Plant Physiology

Hawes, Martha; Ronald, Pamela

doi: 10.1104/pp.109.900297pmid: 19656918

The incorporation of resistance genes into agronomically important crop plants is the most economically effective method for controlling plant disease. This biological disease control strategy is heritable and, therefore, inexpensive and permanently available once introduced (Keen et al., 1993). In 1993, Noel Keen and colleagues wrote an insightful review that outlined the benefits to be gained if only cloned resistance genes were available for deployment against plant pathogens. Practically before the ink was dry, the doorway to a new era opened with the news from Steve Tanksley's laboratory at Cornell that the tomato PTO gene conferring resistance to strains of the bacterial pathogen Pseudomonas syringae pv tomato carrying the corresponding avirulence gene AvrPto was a kinase (Ronald et al., 1992; Martin et al., 1993). This discovery was quickly followed by cloning and functional characterization of a number of diverse classes of genes for resistance to other bacterial pathogens, including intracellular proteins carrying nucleotide-binding sites and Leu-rich repeat motifs (NBS-LRRs; Bent et al., 1994; Mindrinos et al., 1994; Whitham et al., 1994) and extracellular pattern recognition receptors (PRRs; Song et al., 1995). Many plant pathologists now recognize two broad classes of the plant immune system termed pathogen-triggered immunity controlled by PRRs and effector-triggered immunity controlled by NBS-LRRs (Chisholm et al., 2006; Jones and Dangl, 2006). PRRs respond to microbe- or pathogen-associated molecular patterns that are highly conserved within a class of microbes such as flagellin (Medzhitov and Janeway, 1997; Gómez-Gómez and Boller, 2000). The NBS-LRR proteins recognize pathogen effectors, typically secreted by type III secretion systems. This focus issue provides an update on the tools now in hand to combat bacterial pathogens and the insights gained in the ensuing years. With complete genome sequences of several host and pathogen partners now available, there are literally hundreds of candidate genes with potential applications in crop protection. These include genes with sequence similarity to known NBS-LRR genes and PRRs as well as genes controlling plant responses to hormones involved in disease resistance responses. In addition to well-established players including jasmonic acid, ethylene, and salicylic acid, Lamb and coworkers report in the current issue that altered expression of genes controlling abscisic acid synthesis can increase resistance to certain pathogens while increasing susceptibility in others. Bent and coworkers note that the ability of plants to respond rapidly with global changes in physiology was recognized decades before any cloned genes were available; and while there are “few truly new questions” to be asked, focusing attention on underexplored niches in research and biology is likely to amplify the information to be gained. On the pathogen side, Collmer and colleagues consider that the greatest impact of a genomics approach has been the discovery that pathogens express many genes encoding many known or predicted effectors. They suggest that the greatest challenge will be to define how they work together to facilitate access to their host plants. Discovery of the “HRP” cluster and its relationship with the Yersinia outer membrane proteins in 1986 led quickly to the recognition that type III secretion is a central player in plant pathogenesis (Galan and Collmer, 1999). Expression of a single secreted protein, harpin, could confer the ability to induce the cell death response that is a hallmark of disease resistance, and harpin was soon made available for commercial disease control (Wei et al., 1992). This rapid translation of discovery into application made it appear that solutions to plant disease would be forthcoming with great efficiency. At present, exactly how or even where in the cell (or outside the cell, as the case may be) harpin may function remains under investigation. Michelmore and colleagues used a global profiling analysis to show that many effectors, like harpin, can trigger defense responses in diverse species. On the other hand, functional redundancy means that most are dispensable and, as White and coworkers put it, challenges for the future include establishing whether they work as “wrecking balls or guided missiles” in subverting plant metabolism. The Agrobacterium system continues to be in a class by itself with regard to insights into cross-species transport and transformation, fundamental plant physiology and development, as well as applications ranging from basic research to crop improvement. LITERATURE CITED Bent AF, Kunkel BN, Dahlbeck D, Brown KL, Schmidt R, Giraudat J, Leung J, Staskawicz BJ ( 1994 ) RPS2 of Arabidopsis thaliana: a leucine-rich repeat class of plant disease resistance genes. Science 265 : 1856 – 1860 Crossref Search ADS PubMed Chisholm ST, Coaker G, Day B, Staskawicz BJ ( 2006 ) Host-microbe interactions: shaping the evolution of the plant immune response. Cell 124 : 803 – 814 Crossref Search ADS PubMed Galan JE, Collmer A ( 1999 ) Type III secretion machines: bacterial devices for protein delivery into host cells. Science 284 : 1322 – 1328 Crossref Search ADS PubMed Gómez-Gómez L, Boller T ( 2000 ) FLS2: an LRR receptor-like kinase involved in the perception of the bacterial elicitor flagellin in Arabidopsis. Mol Cell 5 : 1003 – 1011 Crossref Search ADS PubMed Jones J, Dangl J ( 2006 ) The plant immune system. Nature 444 : 323 – 329 Crossref Search ADS PubMed Keen NT, Bent A, Staskawicz B ( 1993 ) Plant disease resistance genes: interactions with pathogens and their improved utilization to control plant diseases. In I Chet, ed, Biotechnology in Plant Disease Control. Wiley-Liss, New York, pp 65–68 Martin GB, Brommonschenkel SH, Chunwongse J, Frary A, Ganal MW, Spivey R, Wu T, Earle ED, Tanksley SD ( 1993 ) Map-based cloning of a protein kinase gene conferring disease resistance in tomato. Science 262 : 1432 – 1436 Crossref Search ADS PubMed Medzhitov R, Janeway CR ( 1997 ) Innate immunity: impact on the adaptive immune response. Curr Opin Immunol 9 : 4 – 9 Crossref Search ADS PubMed Mindrinos M, Katagiri F, Yu GL, Ausubel FM ( 1994 ) The A. thaliana disease resistance gene RPS2 encodes a protein containing a nucleotide-binding site and leucine-rich repeats. Cell 78 : 1089 – 1099 Crossref Search ADS PubMed Ronald PC, Salmeron