Norwich, NR4 7UH UK

Ralf Koebnik Institut de recherche pour le développement UMR ‘Résistance des Plantes aux Bioagresseurs’ 911 Avenue Agropolis 34394 Montpellier France

Thomas Kroj UMR Biologie et Génétique des Interactions Plante-Parasite Campus International de Baillarguet F-34398 Montpellier France

Marc-Henri Lebrun UR 1290 INRA BIOGER Campus AgroParisTech Thiverval-Grignon France and UMR 5140 CNRS UCB BCS Microbiologie Adaptation Pathogénie Bayer Cropscience Lyon, France

Magdalen Lindeberg Department of Plant Pathology and Plant-Microbe Biology Plant Science Building Cornell University Ithaca, NY 14853 USA

Francis Martin UMR INRA-UHP 1136 Interactions Arbres/Micro-organismes Centre INRA de Nancy 54280 Champenoux France

Gregory Martin Boyce Thompson Institute for Plant Research and Department of Plant Pathology and Plant-Microbe Biology Cornell University Ithaca, NY 14853 USA

Thomas Mentlak School of Biosciences University of Exeter Geoffrey Pope Building Exeter, EX4 4QG UK

Melissa G. Mitchum Division of Plant Sciences and Bond Life Sciences Center University of Missouri Columbia, MO 65211 USA

Ricardo Oliva The Sainsbury Laboratory Norwich, NR4 7UH UK

Jonathan M. Plett UMR INRA-UHP 1136 Interactions Arbres/Micro-organismes Centre INRA de Nancy 54280 Champenoux France

Sylvain Raffaele The Sainsbury Laboratory Norwich, NR4 7UH UK

Marie-Noëlle Rosso INRA UMR 1301 CNRS UMR 6243 UNSA 400 route des Chappes F-06903 Sophia-Antipolis France

Thierry Rouxel INRA-Bioger Campus AgroParisTech BP 01 78850 Thiverval-Grignon France

Kerstin Schipper Max Planck Institute for Terrestrial Microbiology Department of Organismic Interactions Karl-von-Frisch-Strasse 10 D-35043 Marburg Germany

Sebastian Schornack The Sainsbury Laboratory Norwich, NR4 7UH UK

María Eugenia Segretin The Sainsbury Laboratory Norwich, NR4 7UH UK Present address: Laboratorio de Biotecnología Vegetal INGEBI-CONICET Vta. Obligado 2490 2do. piso (C1428ADN) Ciudad de Buenos Aires Argentina

Geert Smant Laboratory of Nematology Wageningen University Binnenhaven 5 6709PD Wageningen The Netherlands

Nicholas J. Talbot School of Biosciences University of Exeter Geoffrey Pope Building Exeter, EX4 4QG UK

Brett M. Tyler Virginia Bioinformatics Institute and Department of Plant Pathology Physiology and Weed Science Virginia Polytechnic Institute and State University Washington Street Blacksburg, VA 24061 USA Present address: Center for Genome Research and Biocomputing and Department of Botany and Plant Pathology 3021 Agriculture and Life Sciences Building Oregon State University Corvallis, Oregon, 97331-7303 USA

Pierre J.G.M. de Wit Wageningen University Laboratory of Phytopathology P.O. Box 6798PB Wageningen, The Netherlands

Foreword

Effectors in Plant–Microbe Interactions: Past to Present

Brian Staskawicz

Department of Plant and Microbial Biology, University of California Berkeley, Berkeley, CA 94720, USA

The basic understanding of why a phytopathogen can cause disease on only a few species of any particular plant has long intrigued plant pathologists. In fact, if one looks at all the potential disease-causing agents of plants, the ability of a pathogen to cause disease is often the exception as most plants are able to recognize and actively defend themselves against most pathogens in nature. Early work by E.C. Stakman at the University of Minnesota in early twentieth century established the concept of the “physiological race” of a single species of rust (Stakman, 1914). He demonstrated that physiological races derived from the sexual cycle of Puccinia graminis gave rise to distinct strains that varied in their ability to cause disease when inoculated on various wheat varieties. This observation was critical to the concept that resistance to cereal rust pathogens was race specific and that knowledge of the genetic variation in rusts was essential to the successful breeding for disease resistance. It was then Harold Flor in the 1940s with his work on flax rust who provided a genetic explanation for Stakman's “physiological race” concept (Flor, 1942). His work established that single gene differences in both the host and pathogen controlled whether a flax rust strain caused disease on a particular cultivar of flax. Building on these prior observations and work by Al Ellingboe along with the discovery of recombinant DNA and gene cloning, I set out with Douglas Dahlbeck and Noel Keen in the early 1980s to clone a gene that defined the “physiological race” that Stakman and Flor had previously described and genetically characterized. The cloning of an “avirulence” gene from a Pseudomonas syringae pv. glycinea race established that a single gene in the pathogen controlled whether this bacterium caused disease on a particular cultivar of soybean (Staskawicz et al., 1984). In this case, the avirulence gene was recognized as a single resistance gene in soybean. However, it was not until several years later that it was established that these so-called avirulence genes also played a major role in the virulence of the pathogen. This was accomplished once methods had been established for performing site-directed gene mutations in phytopathogenic bacteria such that isogenic strains could be constructed and evaluated on hosts that did not contain the cognate resistance gene. Mutations in the avrBs2 gene resulted in lower bacterial growth populations on pepper plants that did not contain the cognate Bs2 gene (Kearney and Staskawicz, 1990). Once it was established that avirulence genes could be isolated in this manner, it was not long before several more examples were published. The concept that avirulence genes also had a role in virulence was further strengthened by the discovery that the “Hrp” gene in Xanthomonas, Ralstonia, and Pseudomonas turned out to be highly homologous to the type three secretion systems genes that had been earlier established in animal bacterial pathogens (Fenselau et al., 1992; Gough et al., 1992). Since the medical field used the term “effector protein” to describe proteins that were delivered via the bacterial type three secretion systems, phytopathologists also adopted this term to be consistent with the medical field. Since the original discovery of phytopathogenic effectors, it has become apparent that all classes of plant pathogens employ effectors to either modulate or suppress plant innate immune functions (Dodds and Rathjen, 2010). Since the field has rapidly expanded over the last 5 years, the publishing of this book is timely as it brings together a wealth of information and points of view on a wide range of pathogen effectors. There is no question that we have learned a great deal about the mode of action of pathogen effectors to date, but this field is in its infancy and surely will flourish in the years to come. The combination of molecular, cellular, genomic, and structural studies will be paramount to this effort. As for the future, the sequencing of field isolates of naturally occurring pathogens will shed new light on pathogen diversity and will provide novel insights into the evolution and function of pathogen effectors in agricultural systems. This, in turn, will greatly benefit the deployment of durable disease-resistance strategies to control disease in an environmentally sustainable manner. One can only hope that translational approaches will be employed to solve important disease problems that are currently present and for new diseases that will emerge in the future.