JM, Carland FM, Staskawicz BJ ( 1992 ) The cloned avirulence gene avrPto induces disease resistance in tomato cultivars containing the Pto resistance gene. J Bacteriol 174 : 1604 – 1611 Crossref Search ADS PubMed Song WY, Wang GL, Chen LL, Kim HS, Pi LY, Holsten T, Gardner J, Wang B, Zhai WX, Zhu LH, et al ( 1995 ) A receptor kinase-like protein encoded by the rice disease resistance gene, Xa21. Science 270 : 1804 – 1806 Crossref Search ADS PubMed Wei ZM, Laby RJ, Zumoff CH, Bauer DW, He SY, Collmer A, Beer SV ( 1992 ) Harpin, elicitor of the hypersensitive response produced by the plant pathogen Erwinia amylovora. Science 257 : 85 – 88 Crossref Search ADS PubMed Whitham S, Dinesh-Kumar SP, Choi D, Hehl R, Corr C, Baker B ( 1994 ) The product of the tobacco mosaic virus resistance gene N: similarity to toll and the interleukin-1 receptor. Cell 78 : 1101 – 1115 Crossref Search ADS PubMed Author notes www.plantphysiol.org/cgi/doi/10.1104/pp.109.900297 © 2009 American Society of Plant Biologists This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Collmer, Alan; Schneider, David J.; Lindeberg, Magdalen

doi: 10.1104/pp.109.140327pmid: 19515788

Virtually all crop plants are attacked by pathogenic microbes, including bacteria, fungi, oomycetes, and nematodes. In many cases, bacterial diseases are still poorly controlled with century-old agents like copper, and cause serious losses, as seen with the recent citrus canker outbreak in Florida (Schubert et al., 2001). Pathogenicity has evolved independently in diverse phylogenetic lineages within the bacteria, resulting in a range of pathogenic lifestyles that parallels those that have evolved in the fungi and oomycetes. Thus, the study of bacterial pathogens is casting light into the interactions of plants with all microbes. This window of investigation was flung open in 2000 with publication of the first complete genome sequence of a phytopathogen, Xylella fastidiosa 9a5c (Simpson et al., 2000). Because it is a xylem-limited, nutritionally fastidious pathogen, its mechanisms of pathogenicity had been a complete mystery prior to analysis of the genome sequence. Subsequent sequencing of experimentally tractable pathogens in the genera Ralstonia (Salanoubat et al., 2002), Xanthomonas (Da Silva et al., 2002), Pseudomonas (Buell et al., 2003), and Pectobacterium (Bell et al., 2004) had a similarly revolutionary impact, revealing unexpected complexities in their virulence systems. Of particular importance to plant biologists is the genome-enabled, comprehensive identification of proteins and toxins that directly interact with plants and are referred to here as effectors (Hogenhout et al., 2009). Because effectors act during infection and outside of the bacterium, genes encoding them can be systematically identified on the basis of expression in planta and passage of their products through secretion pathways known to be trafficked by virulence proteins. Of the seven secretion systems described to date in gram-negative bacteria, the type II secretion system (T2SS) and type III secretion system (T3SS) are responsible for extracellular localization of the majority of critical virulence factors. Agrobacterium tumefaciens, an important pathogen that relies on the type IV pathway to deliver nucleoprotein T-DNA complexes into plant cells, has also been sequenced (Goodner et al., 2001; Wood et al., 2001) and is addressed in another Update article in this Focus issue. Small molecule effector candidates, which do not typically rely on a dedicated secretion system, can be identified through characterization of nonribosomal peptide and polyketide synthetases identified in the genome (Bender et al., 1999). Since each effector interacts with at least one plant molecule, and these targets involve diverse organelles and metabolic processes, the newly expanded collection of effectors provides a vast new tool box for plant biologists, revealing the unexpected extent of pathogen manipulations of plants (Speth et al., 2007). Importantly, complete genome sequences also enable investigation of how all of the effector parts work together to enable parasitism in different plants and specific plant niches. This Update highlights insights gained from fully sequenced bacterial pathogen genomes that are of particular relevance to plant biologists. We will describe the range of bacterial phytopathogens and their lifestyles in plants, lessons gained from type III effector repertoires (a focus of much study during this period), major insights arising from each of the phytopathogen groups with completely sequenced genomes, and future challenges. THE DIVERSITY OF PHYTOPATHOGENIC BACTERIA AND AN EMERGING BIG PICTURE OF THEIR INTERACTIONS WITH PLANTS Plant pathogens of all classes are now considered to have two broadly different pathogenic lifestyles, with necrotrophs gaining nutrients from rapidly killed tissue and biotrophs gaining nutrients from living host tissue (or in the case of hemibiotrophs, from living tissue that dies in a later stage of pathogenesis; Glazebrook, 2005). The phytopathogenic bacteria with complete and published sequences span this continuum of pathogenic lifestyles, summarized in Table I Table I. Salient properties of representative phytopathogenic bacteria with published genome sequences Host names followed with an ellipsis denote a broader host range. Column headings: V, obligately vectored by insects (as denoted by +); T, tissue primarily colonized in plant (M, mesophyll; X, xylem; P, phloem); CW, cell wall-degrading enzymes; TE, translocated effectors acting within plant cells; SP, extracellular Ser proteases; SM, small molecules. Symbols in Effectors columns: +, one or a few present; ++ many present and major role; −, none present; blank cell, not studied. References are in text. Bacterial Groups and Representative Pathogens . Disease . Host(s) . V . T . Effectors . . . . . . . . . CW . TE . SP . SM . Proteobacteria (necrotrophs)     P. atrosepticum Black leg Potato MX ++ + + Proteobacteria (hemibiotrophs)     P. syringae pv tomato Bacterial speck Tomato, Arabidopsis, other Brassica M + ++ +     R. solanacearum Bacterial wilt Tomato, Arabidopsis… X ++ ++ +     X. axonopodis pv citri Canker Citrus M ++ ++     X. campestris pv campestris Black rot Arabidopsis, other Brassica X ++ ++     X. fastidiosa Variegated chlorosis Citrus + X + − Actinobacteria     C. michiganensis subsp. michiganensis Wilt and canker Tomato X + − ++     C. michiganensis subsp. sepedonicus Ring rot