References

1. Stakman, E. (1914) A Study in cereal rusts, physiological races. University of Minnesota Agricultural Experiment Station Technical Bulletin 138, 1–56.

2. Flor, H.H. (1942) Inheritance of pathogenicity in Melampsora lini. Phytopathology 32, 653–669.

3. Staskawicz, B., Dahlbeck, D., & Keen, N. (1984) Cloned avirulence gene of Pseudomonas syringae pv. glycinea determines race-specific incompatibility on Glycine max (L.) Merr (Translated from ENG). Proceedings of the National Academy of Sciences USA 81(19), 6024–6028.

4. Kearney, B. & Staskawicz, B.J. (1990) Widespread distribution and fitness contribution of Xanthomonas campestris avirulence gene avrBs2. Nature 346, 385–386.

5. Fenselau, S., Balbo, I., & Bonas, U. (1992) Determinants of pathogenicity in Xanthomonas campestris pv. vesicatoria are related to proteins involved in the secretion in bacterial pathogens of animals. Molecular Plant-Microbe Interactions 5, 390–396.

6. Gough, C.L., Genin, S., Zischek, C., & Boucher, C.A. (1992) hrp genes of Pseudomonas solanacearum are homologous to pathogenicity determinants of animal pathogenic bacteria and are conserved among plant pathogenic bacteria. Molecular Plant-Microbe Interactions 5(5), 384–389.

7. Dodds, P.N. & Rathjen, J.P. (2010) Plant immunity: towards an integrated view of plant-pathogen interactions. Nature Reviews Genetics 11(8), 539–548.

Preface

Every single plant in nature is closely associated with mutualistic microbes, particularly fungi and bacteria. In addition, plants are repeatedly attacked by a multitude of pathogens and pests, including bacteria, fungi, oomycetes, nematodes, and insects. Deciphering how plants interact with both mutualistic and parasitic microbes is central to understanding their biology. One could almost argue that plant biology should be viewed as a subdiscipline of plant–microbe interactions. Identifying the plant–microbe cross talks is also crucial for a better understanding of the processes regulating the complex interactions between entangled plant and microbial communities in ecosystems.

The field of plant–microbe interactions has significantly matured in recent years. All major classes of molecular players both from plants (surface and intracellular immune receptors) and microbes (microbial pattern molecules and effectors) have now been revealed. This book focuses on effectors, secreted microbial molecules that alter plant processes and facilitate colonization. Effectors are central to our newly integrated view of plant–microbe interactions. Effectors have evolved to facilitate parasitism, for example, by suppressing host immunity in a variety of ways. However, they can also “trip on the wire” and activate plant immune receptors, a response known as effector-triggered immunity. These are complex interactions and the coevolutionary dynamics between plants and microbes have left striking marks in their genomes. Our goal was to take stock of current knowledge on effectors of plant-associated organisms and illustrate the diverse and complex ways in which effectors interact with their host plants.

The book opens with general reviews on plant immunity and how it is targeted by microbial effectors (Chapters 1 and 2). The field of effector biology has greatly benefited from genome-wide analyses, which result in complete catalogs of effector genes. Chapters 3–5 report on genome-wide analyses and evolution of effectors genes. These chapters nicely illustrate how comparative genomics greatly contributed to our understanding of effector evolution. Chapters 6–8 describe how effectors function in suppressing host immunity and how they are perceived by plant immune receptors. How effectors traffic inside plant cells is covered by Chapters 9 and 10. Finally, the closing Chapters 11–15 cover emerging topics. Effectors have been reported in a number of plant–microbe systems, including bacterial and fungal symbioses, as well as nematode and insect pests.

Effector biology is a new and fast-paced field of research. As with all emerging fields of science, consensus among researchers has not always been reached and some topics remain controversial. Readers will surely notice more than one example throughout the book. We elected to keep such “inconsistencies” rather than enforce an arbitrarily sanitized version. We hope that such differences between authors will be informative of the current dynamic state of our science.

Books may have become less fashionable in the age of tweeting and microblogging. However, we hope that there is value in a document that summarizes the current state of the field of effector biology and provides a handy complement to the literature for both novice and experienced scientists.

Francis Martin and Sophien Kamoun

Section 1

Plant Immune Response Pathways

1

Innate Immunity: Pattern Recognition in Plants

Delphine Chinchilla and Thomas Boller

1.1 Pattern Recognition through MAMPs (Microbe-Associated Molecular Patterns)

Classic work attempted to define and characterize the so-called “elicitors,” pathogen-derived molecules that would elicit a defense response in plants (Darvill and Albersheim, 1984; Boller, 1995). In the case of oomycetes and fungi, these “elicitors” turned out to be characteristic microbial structures derived from their cell walls, such as the heptaglucan epitope of Phytophthora megasperma (see Darvill and Albersheim, 1984) and chitin fragments (Felix et al., 1993), or from microbial membranes, such as arachidonic acid (Preisig and Kuc, 1985) and ergosterol (Granado et al., 1995). Thus, plants appeared to perceive microbes through common patterns that were not specifically associated with pathogens (Boller, 1995). However, although these elicitors were able to induce a vigorous defense response, their importance for actual plant–pathogen interactions remained elusive.

The appreciation of these “general elicitors” changed when a similar principle was described in the field of (human) immunology: In the evolutionarily ancient “innate immunity,” a group of receptors named “pattern recognition receptors (PRRs)” was found to recognize conserved molecular patterns of microbes that are essential for their survival, the so-called “pathogen-associated molecular patterns (PAMPs)” (Medzhitov and Janeway, 2000, Janeway and Medzhitov, 2002). Interestingly, both plants (Gómez-Gómez and Boller, 2000) and animals (Hayashi et al., 2001) were found to possess specific PRRs for bacterial flagellin, namely flagellin sensing 2 (FLS2) and Toll-like receptor 5 (TLR5). This highlighted the similarities of plant and animal innate immunity (Asai et al., 2002), particularly because it appeared that the two PRRs had arisen by convergent evolution rather than from a primeval eukaryotic PRR (see Ausubel, 2005; Boller and Felix, 2009). As well as illustrated by the PRRs for flagellin, it is apparent that the molecular patterns recognized are characteristic of whole classes of microbes, independent of whether they are pathogenic or not, and therefore should more precisely be called “microbe-associated molecular patterns (MAMPs),” a term we will use throughout this chapter (see also Mackey and McFall, 2006; Boller and Felix, 2009; Boller and He, 2009). In fact, well-adapted pathogens might alter and camouflage the molecular patterns that lead to recognition by the PRRs, as illustrated by the changes in the flagellin genes of some plant pathogenic bacteria, such that true pathogens may no longer present the MAMP in question (Felix et al., 1999; Pfund et al., 2004; Sun et al., 2006).