Potato X + − ++     L. xyli subsp. xyli Ratoon stunting Sugarcane + X + − + Firmicutes     Candidatus P. asteris Yellows Wide host range + P − + Bacterial Groups and Representative Pathogens . Disease . Host(s) . V . T . Effectors . . . . . . . . . CW . TE . SP . SM . Proteobacteria (necrotrophs)     P. atrosepticum Black leg Potato MX ++ + + Proteobacteria (hemibiotrophs)     P. syringae pv tomato Bacterial speck Tomato, Arabidopsis, other Brassica M + ++ +     R. solanacearum Bacterial wilt Tomato, Arabidopsis… X ++ ++ +     X. axonopodis pv citri Canker Citrus M ++ ++     X. campestris pv campestris Black rot Arabidopsis, other Brassica X ++ ++     X. fastidiosa Variegated chlorosis Citrus + X + − Actinobacteria     C. michiganensis subsp. michiganensis Wilt and canker Tomato X + − ++     C. michiganensis subsp. sepedonicus Ring rot Potato X + − ++     L. xyli subsp. xyli Ratoon stunting Sugarcane + X + − + Firmicutes     Candidatus P. asteris Yellows Wide host range + P − + Open in new tab Table I. Salient properties of representative phytopathogenic bacteria with published genome sequences Host names followed with an ellipsis denote a broader host range. Column headings: V, obligately vectored by insects (as denoted by +); T, tissue primarily colonized in plant (M, mesophyll; X, xylem; P, phloem); CW, cell wall-degrading enzymes; TE, translocated effectors acting within plant cells; SP, extracellular Ser proteases; SM, small molecules. Symbols in Effectors columns: +, one or a few present; ++ many present and major role; −, none present; blank cell, not studied. References are in text. Bacterial Groups and Representative Pathogens . Disease . Host(s) . V . T . Effectors . . . . . . . . . CW . TE . SP . SM . Proteobacteria (necrotrophs)     P. atrosepticum Black leg Potato MX ++ + + Proteobacteria (hemibiotrophs)     P. syringae pv tomato Bacterial speck Tomato, Arabidopsis, other Brassica M + ++ +     R. solanacearum Bacterial wilt Tomato, Arabidopsis… X ++ ++ +     X. axonopodis pv citri Canker Citrus M ++ ++     X. campestris pv campestris Black rot Arabidopsis, other Brassica X ++ ++     X. fastidiosa Variegated chlorosis Citrus + X + − Actinobacteria     C. michiganensis subsp. michiganensis Wilt and canker Tomato X + − ++     C. michiganensis subsp. sepedonicus Ring rot Potato X + − ++     L. xyli subsp. xyli Ratoon stunting Sugarcane + X + − + Firmicutes     Candidatus P. asteris Yellows Wide host range + P − + Bacterial Groups and Representative Pathogens . Disease . Host(s) . V . T . Effectors . . . . . . . . . CW . TE . SP . SM . Proteobacteria (necrotrophs)     P. atrosepticum Black leg Potato MX ++ + + Proteobacteria (hemibiotrophs)     P. syringae pv tomato Bacterial speck Tomato, Arabidopsis, other Brassica M + ++ +     R. solanacearum Bacterial wilt Tomato, Arabidopsis… X ++ ++ +     X. axonopodis pv citri Canker Citrus M ++ ++     X. campestris pv campestris Black rot Arabidopsis, other Brassica X ++ ++     X. fastidiosa Variegated chlorosis Citrus + X + − Actinobacteria     C. michiganensis subsp. michiganensis Wilt and canker Tomato X + − ++     C. michiganensis subsp. sepedonicus Ring rot Potato X + − ++     L. xyli subsp. xyli Ratoon stunting Sugarcane + X + − + Firmicutes     Candidatus P. asteris Yellows Wide host range + P − + Open in new tab . The soft-rot enterobacterium Pectobacterium atrosepticum SCRI1043 (Bell et al., 2004) was the first sequenced representative of the necrotrophs. The pioneer hemibiotrophs were Ralstonia solanacearum GMI1000 (Salanoubat et al., 2002), Xanthomonas campestris pv campestris ATCC33913 (Da Silva et al., 2002), Xanthomonas axonopodis pv citri 306 (Da Silva et al., 2002), and Pseudomonas syringae pv tomato DC3000 (Buell et al., 2003). Although sharing the basic property of dependence on type III effectors, the hemibiotrophic pathogens have interesting differences in their interactions with plants. For example, R. solanacearum is a wilt pathogen with a potentially broad host range that colonizes the xylem after entering plants through roots. X. campestris pv campestris also can colonize the xylem, but it attacks aerial parts of the plant and has a narrow host range. In further contrast, X. axonopodis pv citri and P. syringae are narrow-host-range pathogens that colonize the mesophyll rather than the xylem. All of these bacteria are gram negative, with R. solanacearum in the β-Proteobacteria and the others in the γ-Proteobacteria. Clavibacter michiganensis subsp. michiganensis NCPPB382 (Gartemann et al., 2008), C. michiganensis subsp. sepedonicus ATCC33113 (Bentley et al., 2008), and Leifsonia xyli subsp. xyli CTCB07 (Monteiro-Vitorello et al., 2004) are the pioneer, sequenced bacteria in the high G + C content, gram-positive group Actinobacteria. These bacteria are closely related and are all xylem colonizers. Analogous to X. fastidiosa, L. xyli subsp. xyli is a fastidious bacterium with a small genome size (2.6 Mb) that appears to be a product of reductive evolution (Moreira et al., 2004). The end point in the continuum toward biotrophic specialization for life within a specific plant niche and associated genome reduction is represented by wall-less bacteria in the low G + C content, gram-positive group, Firmicutes. These bacteria are obligate colonizers of the phloem and their phloem-feeding insect vectors (Hogenhout and Loria, 2008). Notably, the Candidatus Phytoplasma asteris OY-M pioneer genome is only 860 kb (Oshima et al., 2004). In contrast, the largest genome among this pioneering set of phytopathogens is that of P. syringae pv tomato DC3000, which is 6.5 Mb. A conceptually and agriculturally significant feature of these pathogenic lifestyles and phylogenetic groups is that the plant defenses against necrotrophs and (hemi)biotrophs are different and antagonistic. Specifically, major gene resistance is only effective against hemibiotrophic gram-negative bacteria injecting type III effectors that are recognized by cytoplasmic resistance (R) proteins in the host (Spoel et al., 2007; Poland et al., 2009). A recently formulated model for the interactions of plants with hemibiotrophic Proteobacteria has broad explanatory power (Jones and Dangl, 2006; Gohre and Robatzek, 2008). According to this model, bacteria in the apoplast display pathogen (or microbe)-associated molecular patterns (PAMPs), such as flagellin, lipopolysaccharide, peptidoglycan, and elongation factor EF-Tu, which are recognized at the plant cell surface by pattern recognition receptors that elicit PAMP-triggered immunity (PTI). Pathogens defeat this defense by injecting type III effectors that suppress PTI. The T3SS, required for delivery of these effectors and thus essential to virulence in all hemibiotrophic Proteobacteria pathogens, is discussed in another Update in this issue. Although effectors suppress PTI, they may also elicit defenses. Specifically, if the host carries an appropriate R gene for detecting the activity of one or more of these effectors inside plants cells, effector-triggered immunity (ETI) is activated, typically manifested as a defense-related programmed cell death referred to as the hypersensitive response. PTI is addressed in more detail in another Update in this issue but this model, based on plant interaction with hemibiotrophic Proteobacteria, raises important questions about other groups of phytopathogenic bacteria. Are PAMPs also perceived in the xylem or the phloem? How does Clavibacter, which does not appear to inject effectors, evade or suppress PTI? How do necrotrophs defeat PTI? What determines the differing levels of host specificity and tissue specificity observed in various pathogen groups? THE TYPE III EFFECTOR REPERTOIRES OF HEMIBIOTROPHIC PROTEOBACTERIA The greatest impact of bacterial phytopathogen genome sequencing so far has been the discovery of large numbers of type III effectors in various phytopathogenic Proteobacteria. Because of gene duplication and functional redundancy, many type III effectors are individually dispensable. In hindsight, it is not surprising that the corresponding genes were largely missed in pregenomics era screens for mutants with reduced virulence. Genomics bypassed this problem by enabling various functional screens and characterization of sequence patterns to identify all probable candidates in each strain, many of which have been experimentally validated. Patterns used have included promoter motifs, amino acid biases associated with type III targeting signals, motifs predicting eucaryote-like functions, and presence in genomic islands (Cunnac et al., 2004; Lindeberg et al., 2006; Vinatzer and Yan, 2008; Furutani et al., 2009). These efforts have revealed complex effector repertoires for several strains, as well as provisional super repertoires for the pangenomes of three phytopathogen groups, as tabulated in recent reviews addressing Xanthomonas spp. (Kay and Bonas, 2009), R. solanacearum (Poueymiro and Genin, 2009), and the P. syringae pathovars (Cunnac et al., 2009). Effector repertoires are highly variable, even for pathogens of a single plant species. The relative size and variability of these repertoires can be seen in three well-studied tomato (Solanum lycopersicum) pathogens: X. campestris pv vesicatoria 85 to 10 (17 effectors confirmed), R. solanacearum GMI1000 (28 confirmed + 46 candidates), and P. syringae pv tomato DC3000 (28 confirmed; Cunnac et al., 2009; Kay and Bonas, 2009; Poueymiro and Genin, 2009). Of the 17 effectors injected by X. campestris pv vesicatoria, five have homologs in R. solanacearum and three have homologs in P. syringae pv tomato (Kay and Bonas, 2009). This variability in type III effector repertoires is particularly striking in two strains of P. syringae pv tomato: DC3000 and T1. Both strains cause bacterial speck of tomato, but only half of their effector repertoires are shared (Almeida et al., 2009). Complete genome sequences are now published for seven Xanthomonas strains representing diverse species and pathovars, and three pathovars of P. syringae (Da Silva et al., 2002; Buell et al., 2003; Feil et al., 2005; Joardar et al., 2005; Lee et al., 2005; Qian et al., 2005; Thieme et al., 2005; Salzberg et al., 2008; Vorholter et al., 2008). Comparison of their effector repertoires similarly reveals a high degree of variability, with no obvious correlation between repertoire composition and host specificity or tissue specificity (Salzberg et al., 2008; Almeida et al., 2009). Thus, it appears that there are many ways for bacteria to defeat plant defenses with type III effectors. Enormous progress has been made in identifying the diverse biochemical activities, subcellular targets, and host interactors of type III effectors. These properties of individual effectors are summarized in recent reviews (Block et al., 2008; Gohre and Robatzek, 2008; Cunnac et al., 2009; Kay and Bonas, 2009; Poueymiro and Genin, 2009), and we are limited here to a few highlights that indicate the scope and sophistication of the effector assault on plants. In general, type III effectors manipulate host cell protein turnover, RNA synthesis and stability, and protein phosphorylation (Block et al., 2008). Effectors can insert themselves into host protein processing and targeting pathways, resulting in localization to the plasma membrane, chloroplasts, the nucleus, and other subcellular sites (Gohre and Robatzek, 2008; Dowen et al., 2009). Their molecular targets can range from pattern recognition receptors essential for PAMP perception to promoters for genes controlling cell size (Gohre and Robatzek, 2008). The biological function of most type III effectors appears to be suppression of PTI and/or ETI, and some effectors, such as AvrPtoB, have multiple domains that suppress both defenses (Abramovitch and Martin, 2005; Rosebrock et al., 2007). The bewildering range of activities and subcellular and molecular targets of type III effectors is best appreciated by scanning the tables of recent reviews (as cited above). However, it is worth noting that only a small fraction of the known effectors have been characterized, and the known effectors represent only a fraction of the total that are likely encoded in the pangenomes of the hemibiotrophic Proteobacteria. THE IMPORTANCE OF REDUNDANT EFFECTOR GROUPS IN PHYTOPATHOGENIC BACTERIA The model described above of pathogenesis based on translocated effectors suppressing PTI while being under R protein surveillance evokes a coevolutionary war between plants and pathogens that can generate large and polymorphic repertoires of effectors and R proteins (McHale et al., 2006; Stavrinides et al., 2008). Importantly, this process appears to generate effector repertoires that are collectively essential but where individual effectors are typically dispensable. Because of this dispensability, effector genes can be lost with little or no virulence penalty by pathogen populations facing a cultivar that relies on R-gene-mediated resistance. Thus, R-gene-mediated resistance