The variety of MAMPs is large, as summarized in Table 1.1. Typically, for a given class of microbes, a given plant species can perceive several different MAMPs. This redundancy guarantees a robust recognition of the microbe.

Table 1.1 Overview of PRRs/binding sites for MAMPs/DAMPs characterized in plants.

While MAMPs are generally characteristic of a whole class of microorganisms, many of these MAMPs are not perceived in a general way by most plants, but only by a few of them, e.g., by most members of an order or family (reviewed in Boller and Felix, 2009). For example, recognition of the active epitope of elongation factor-Tu (EF-Tu), called elf18, is restricted to the Brassicaceae family (Kunze et al., 2004, Zipfel et al., 2006) and bacterial cold shock protein (CSP) is active only in Solanaceae (Felix and Boller, 2003). From an evolutionary point of view, it is probable that perception of EF-Tu and CSP are more recent systems than the perception of flagellin, which is common to numerous plant species (Albert et al., 2010a). There may be an advantage for a given host plant, in terms of coevolution, to recognize a MAMP that most other plants do not.

1.2 Some Classical MAMP-Receptor Pairs

1.2.1 MAMP Receptors: PRRs

In animals, PRRs can be separated into surface-located receptors, called Toll-like receptors (TLRs) and intracellular receptors of the NOD-like family (Takeuchi and Akira, 2010). In plants, the PRRs identified so far are all located at the plasma membrane (Zipfel, 2008). There is currently no example of intracellular recognition of a MAMP in terms of the above definition, although plants possess specialized intracellular receptors of the NOD-like family to perceive effectors (see Chapter 2).

Most plant PRRs described so far belong to the class of receptor-like kinase (RLKs) (Shiu and Bleecker 2001; Shiu et al., 2004): these proteins contain an ectodomain probably acting as the binding site for the respective ligand, followed by a single pass transmembrane domain and a cytoplasmic protein kinase domain, which is likely to function in intracellular signal transduction. Many RLKs are induced by biotic stresses, including MAMP treatments (Zipfel et al., 2004, 2006; Kemmerling et al., 2007), and some were shown to be dispensable for plant development and thus are good candidates as PRRs (Lehti-Shiu et al., 2009).

Three typical leucine-rich repeat (LRR)-RLKs acting as PRRs (which can be referred as LRR-receptor kinases) are FLS2 and EFR, the Arabidopsis receptors for flagellin (Gómez-Gómez and Boller, 2000) and EF-Tu (Zipfel et al., 2006), respectively, and Xa21, an RLK of rice that has long been considered an unusual “resistance gene” product (Song et al., 1995) but was recently shown to perceive ax21 (Lee et al., 2009), a peptide universally conserved in Xanthomonas oryzae that should be considered as a MAMP (reviewed by Ronald and Beutler, 2010; see also Table 1.1 for further examples).

Other functional PRRs are members of a second class of receptor proteins called receptor-like proteins (RLPs). These proteins have a similar structure to RLKs but lack kinase domains; instead, these proteins often exhibit a short cytoplasmic domain with no signaling signature. This suggests different molecular mechanisms of receptor activation than those controlling RKs; they probably have to interact with other membrane proteins to transmit the signal across the membrane (Wang et al., 2008a). RLPs have a similar structure to the animal TLRs, but in contrast to TLRs that are encoded by 10–12 different genes in mammals (Leulier and Lemaitre, 2008), RLPs expanded into a larger family, with 57 members in the Arabidopsis genome (Wang et al., 2008a). The apparent expansion of the families of RLKs and RLPs may indicate that in evolution these receptors have become one of the preferred systems for non self-perception in plants. To date, two RLPs have been identified as PRRs for individual MAMPs: (1) the receptor for the fungal MAMP xylanase (ethylene (ET)-inducing xylanase, EIX) in tomato, named LeEIX (Ron and Avni, 2004); (2) and the chitin-binding site CEBiP (chitin elicitor binding protein) in rice (Kaku et al., 2006).

PRRs are often identified on the basis of genetics, and only for a small number, biochemical evidence has been provided to demonstrate direct interaction between the receptor and its respective ligand (Table 1.1). However, technical advances in methods, such as affinity chromatography, chemical cross-linking, and immunoprecipitation, have allowed the unequivocal identification of PRR–MAMP interactions and will hopefully shape the future of PRR characterizations (Chinchilla et al., 2006; Kaku et al., 2006; Shinya et al., 2010).

1.2.2 Flagellin Perception in Arabidopsis through FLS2: A Paradigm for MAMP Recognition in Plants

Flagellin, the best-characterized MAMP in plants, was serendipitously identified as the active elicitor in a “harpin” preparation from the plant pathogen Pseudomonas syringae (Felix et al., 1999). Flagellin is the main building block of the flagellum, which allows bacteria to “swim”; typically, a flagellum consists of 10,000 monomers of flagellin. Due to its essential role in bacterial motility, and also due to its abundance and surface exposure, flagellin represents a perfect “molecular pattern” to detect the approach of potentially pathogenic bacteria. The “epitope” of flagellin perceived by plants is located at the N-terminus of the protein and is strongly conserved in bacteria (Felix et al., 1999). Its effect can be mimicked by a synthetic peptide called flg22. Most higher plants, including both gymnosperms and angiosperms, share the capacity to perceive flg22 as a MAMP (Albert et al., 2010a).