is often defeated after a few years of agricultural use, as has been observed with many important bacterial, fungal, and oomycete pathogens (Jones and Dangl, 2006; Poland et al., 2009). The availability of complete genome sequences and complete type III effector repertoires has enabled investigation of an important property of most effector repertoires, that is, the dispensability of individual effectors. Such dispensability has been inferred from the results of numerous mutant screens involving various hemibiotrophic Proteobacteria and has been systematically explored with R. solanacearum GMI1000 and P. syringae pv tomato DC3000. Mutagenesis of 42 effector and effector candidate genes in GMI1000 revealed that only two had a virulence phenotype in host tomato, manifested as only a slight delay in symptom development (Cunnac et al., 2004). Similarly, a screen of P. syringae pv tomato DC3000 transposon mutants with a sensitive virulence assay based on dip-inoculated Arabidopsis (Arabidopsis thaliana) plants yielded only a single effector gene, which only quantitatively contributes to lesion formation (Brooks et al., 2004; Badel et al., 2006). Through the efforts of multiple research groups, the type III effector repertoire of DC3000 has been particularly well established and is thought to comprise 28 actively deployed effectors (Lindeberg et al., 2006). Combinatorial deletions involving 20 of the active effector genes have revealed a redundancy-based structure in the effector repertoire, such that some deletions diminish growth in planta only in combination with other deletions (Kvitko et al., 2009). It was found that two redundant effector groups are particularly important in promoting DC3000 growth in planta, and based on the known activities of some of the members, these internally redundant groups were proposed to target different high-level processes in PTI: perception of PAMPs and vesicle trafficking of antimicrobial factors. These observations suggest that successful pathogenesis by DC3000 depends on blocking of a few key defense processes, with each process targeted redundantly. Host surveillance of the effectors and, possibly, the polymorphic nature of effector targets may explain the apparent need for redundancy. The phenomenon of redundant effector groups appears widespread in phytopathogenic bacteria. For example, the extracellular components of the T3SS that have been genetically implicated in forming the translocation pore in the host plasma membrane are conserved and individually essential in animal pathogens but more numerous, variable, and individually dispensable in plant pathogens (Kvitko et al., 2007). R. solanacearum and many Xanthomonas spp. also have candidate redundant effector groups represented by the multiple copies of F-box-containing GALA family effectors and transcription activator-like effectors that their respective genomes encode (Angot et al., 2006). Experiments thus far with both effector families indicate that combinatorial mutations are required to produce significant virulence phenotypes (Yang et al., 1996; Angot et al., 2006). As will be discussed in the next section, redundant groups of candidate effectors (broadly defined) can also be found in Pectobacterium and Clavibacter spp. Systematic identification of redundant effector groups has the potential to reveal processes within plants that contribute to defense against microbial pathogens and may provide clues to the specific plant proteins targeted by and/or involved in the recognition of individual effectors. For example, AvrPto and AvrPtoB, though showing no sequence similarity, are members of a redundant effector group that inhibits the kinase activity of the FLS2 pattern recognition receptor and interacts with Pto, now thought to function as a decoy kinase under R protein surveillance (Shan et al., 2008; Xiang et al., 2008; Zhou and Chai, 2008). Identification of redundant effector groups may also have practical utility in suggesting more durable combinations of R genes less likely to be overcome by pathogen mutation. Ultimately, the genomics-enabled characterization of type III effector repertoires and interacting plant proteins is serving to unravel the complex network of host-pathogen interactions in a manner that is biologically significant and agriculturally useful. OTHER LESSONS FROM PATHOGEN GENOMES The P. atrosepticum genome highlights two other important classes of effectors: pectic enzymes and phytotoxins. Like other soft-rot enterobacteria, P. atrosepticum (formerly Erwinia carotovora subsp. atroseptica) produces multiple pectic enzymes, secreted via the T2SS (Toth and Birch, 2005). Studies with various soft-rot enterobacteria involving mutations in T2SS genes and either single or multiple pectic enzyme genes suggest that pectic enzymes are collectively essential but individually dispensable for the maceration of parenchymatous plant tissues characteristic of soft-rot diseases (Toth et al., 2006). The P. atrosepticum genome sequence revealed 11 new putative pectic enzyme genes, thus bringing the repertoire to 20 (Bell et al., 2004). However, there is no evidence that plant surveillance or inhibitors are driving amplification of the P. atrosepticum pectic enzymes or causing diversification in the repertoire encoded by related species (Glasner et al., 2008). Subtle specialization for substrates and reaction conditions in the cell walls of different plants may account for the amplification of effectors in this class. In contrast, T2SS mutants are only partially reduced in virulence, and pectic enzymes appear to have only a minor role in P. syringae pathogenesis (Bauer and Collmer, 1997; Bronstein et al., 2005). A surprising discovery in the P. atrosepticum genome was the presence of homologs of P. syringae genes directing biosynthesis of the toxins syringomycin and coronafacic acid (Bell et al., 2004), mutations that strongly reduced black leg disease in potato (Solanum tuberosum; Bell et al., 2004). In P. syringae pv tomato DC3000, coronatine, an amide-linked conjugate of coronafacic acid and an Ile derivative, mimics jasmonic acid Ile, promotes the opening of stomates and bacteria entry, and suppresses salicylic acid-dependent plant defenses (Brooks et al., 2005; Melotto et al., 2008). The biosynthetic capacity for this toxin family is encoded in a horizontally acquired region in DC3000 and is present in only a subset of other P. syringae pathovars and in only P. atrosepticum among the sequenced