Among responses typical for MAMPs (see also next Section 1.3), a prolonged treatment with flg22 induces a growth arrest in seedlings. Using this growth defect as a readout, a genetic screen was conducted in Arabidopsis that allowed the identification of FLS2 as a gene essential for flagellin responses (Gómez-Gómez and Boller, 2000). Using a biochemical approach with radiolabeled flg22, it was shown that these mutants were not only impaired in flg22 signaling but also in flg22 binding (Gómez-Gómez et al., 2001). Both Arabidopsis and tomato perceive flg22, but interestingly these two species show different specificities of binding and responses to derivatives of flg22 (Meindl et al., 2000; Bauer et al., 2001). Transfer of Arabidopsis FLS2 into tomato cells (having an endogenous flagellin receptor) induced new recognition specificities representative of the flagellin receptor from Arabidopsis indicating that AtFLS2 is the “bona fide” receptor for bacterial flagellin, controlling binding of ligand and activation of responses (Chinchilla et al., 2006). This was corroborated by cross-linking experiments with labeled flg22, followed by immunoprecipitation with antibodies specific for FLS2, which showed unequivocally that FLS2 interacts directly with its flg22 ligand (Chinchilla et al., 2006). Several orthologs of FLS2 were identified in tomato (LeFLS2; Robatzek et al., 2007), Nicotiana benthamiana (NbFLS2; Hann and Rathjen, 2007), and rice (OsFLS2; Takai et al., 2008). In rice, responses to flg22 are weak, but OsFLS2 is able to functionally complement Arabidopsis fls2 mutants, confirming that it is indeed a flagellin receptor (Takai et al., 2008). Comparison of orthologs of FLS2 indicated a highly conserved structure for this protein, including a large ectodomain of 28 LRRs, except for OsFLS2 lacking LRR3, and other conserved domains at the N- and C-terminus, indicating in particular that the large LRR domain is of functional relevance (Boller and Felix 2009).

It remains an open question where the exact binding site of flg22 lies within the LRR domain of FLS2. A random mutagenesis approach using the defined individual LRR domains of AtFLS2 indicated that LRR9 to LRR15 play an important role for FLS2 function (Dunning et al., 2007). Protein crystallography would seem a method of choice to delineate the ligand–receptor interaction; however, up to now, attempts to functionally express the extracellular flg22-binding site of FLS2 in heterologous systems were unsuccessful, perhaps because of its high degree of glycosylation in vivo (see Chinchilla et al., 2006).

The analysis of chimeric receptors is another approach to understand how the large ectodomains of the PRRs function in ligand binding and receptor activation. This is exemplified by a study where the ectodomain of the brassinolide receptor BRI1, a typical LRR-RK (Li and Chory, 1997) was fused to the cytoplasmic domain of the rice PRR Xa21 (Song et al., 1995). Rice cells expressing this chimeric receptor were able to induce defense responses after application of exogenous brassinolides (He et al., 2000). More recently, a role of the wall-associated kinase 1 (WAK1) in the perception of oligogalacturonides (damage-associated molecular patterns (DAMPs) released from the plant cell wall, which activate the plant-immune response) was demonstrated using chimeric receptors between the ectodomain of WAK1 and the protein kinase domain of EF-Tu receptor (EFR) (Brutus et al., 2010). In this work, as a proof of concept, functional chimeric receptors inducing an immune response were also constructed between the ectodomain of FLS2 and the protein kinase domain of EFR (Brutus et al., 2010). A refined analysis employing chimeras within the extracellular LRR domains of EFR and FLS2 allowed mapping of subdomains relevant for ligand binding and receptor activation in EFR (Albert et al., 2010b). Work is in progress to map such domains in FLS2, using the orthologs of the flagellin receptor from Arabidopsis and tomato, which show distinct specificities for ligand binding and responses (Robatzek et al., 2007).

1.2.3 EFR: An Evolutionarily Young but Efficient PRR Perceiving Bacterial EF-Tu

A protein able to induce defense responses was isolated from an extract of an Escherichia coli strain mutated for the flagellin synthesis gene FliC; it turned out to be the bacterial EF-Tu (Kunze et al., 2004). This new MAMP plays a crucial role in protein synthesis and belongs to the most abundant and most highly conserved bacterial proteins. Peptides representing the N-terminus of bacterial EF-Tu, namely elf18 and elf26, require N-terminal acetylation for full activity—a typical modification of bacterial but not of eukaryotic EF-Tu. Interestingly, elf18 and elf26 are recognized as elicitors only by plants from the Brassicaceae family. No activity could be measured in any other plant families tested to date (Kunze et al., 2004; Zipfel et al., 2006; Albert et al., 2010a). This indicates that evolution has shaped the recognition specificity of EFR—structurally a close relative of Xa21 (Boller and Felix, 2009)—only after the emergence of Brassicaceae, about 40 million years ago.

Interestingly, elf18 induces very similar responses as flg22, particularly with regard to altered gene expression (Zipfel et al., 2006). The observation that treatment with flg22 or elf18 induced transcript accumulation of FLS2 led to the hypothesis that RLKs induced by MAMPs are potential PRRs. Along this hypothesis, and focusing on the class of LRR-RLKs, Zipfel and colleagues established a collection of knock out mutants of Arabidopsis affected in the LRR-RLKs induced by flg22. This collection was screened for responsiveness to elf18, which allowed the identification of the EF-Tu receptor, called “EFR” (Zipfel et al., 2006). The EFR protein has a similar structure as FLS2: both belong to the LRR-RLK subfamily XII and the ectodomain of EFR consists of 21 LRRs. Transfer of the EFR gene to heterologous plants “blind” for elf18, such as N. benthamiana or tomato, resulted in responsiveness to this elicitor, demonstrating EFR is the receptor for EF-Tu, and also in an enhanced resistance to several bacterial pathogens, demonstrating the biological relevance of MAMP perception in disease resistance (Zipfel et al., 2006; Lacombe et al., 2010).

Recent genetic screens aiming at identifying new regulators of elf18 signaling allowed the identification of an element of the secretory pathway important for maturation of the EFR receptor, namely the ER quality control system (ER-QC; Nekrasov et al., 2009; Li et al., 2009b; Saijo et al., 2009; Lu et al., 2009; Haweker et al., 2010). Intriguingly, neither FLS2 nor the RLK CERK1 (chitin elicitor receptor kinase1) involved in chitin signaling (see also Section 1.2.6) seem to be affected by these mutations.

1.2.4 A newly Recognized MAMP–PRR Pair: The Rice LRR-RK Xa21 Recognizes ax21 from Xoo

The receptor kinase Xa21 is among the first receptor kinases cloned in plants (Song et al., 1995). It was shown to provide rice cultivars with considerable resistance to X. oryzae pv oryzae (Xoo), a pathogen causing bacterial blight. The Xa21 gene was for a long time considered to be a resistance gene involved in race specific resistance but now it is considered a PRR (Park et al., 2010; Ronald and Beutler, 2010).

This is because the ligand of this receptor was recently identified as the sulfated peptide axYS22, more commonly called “ax21”, for activating Xa21 immunity (Lee et al., 2009). This microbial molecule appears to be conserved in all Xanthomonas species and may play a role in quorum sensing (Lee et al., 2006); thus, it resembles a MAMP rather than an “avirulence” determinant.

Some Xanthomonas strains evade ax21 recognition by avoiding sulfation of the secreted peptide, a modification that appears to be crucial for recognition by Xa21 (da Silva et al., 2004). In general, posttranslational modifications of MAMPs, such as acetylation and sulfation, seem to emerge as an important way to modulate MAMP recognition in plants (Kunze et al., 2004; Lee et al., 2009).

Numerous genetic, molecular, and biochemical studies have been conducted on the Xa21 immunity (reviews: Park et al., 2010; Ronald and Beutler, 2010). Among them is the finding that a kinase dead version of Xa21 is still partially active (Liu et al., 2002). Instead, activation of the receptor complex via phosphorylation may be controlled by an unknown kinase (Park et al., 2010). In addition, several interactors of Xa21 were identified by diverse approaches (Park et al., 2010). In particular, an ATPase called XB24 (for Xa21 binding 24) was shown to interact with Xa21 to promote its autophosphorylation (Chen et al., 2010). In plants, silencing of XB24 enhances Xa21 immunity; thus, activation of Xa21 following ax21 perception may be the result of the inactivation or the release of XB24 from the receptor complex. As discussed in Section 1.2.3, Xa21 accumulation seems to be regulated by ER-QC and ER-associated degradation systems, as indicated by the identification of several regulators of these pathways in the Xa21 receptor complex (Park et al., 2010).

1.2.5 BAK1—A Positive Regulator of PRRs

Our understanding of PRR activation and signal transduction made an important step forward with the identification of a second RLK involved in flagellin signaling, called BAK1 (Chinchilla et al., 2007; Heese et al., 2007).

The BRI1-associated kinase 1 protein was first identified as a coreceptor for the brassinosteroid receptor BRI1 in Arabidopsis (Li et al., 2002; Nam and Li, 2002). Surprisingly, BAK1 seems to be shared by several signaling pathways controlling developmental as well as defense responses (reviewed in Chinchilla et al., 2009; see also Table 1.1). Consistent with a role in plant immunity, plants depleted for BAK1 show reduced responses to flg22 and exhibited more symptoms to virulent bacteria (Chinchilla et al., 2007; Heese et al., 2007; Kemmerling et al., 2007).

BAK1 is localized at the plasma membrane of plant cells (Li et al., 2002; Nam and Li, 2002) and further biochemical analysis demonstrated that BAK1 interacts with FLS2 in a flg22-dependent manner (Chinchilla et al., 2007; Heese et al., 2007): this oligomerization process is extremely quick, occurring within seconds of elicitation with flg22 (Schulze et al., 2010). However, BAK1 seems to be dispensable for flg22 binding (Chinchilla et al., 2007). Using an in vivo phospholabeling approach, phosphorylation events were detected in both BAK1 and FLS2 very rapidly after flg22 perception (∼15s; Schulze et al., 2010). These data indicate that the flagellin receptor is activated in a similar way as animal tyrosine receptor kinases (Lemmon and Schlessinger, 2010) or the RK BRI1 in Arabidopsis (Wang et al., 2008b). In the latter model, the ligand-binding RK BRI1 is activated after BL perception, which promotes its association with BAK1. In this heterocomplex, BRI1 and BAK1 phosphorylate each other in “trans” to amplify the BL signal. In absence of BAK1, BRI1 can still exhibit some kinase activity and BL signaling is functional but at a lower level (Wang et al., 2008b).

Recently, the kinase activity of BAK1 was shown to be essential for flg22 signaling, although not for its heteromerization with FLS2 (Schulze et al., 2010). Consistently, treatment of Arabidopsis cells with a general kinase inhibitor did not affect the FLS2–BAK1 complex formation. This is in contrast to the BRI1–BAK1 complex, which requires BRI1 kinase activity for its formation or stability (Wang et al., 2008b). It remains to be tested if FLS2 kinase activity is required for signaling and if this activity is promoted by BAK1 in vivo.

Several studies support a role of BAK1 as a central regulator of PRRs (see Table 1.1). For example, activation of EFR also involves BAK1 (Chinchilla et al., 2007; Schulze et al., 2010). Furthermore, BAK1 seems to play a role in DAMP signaling since it is phosphorylated upon perception of Pep1 (Huffaker et al., 2006) in Arabidopsis and interacts with the Pep1 receptors, PEPR1 and PEPR2 (Yamaguchi et al., 2006, 2010), and two other LRR-RKs (Schulze et al., 2010; Postel et al., 2010). Since BAK1 appears to regulate several LRR-RKs, it will be interesting to test if OsBAK1, the homolog of BAK1 recently identified in rice (Li et al., 2009a), is involved in Xa21-mediated immunity. The role of BAK1 in PRR regulation is not restricted to LRR-RKs; a recent report indicated a role of BAK1 in regulation of xylanase perception controlled by the RLP LeEIX1 and LeEIX2 (Bar et al., 2010). Interestingly, chitin perception, which involves a different class of receptor kinase (LysM RKs), does not require BAK1 (Shan et al., 2008; Schulze et al., 2010). Finally, BAK1 also plays a role in resistance to fungi and oomycetes, because plants depleted for BAK1 were more susceptible to several of these pathogens, including Verticillium, Alternaria, and Hyaloperonospora parasitica (see Chinchilla et al., 2009).

BAK1 is a member of a small gene family of five members, called the SERKs, initially defined as “somatic embryogenesis-receptor kinases” (see Boller and Felix 2009; Chinchilla et al., 2009). Two of them, BAK1 (SERK3) and SERK4, share a particularly high homology on the level of amino acid sequences. Intriguingly, bak1(serk3) serk4 double mutants of Arabidopsis are lethal at the seedling stage, displaying constitutive defense-gene expression, callose deposition, reactive oxygen species (ROS) accumulation, and spontaneous cell death even under sterile growing conditions (He et al., 2007). Thus, BAK1 and its SERK4 homolog must be involved in cell death control as well (see Chinchilla et al., 2009).