Pectobacteria spp. (Hwang et al., 2005; Glasner et al., 2008; Lindeberg et al., 2008). Like P. syringae, P. atrosepticum encodes a functional T3SS, but the only effector candidate is a homolog of the widespread P. syringae effector AvrE (Bell et al., 2004). In summary, although P. atrosepticum and P. syringae are in different bacterial families, they appear to have acquired effector genes by horizontal transfer, with their respective effector repertoires being differently expanded in association with their distinct pathogenic lifestyles. Comparison of phylogenetically divergent organisms with similar pathogen lifestyles provides yet another opportunity for exploring the nature and evolution of bacterial pathogenesis through comparative genomics. For example, the >50 P. syringae pathovars and >100 Xanthomonas species/pathovars are all T3SS-dependent, host-specific pathogens that are often good epiphytes and commonly cause diseases characterized by scattered lesions on foliage (although some Xanthomonas spp. also invade the xylem and cause extensive tissue death). Most crops are attacked by at least one member of each group. Importantly, genome sequencing suggests that phytopathogenicity in these two genera has evolved convergently. For example, not only are their type III effector repertoires largely different, but they also possess independently acquired and distinct T3SS (Alfano and Collmer, 1997). Thus, functional genomic comparisons of strains with common hosts from the parallel pseudomonad and xanthomonad series could indicate the potential range of interactions plants may have with microbes that attack via translocated effector proteins. R. solanacearum represents a versatile pathogen, unique among the major bacterial phytopathogens given its ability to attack plants via the roots (Genin and Boucher, 2004) and particularly devastating to a variety of tropical crops. The bacterium has large repertoires of candidate effectors traveling the T2SS and the T3SS, and appropriate mutants indicate that the two pathways and their respective repertoires are important for pathogenesis (Poueymiro and Genin, 2009). The type III effector repertoire features three expanded families with multiple members as well as multiple effectors with repeat domains implicated in the recognition of host proteins (Poueymiro and Genin, 2009). The GALA proteins mentioned above provide a good example, with strain GMI1000 producing seven such proteins predicted to target host proteins for degradation via the 26S proteosome (Angot et al., 2006). Analogous to P. syringae pv tomato in its manipulation of hormone signaling with a small-molecule effector, GMI1000 produces ethylene, which affects the expression of ethylene-response host genes during infection (Valls et al., 2006). Genomic analyses suggest that effector families may be similarly subject to amplification in gram-positive pathogens as well. Pregenomics research had identified two C. michiganensis subsp. michiganensis proteins, CelA endo-β1 to 4 glucanase and Pat-1 Ser protease (Hogenhout and Loria, 2008), which appear to be effectors given their role in virulence and predicted extracellular location. One or more yet to be identified proteins in the extracellular fluids from both subspecies have additionally been shown to elicit the hypersensitive response in tobacco (Nicotiana tabacum; Nissinen et al., 1997; Alarcon et al., 1998). Genome sequences reveal that homologs of celA family members and pat-1 family members are present in subspecies michiganensis and sepedonicus, as well as in L. xyli subsp. xyli (Monteiro-Vitorello et al., 2004; Bentley et al., 2008; Gartemann et al., 2008). Intriguingly, subspecies michiganensis encodes at least 28 Ser proteases of which 10 are Pat-1-like members of the Chp family. These gene families have several characteristic properties of effector genes including an atypical G + C content suggestive of horizontal acquisition and a demonstrated role in elicitation of the hypersensitive response on nonhost plants for at least one of the family members. Subspecies sepedonicus harbors 11 Chp family genes, and L. xyli subsp. xyli has one. It is tempting to speculate that the Chp family Ser proteases are acting extracellularly to degrade either pattern recognition receptors or antimicrobial peptides/proteins and that ETI surveillance of these proteins or their activities has driven amplification of the family. The Chp Ser proteases thus could provide new clues to how plants defend the xylem against pathogens. It is also noteworthy that C. michiganensis subsp. michiganensis and L. xyli subsp. xyli encode tomatinase and a fatty acid desaturase, respectively. Tomatinase can release defense-suppressive products from the antifungal saponin tomatine (Bouarab et al., 2002), and the fatty acid desaturase may produce abscisic acid, which could contribute to the major symptom of ratoon stunting of sugarcane (Saccharum officinarum; Monteiro-Vitorello et al., 2004). Thus, both of these proteins have the potential to generate small-molecule effectors. In contrast to the other pathogens discussed, the phytoplasmas can inhabit the intracellular space of both insects and plants and may evade plant defenses in large part through absence of PTI-triggering PAMPs that have been lost during genome reduction. Nonetheless, identification of extracellular effector candidates on the basis of conserved targeting motifs (in this case, N-terminal signal peptides facilitating secretion via a functional Sec pathway) represents a valuable approach for effector identification. Eukaryotic nuclear localization signals have been identified in two of the effector candidates with SAP11 from Candidatus P. asteris AY-WB shown to induce necrosis and alter transcription (Bai et al., 2006; Hogenhout and Oshima, 2008). NEW CHALLENGES A primary challenge of genomics research involves discovery of patterns in the DNA and protein sequences. Fortunately, because typical effector genes encoding type III effectors, cell wall-degrading enzymes, extracellular proteases, and biosynthetic enzymes for toxins and phytohormones carry a variety of predictive patterns, we have been able to make substantial progress toward comprehensive identification of effectors in the pioneer pathogen genomes. The subsequent sequencing of strains related to the pioneers has enabled a second round of pattern searching focused on differences in virulence and host or tissue specificity and an initial glimpse of the pangenomes for