1.2.6 Chitin Perception in Plants: A New Scenario for Molecular Events of MAMP Perception

Chitin is a polymer of N-acetylglucosamine found in fungal cell walls, insect exoskeletons, and crustacean shells, but not in plants. Plants do not have chitin but instead possess chitinases that degrade chitin. It was hypothesized that plant chitinases can degrade chitin in the cell wall of the invading fungus and release short fragments of chitin (chito-oligosaccharides) that can act as MAMPs (reviewed in Boller, 1995).

Similar to flg22, chitin oligosaccharides are active in a wide range of plants, including both dicots (Felix et al., 1993) and monocots (Shibuya and Minami, 2001). Much effort has been made since many years to identify and characterize high affinity binding sites for chitin oligomers in membrane fractions of diverse plant species such as rice (Shibuya et al., 1993) and tomato (Baureithel et al., 1994). But the molecular identity of the first chitin-binding protein was discovered only recently with the isolation of CEBiP from extracts of rice culture cells using chitin affinity chromatography (Kaku et al., 2006).

The CEBiP from rice possesses two “lysine motifs” (LysM) in its ectodomain. In legumes, LysM motifs were identified in the plant receptor kinases involved in the recognition of Nod factors, lipo-chitooligosaccharides secreted by symbiotic rhizobium bacteria to establish the nitrogen-fixing nodule symbiosis (Limpens et al., 2003; Madsen et al., 2003; Radutoiu et al., 2003). In Lotus japonicus, NFR1 and NFR5 are the candidate Nod Factor receptors. L. japonicus (Lj) and Lotus filicaulis (Lf) associate with different symbiotic strains of Rhizobium due to different Nod factor recognition specificities. By domain swapping of LysM motifs from LjNFR1/5 and LfNFR1/5, it was shown that the second LysM domain of NFR5 is involved in this recognition process; thus, this domain appears to be capable of binding carbohydrate molecules in a highly specific way (Radutoiu et al., 2007) and may be involved, remarkably, in recognition of both friends and foes in plants (Knogge and Scheel, 2006).

Since CEBiP does not contain a cytoplasmic signaling domain, in contrast to LysM-domain receptor kinases such as NFR1 and NFR5, it is likely that it cooperates with other proteins to transmit the signal from the plasma membrane to the cytoplasm. One LysM RK was characterized as the CERK1 in Arabidopsis and more recently in rice: plants affected in cerk1 expression are unable to respond to chitin, indicating that this LysM RK is involved in chitin perception and/or signaling (Miya et al., 2007; Wan et al., 2008; Shimizu et al., 2010). Interestingly, in rice, OsCERK1 can form heteromers with CEBiP in vivo, in a ligand-dependent manner (Shimizu et al., 2010). But does OsCERK1 contribute to ligand binding? The main band cross-linked to labeled chitin was found to be CEBiP, this cross-linking signal was not affected in knockdown lines of OsCERK1, indicating that CEBiP is the major molecule that binds chitin oligosaccharides on the rice cell surface (Miya et al., 2007; Shimizu et al., 2010). In contrast, recent studies in Arabidopsis report the capacity of AtCERK1 to bind chitin (Iizasa et al., 2010; Petutschnig et al., 2010). However, some other data speak against CERK1 as a binding site specific for chitin: notably AtCERK1 was shown to be targeted by a bacterial effector and thus might be involved in signaling of a bacterial MAMP as well (Gimenez-Ibanez et al., 2009).

Although the role of CERK1 in ligand binding remains elusive, it exhibits a clear kinase activity in vitro (Miya et al., 2007) and this kinase activity is essential for activation of early defense responses in Arabidopsis (Petutschnig et al., 2010). Moreover, similar to other RKs, such as FLS2 and BAK1, CERK1 is phosphorylated in vivo in its cytoplasmic domain, as shown by proteomic analysis of CERK1 in chitin-treated cells of Arabidopsis (Petutschnig et al., 2010).

Currently, it is difficult to draw a clear model for chitin perception based on the divergent findings in Arabidopsis and rice. CERK1 and CEBiP are clearly essential for chitin signaling, and it is tempting to imagine that these membrane proteins collaborate to form a functional receptor for chitin in rice. But in contrast to the BRI1–BAK1 and FLS2–BAK1 models where activation of receptors seems to occur via transphosphorylation events between the kinase domains of both RKs in the cytoplasm, it is more difficult to forecast the molecular mechanisms controlling activation of chitin receptor elements. Since CERK1 can form homodimers (Shimizu et al., 2010), it is possible that chitin perception activates CERK1 via transphosphorylation events within these homodimers. Alternatively, another unknown protein (kinase) present in the chitin receptor complex may activate CERK1 in response to ligand binding. More work needs to be done to clarify the mechanism of chitin signaling, a fundamental and very interesting example of MAMP perception.

1.3 Physiological Responses and Signaling Events Induced by Elicitors

Recognition of MAMPs through PRRs leads to a number of responses, in a well-ordered temporal pattern, and culminates in a state of “PTI” (originally defined as PAMP-triggered immunity, but now better redefined as “pattern-triggered immunity”). In the following, an overview is provided on the elements of PTI.

1.3.1 Immediate Early Responses

Ion Fluxes

Among the earliest and most easily recordable physiological responses to MAMPs is an alkalinization of the growth medium due to changes of ion fluxes across the plasma membrane in plant cell cultures (Boller, 1995; Boller and Felix, 2009). This response starts after a lag phase of ∼0.5–2.0 minutes and is certainly the easiest readout to identify new MAMPs from microbial extracts.

Rapid changes in ions include increased influx of H+ and Ca2+ and a concomitant efflux of K+; and an efflux of anions, in particular of nitrate (Wendehenne et al., 2002; Jeworutzki et al., 2010). These ion fluxes lead to membrane depolarization as recorded, for example, in electrophysiological studies with soybean cells challenged with the fungal MAMP, heptaglucan (Mithöfer et al., 2005) and more recently with mesophyll cells and root hairs from Arabidopsis treated with flg22 and elf18 (Jeworutzki et al., 2010).

In eukaryotes, the Ca2+ ion is a ubiquitous intracellular second messenger involved in numerous signaling pathways regulating developmental as well as defense processes. Variations in the cytosolic concentration of Ca2+ ([Ca2+]cyt) couple a large array of signals and responses in plants, although the way this response activates specific targets and responses remains unclear. Specificity of [Ca2+]cyt may be due to the time course of [Ca2+]cyt variations and the location of the [Ca2+]cyt increase (Garcia-Brugger et al., 2006).

Different Ca2+ signatures (varying with amplitude, frequency, time, and location) have been associated with diverse MAMPs (Lecourieux et al., 2005; Gust et al., 2007; Aslam et al., 2008; Aslam et al., 2009; Jeworutzki et al., 2010), and these signatures may potentially be decoded by distinct calcium sensors.

Calcium sensors perceive changes in [Ca2+]cyt that directly binds to the EF-hand motif of these proteins to modulate their activity. Best evidence for a role of calcium sensors in PTI is based on a recent study on calcium-dependent protein kinases (CDPKs) (Boudsocq et al., 2010), but other sensors such as calmodulins and calmodulin-binding proteins may also contribute to this regulation (Reddy and Reddy 2004). Using a functional genomic screen and genome-wide gene expression profiling, specific CDPKs, CPK4, CPK5, CPK6, and CPK11, were shown to control ROS production and expression of a subset of genes induced by MAMPs in Arabidopsis (Boudsocq et al., 2010). Moreover, multiple knockout mutants cpk5/cpk6 and cpk5/cpk6/cpk11 are affected in disease resistance to P. syringae. It is still unknown how these CDPKs are regulated, when they are activated, and how they regulate plant defense responses.

Production of Reactive Oxygen Species (ROS)

A characteristic defense response of plants is the rapid and robust production of ROS by host cells, a reaction also known as “oxidative burst” (review: Boller and Felix, 2009). This response occurs after a lag phase of ∼2 minutes in Arabidopsis plants and is transient. ROS are highly toxic intermediates comprising reduced forms of oxygen, such as the superoxide anion and hydrogen peroxide.

The sources of ROS can be diverse: In Arabidopsis, a membrane localized NADPH oxidase called AtRbohD (for respiratory burst oxidase homolog D) appears to be responsible for all flg22-induced ROS produced in the apoplast (Nühse et al., 2007; Mersmann et al., 2010). In other systems, peroxidases may have a role in apoplastic ROS generation (Torres, 2010).

Regulation of NADPH oxidase-dependent ROS production probably involves Ca2+ ions, which may bind to the two EF hands motifs present in the N-terminal region of the protein, thereby regulating these enzymes. In addition, NADPH oxidases and therefore ROS generation appear to be regulated by phosphorylation (Benschop et al., 2007; Nühse et al., 2007; Ogasawara et al., 2008). Two phosphosites were identified in AtRbohD as a result of flg22 perception in Arabidopsis cells that were shown to be essential for ROS production (Nühse et al., 2007). In potato, the production of ROS by the potato NADPH oxidase B appears to be regulated by phosphorylation through two CDPKs, namely StCDPK4 and StCDPK5 (Kobayashi et al., 2007). Consistently, multiple mutants affected in CDPK genes showed impairment in flg22-induced ROS production (Boudsocq et al., 2010).

The role of ROS in disease resistance is not yet well understood. ROS can contribute to defense either directly as an antibiotic agent, or indirectly by promoting oxidative cross-linking in the cell wall (Apel and Hirt, 2004). Furthermore, it may also be involved in the closure of stomata, a defense mechanism restricting bacterial entrance (Melotto et al., 2006; see also Section 1.3.5).

Activation of More Cytoplasmic Kinases: MAPK

Mitogen-activated protein kinase (MAPK) cascades are highly conserved modules in all eukaryotes where they transfer information from sensors to cellular responses. Activation of MAPK cascades involves a phosphorelay mechanism composed of MAPK kinase kinases (MAPKKK), MAPK kinases (MAPKK), and MAPKs.

In plants, MAPK cascades play an important role in signaling in response to biotic and abiotic stresses (Colcombet and Hirt, 2008; Pitzschke et al., 2009).

Two distinct MAPK cascades regulate PTI in Arabidopsis. A first MAPK module MEKK1-MKK4/MKK5-MPK3/MPK6 was originally proposed to be responsible for flg22 signal transduction (Nühse et al., 2000; Asai et al., 2002). However, more recent work demonstrated that although mekk1 mutants were compromised in activation of MPK4, MPK3, and MPK6, responses to flg22 were not affected (Ichimura et al., 2006; Suarez-Rodriguez et al., 2007). Thus, the MEKK protein activating the MKK4/5-MPK3/MPK6 cascade for positive regulation of defense responses remains unknown. MEKK1 forms a second cascade with MKK1/2-MPK4. This cascade is thought to negatively regulate immunity because loss-of-function mutations in these kinases lead to constitutive activation of defenses and a dwarf phenotype associated with mutants accumulating salicylic acid (SA) and other defense-related compounds (Ichimura et al., 2006; Suarez-Rodriguez et al., 2007; reviewed by Pitzschke et al., 2009). However, the mechanisms by which MPK4 regulates immunity are largely unknown.

Importantly, the activation mechanism for neither of the two MAPK modules is known to date. Is it possible that PRRs at the plasma membrane directly activate top-level member of the cascades (the MAPKKK, also called MEKK) directly (occurring ∼1–2 minutes after MAMP perception)? Or, is activation mediated by other yet unidentified kinases?

Activation of Membrane Kinases: BIK1

The membrane-anchored kinase BIK1 was recently identified as a new component involved in early steps of MAMP signaling (Lu et al., 2010; Zhang et al., 2010). Botrytis-induced Kinase1 was originally identified as an Arabidopsis gene that is transcriptionally regulated by pathogen or elicitor treatment (Veronese et al., 2006). More recent studies showed that BIK1 is phosphorylated upon flg22 treatment (as observed by mobility shift in Western blot analysis) (Lu et al., 2010; Zhang et al., 2010). This modification of BIK1 peaks at ∼5–10 minutes after treatment and can thus be distinguished from the phosphorylation events between FLS2 and BAK1 which were observed ∼15 seconds after elicitation (Schulze et al., 2010). Interestingly, BIK1 interacts with the BAK1-FLS2 complex in nonstimulated cells and seems to be released from this complex upon 10 minutes of flagellin treatment (Lu et al., 2010; Zhang et al., 2010). In vitro, BIK1 can phosphorylate BAK1 and FLS2 and may be a direct substrate for BAK1, suggesting a role of BIK1 in the regulation of the flagellin receptor and/or signaling.

Consistently, Arabidopsis bik1 mutants are impaired in some responses (ROS production and callose) induced by flg22 and elf18, but also chitin (in contrast to bak1) (Lu et al., 2010; Zhang et al., 2010). Since bik1 mutants showed high SA content, bik1 sid2 double mutants, which exhibit normal accumulation of SA, were generated to study the effect of BIK1 in respect to bacterial growth. As expected, these mutants had a defect in flg22-induced resistance to P. syringae (Zhang et al., 2010)

Overall, these studies report on BIK1 as an interesting and new signaling element of PTI although further studies are necessary to get a more comprehensive view on its role in MAMP signaling and immunity.