several species. In the near future, we can expect publication of pioneer genome sequences for other important pathogens, such as Streptomyces scabies, Erwinia amylovora, Pantoea stewartii, and Dickeya dadantii (formerly Erwinia chrysanthemi). Furthermore, next-generation sequencing methods have the potential to yield low-cost draft genomes for a virtually unlimited set of relatives for each pioneer. These advances, coupled with continuing refinements in the iterative process of effector function analysis and pattern recognition, should yield the complete effecterome for each of these pathogen groups. The next fundamental challenge will be to discern patterns in these effector repertoires that underlie their evolutionary assembly into viable systems for defeat of host defenses and adaption to plant-associated niches. For example, it appears that diverse phytopathogenic bacteria (with the possible exception of the Firmicutes) produce both protein effectors and small molecule effectors. Do these two classes of effectors work coordinately? And, how do effector repertoires function in coordination with the rest of the bacterial genome and physiology? In this regard, it is important to note that effector repertoire composition has so far failed to explain either the host or tissue specificity of different members of the hemibiotrophic Proteobacteria. Furthermore, although R protein surveillance of type III effectors certainly explains race-cultivar specificity in the field, it may not explain the specificity of P. syringae pathovars, R. solanacearum strains, or Xanthomonas spp. for their different plant species. The latter specificity is generally stable in the field despite the observation that loss of just one or two effectors can expand host range to new plant species (Castaneda et al., 2005; Lin and Martin, 2007; Wei et al., 2007; Poueymiro et al., 2009). It is possible that multiple adaptations involving PAMP perception, nutrition, and antimicrobial factors underlie host and tissue specificity. The ultimate challenge of effector identification and functional characterization involves the integration of their various individual roles into a comprehensive picture of host-pathogen interaction. As one means of managing the increasingly complex data on DC3000 effectors, Gene Ontology annotation is being used to systematically document the biological processes, molecular functions, and cellular locations of individual effectors, enabling comparison among effectors deployed by a single strain as well as among those deployed during the course of other, diverse host-pathogen interactions. LITERATURE CITED Abramovitch RB, Martin GB ( 2005 ) AvrPtoB: a bacterial type III effector that both elicits and suppresses programmed cell death associated with plant immunity. FEMS Microbiol Lett 245 : 1 – 8 Crossref Search ADS PubMed Alarcon C, Castro J, Monoz F, Arece-Johnson P, Delgado J ( 1998 ) Protein(s) from the gram-positive bacterium Clavibacter michiganensis subsp. michiganensis induces a hypersensitive response in plants. 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Mol Plant Microbe Interact 9 : 105 – 113 Crossref Search ADS Zhou JM, Chai J ( 2008 ) Plant pathogenic bacterial type III effectors subdue host responses. Curr Opin Microbiol 11 : 179 – 185 Crossref Search ADS PubMed Author notes 1 This work was supported by the National Science Foundation Plant Genome Research Program (grant no. DBI–0605059). * Corresponding author; e-mail [email protected]. The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Magdalen Lindeberg ([email protected]). www.plantphysiol.org/cgi/doi/10.1104/pp.109.140327 © 2009 American Society of Plant Biologists This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Allen, Caitilyn; Bent, Andrew; Charkowski, Amy

doi: 10.1104/pp.109.140004pmid: 19561122

SUMMARY Despite rapid advances on certain aspects of plant pathogenic bacteria, many economically important pathosystems are largely unexplored and biologically relevant life stages of even familiar systems remain poorly understood. We know remarkably little about end-stage disease, latent infections, survival away from the host, interactions among multiple microbes in a plant, and the effects of quantitative virulence factors. While no thoughtful researcher would dispute the effectiveness of reductionist experiments, we propose that this approach be combined with a broader perspective that includes the ecology, histopathology, and community population biology of phytopathogenic bacteria. We offer examples of exciting recent discoveries resulting from this natural history-based approach. In particular, in situ studies using biologically realistic inoculation followed by analyses with microscopy, gene expression profiling, community analyses, or application of key computational tools can offer new insights into old questions. Research that combines cutting-edge tools with a biological perspective is especially lacking on high-impact diseases of subsistence crops. Understanding the biology underlying important practical issues such as copper resistance, eradication from seed and cuttings, and rapid, sensitive detection could be of significant utility. Overall, we endorse a broader biological approach to research on plant pathogenic bacteria. CHOICES, CHOICES In response to significant advances in plant bacteriology, researchers can focus in to more deeply understand the discovery, or they can change the subject and turn to important questions that remain poorly understood. This article encourages the second approach by pointing out some underexplored but important aspects of plant pathogenic bacteria. We first discuss considerations that may aid selection of research topics, and then suggest a necessarily incomplete set of specific questions and approaches that promise fresh and productive research. Ideally, our research programs would be designed to reveal fundamental biology of high-impact plant pathogens, leading to useful disease management strategies. All too often our planning instead brings us to the intersection of the feasible, the fundable, and the familiar—hardly a path to novelty. We suggest that those seeking new directions should instead choose a study system that satisfies at least two of the criteria listed in Table I Table I. Some criteria to identify novel and important systems for research Major disease of major staple crop Disease of understudied staple crop (e.g. plantains [Musa paradisiaca], oil palms [Elaeis