1.3.2 Hormone Changes in Response to MAMPs with a Focus on Ethylene Signaling

The three major plant hormones associated with MAMP perception and biotic stress are SA, jasmonic acid (JA), and ET, but other plant hormones may play a role in the defense response of plants as well (reviewed in Bari and Jones, 2009). Here, we briefly summarize recent advances in our understanding of the role of ET.

ET accumulation is a well-known response to MAMP treatment and is a consequence of the phosphorylation-dependent activation of 1-aminocyclopropane-1-carboxylate (ACC) synthases (reviewed in Boller and Felix, 2009). Interestingly, the MAMP-induced MAPKs MPK3 and MPK6 phosphorylate ACC synthases ACS2 and ACS6 (Liu and Zhang, 2004; Han et al., 2010) as well as EIN3, a transcription factor involved in the ET response, leading to its stabilization (Yoo et al., 2008). But how is ET involved in broad-spectrum resistance? Two recent reports have shown that ET is involved in the regulation of FLS2 gene expression, thereby rendering the plants more sensitive to flg22 perception (Mersmann et al., 2010; Boutrot et al., 2010). Consistently, mutants of the well-known ET regulator EIN2 or the ET receptor ETR1 were nearly insensitive to flg22 and FLS2 transcript levels were strongly reduced. Chromatin immunoprecipitation assays revealed that EIN3 is able to bind the FLS2 promoter (Boutrot et al., 2010). On the basis of this, a simple model was proposed in which ET signaling would control the FLS2 pathway by a positive feedback mechanism (Boutrot et al., 2010). According to this model, plants would maintain a constant pool of FLS2 levels even in the absence of MAMPs due to endogenous ET levels. Upon activation by flg22, FLS2 undergoes endocytosis, which leads to a reduction of the amount of FLS2 at the plasma membrane (Robatzek et al., 2006). However, at the same time, flg22 treatment will activate ACC synthases through phosphorylation by MAPKs, which leads to EIN3 accumulation in the nucleus and thereby induction of FLS2 expression (Chen et al., 2009). Thus, FLS2 level would be regulated by a positive feedback loop driven by ET. The expression of BAK1 is not affected (Mersmann et al., 2010); it remains an open question whether other PRRs are regulated by ET.

1.3.3 Responses at the Level of Gene Expression

Modification of Gene Expression in Response to MAMP Treatments

In Arabidopsis, about a thousand genes are upregulated ∼30–60 minutes after MAMP treatment, as revealed by several transcriptome analyses (Zipfel et al., 2004, 2006; Gust et al., 2007; Wan et al., 2008). Among them are many genes involved in perception and signaling of MAMPs, including PRRs themselves (Navarro et al., 2004), and thus increase the “awareness” for potential pathogens. Other MAMP-induced genes encode enzymes involved in protein degradation, cell wall modification, secondary metabolite biosynthesis, and vesicle trafficking, which may help to arrest, directly or indirectly, the invading microbes (Navarro et al., 2004).

There is a clear overlap between the genes upregulated after flg22, elf18, and chitin treatments, albeit small differences exist (Zipfel et al., 2006; Wan et al., 2008). These data suggest that the signaling pathways triggered by different MAMPs converge. By contrast, only a small number of downregulated genes are common to chitin and flg22/elf18 responses (Wan et al., 2008).

These considerable transcriptional changes in defense genes appear to be mediated by two types of kinases already described above, MAPKs and CDPKs (Asai et al., 2002; Boudsocq et al., 2010). Transcriptome profile comparison suggests that MAPKs and CDPKs are two convergence points of signaling triggered by most MAMPs (Boudsocq et al., 2010). Using a functional screen and diverse MAMP marker genes, this study revealed that CDPK and MAPK cascades act differentially in four MAMP-mediated regulatory programs (CDPK specific/MAPK specific/MAPK dominant and synergistic pathways) to control early genes involved in PTI. In addition, two transcription factors from the WRKY family, WRKY22 and WRKY29, were shown to act downstream of MPK3/MPK6 activation in response to flg22 (Asai et al., 2002).

Role of Silencing in Regulation of PTI

RNA silencing is an inducible defense pathway that uses small interfering RNAs (siRNAs) to specifically target and inactivate invading nucleic acids as a defense against viruses (Ruiz-Ferrer and Voinnet, 2009). Interestingly, there are also endogenous small RNAs that act in reprogramming gene expression in response to pathogen attack. For example, the microRNA miR393 is induced by flg22 in Arabidopsis and negatively regulates auxin signaling by targeting auxin receptors (Navarro et al., 2006). Repression of auxin signaling restricts P. syringae growth, implicating auxin in disease susceptibility and miRNA-mediated suppression of auxin signaling in resistance.

Consistent with a main role of small RNAs in immunity, several components of the silencing machinery were shown to be essential for pathogen resistance (Agorio and Vera, 2007; Navarro et al., 2008). Indeed, AGO1, a main component in the generation of small RNAs, seems to be required for some MAMP responses (Li et al., 2010). Analysis of AGO1-bound small RNAs led to the identification of a number of miRNAs up/downregulated by flg22 treatment; for some of them, a role in PTI was confirmed (Li et al., 2010). Future work will show to what extent small RNAs regulate PTI, and what their target genes are.

1.3.4 Final Outcome of the Response: HR or no HR?

The hypersensitive response (HR) is a form of rapid cell death that may restrict pathogen growth and is often associated with specific resistance (Jones and Dangl, 2006). Most MAMPs do not induce HR in plants; exceptions are the oomycete elicitins in tobacco (Takemoto et al., 2005) or fungal xylanase in tomato (Ron and Avni, 2004). In addition, recent studies have revealed that the prototypic MAMP flagellin can also induce HR in plant cells (Naito et al., 2008). Full-length flagellin from P. syringae pv tabaci 6605 induces an HR in Arabidopsis, in contrast to the classic flg22 from Pseudomonas aeruginosa; this is due to the presence of a single aspartate residue within the core region of the flg22 epitope (Naito et al., 2008). Interestingly, this residue is also important for bacterial virulence. Moreover, although flg22 does not induce cell death in wild type cells from rice, it induces HR in transgenic rice cells overexpressing OsFLS2 (Takai et al., 2008). Thus, the absence or presence of plant cell death should not be used as reliable criterion to distinguish between MAMPs and effectors.

1.3.5 Stomatal Closure