spp.], cassava) Major disease of high-value specialty crop or developing nation crop Effective disease management would expand cropping zone Commodity group or international non-government organization support Current control methods environmentally undesirable Pathogen persistence in environment Pathogen colonization of plant surface or vasculature Pathogen latent or commensal stage Pathogen seed transmissibility Pathogen insect transmissibility System has unique biology (e.g. Agrobacterium tumefaciens) Plant-associated human pathogen Pathosystem has potential impact on medical biology Major disease of major staple crop Disease of understudied staple crop (e.g. plantains [Musa paradisiaca], oil palms [Elaeis spp.], cassava) Major disease of high-value specialty crop or developing nation crop Effective disease management would expand cropping zone Commodity group or international non-government organization support Current control methods environmentally undesirable Pathogen persistence in environment Pathogen colonization of plant surface or vasculature Pathogen latent or commensal stage Pathogen seed transmissibility Pathogen insect transmissibility System has unique biology (e.g. Agrobacterium tumefaciens) Plant-associated human pathogen Pathosystem has potential impact on medical biology Open in new tab Table I. Some criteria to identify novel and important systems for research Major disease of major staple crop Disease of understudied staple crop (e.g. plantains [Musa paradisiaca], oil palms [Elaeis spp.], cassava) Major disease of high-value specialty crop or developing nation crop Effective disease management would expand cropping zone Commodity group or international non-government organization support Current control methods environmentally undesirable Pathogen persistence in environment Pathogen colonization of plant surface or vasculature Pathogen latent or commensal stage Pathogen seed transmissibility Pathogen insect transmissibility System has unique biology (e.g. Agrobacterium tumefaciens) Plant-associated human pathogen Pathosystem has potential impact on medical biology Major disease of major staple crop Disease of understudied staple crop (e.g. plantains [Musa paradisiaca], oil palms [Elaeis spp.], cassava) Major disease of high-value specialty crop or developing nation crop Effective disease management would expand cropping zone Commodity group or international non-government organization support Current control methods environmentally undesirable Pathogen persistence in environment Pathogen colonization of plant surface or vasculature Pathogen latent or commensal stage Pathogen seed transmissibility Pathogen insect transmissibility System has unique biology (e.g. Agrobacterium tumefaciens) Plant-associated human pathogen Pathosystem has potential impact on medical biology Open in new tab . In particular, research is urgently needed on destructive diseases of key tropical subsistence crops, such as Xanthomonas wilt of banana (Musa spp.) and bacterial blight of cassava (Manihot esculenta). A more widespread focus on research to reduce crop losses offers the additional benefit of increasing stakeholder support for plant bacteriology funding. LOOK BACK TO MOVE FORWARD New knowledge and methods create opportunities for progress on old questions, and indeed there are few truly new questions. It is humbling to discover that our scientific predecessors thought deeply and usefully about our subject. Perceptive articles and book chapters that were written long before the advent of PubMed can be overlooked in an online search. Readers curious about plant pathogenic bacteria are encouraged to explore the following and other older sources, which describe key research questions that remain unsolved (Smith, 1920; Walker, 1963; Schuster and Coyne, 1974; Vidaver, 1981; Mount and Lacy, 1982; Starr, 1984; Billing, 1987; Nester et al., 2004). In the same spirit, readers are encouraged to remain open to the curiosity about the natural world that drew us to science. Fancy tools are one route to novel findings, but paradigm-shifting discoveries often come from simple observation. Charles Darwin had travel funds, notebooks, pencils, and a few dead birds. THE PENDULUM IS SWINGING TOWARD DIVERSITY The early years of molecular plant bacteriology explored a wide range of interactions. Conducted without kits, PCR, or commercial DNA sequencing, this research used laborious methods such as screening and characterization of transposon mutants to discover hrp (for host response and pathogenicity) and avr (for avirulence) genes in the interactions between Pseudomonads and bean (Phaseolus vulgaris) plants, dissect the role of cell wall-degrading enzymes in soft-rot enterobacteria, and determine that EPS is key to wilt pathogenesis (Staskawicz et al., 1984; Niepold et al., 1985; Lindgren et al., 1986; Kotoujansky, 1987; Denny and Baek, 1991). The rapid discovery of the unique mechanisms underlying crown gall disease demonstrated how quickly an area could advance given significant investment and competition (Zambryski, 1988). This insight, together with the rise of Arabidopsis (Arabidopsis thaliana) as a host and enthusiasm for model systems in general, has drawn molecular plant bacteriologists to a narrow set of pathosystems. Grant proposals often argue that results obtained with these organisms will be easily applied to economically important plant diseases. However, the pioneering bacterial geneticist Jacques Monod was famously wrong when he said that “anything found to be true of E. coli must also be true of Elephants” (Monod and Jacob, 1961, p. 393). While many mechanisms are common across biological systems, even closely related organisms have adapted and shaped ancestral tools to diverse ends, solving similar problems in strikingly different ways or adapting the same protein for unrelated functions. On the heels of success with model systems, a renewed effort to study diverse plant-bacterial systems is needed precisely because what is true for DC3000 in an Arabidopsis leaf is not always true for Xylella fastidiosa in a grapevine (Vitis spp.). Understanding gene-for-gene resistance to bacterial blight of rice (Oryza sativa) is important, but it is unlikely to elucidate horizontal resistance to bacterial wilt in tomato (Solanum lycopersicum) or lead to greening-tolerant citrus trees (Citrus spp.). BACTERIAL BEHAVIOR IN NATURAL HOSTS UNDER BIOLOGICALLY REALISTIC CONDITIONS IS UNDEREXPLORED Reductionist experiments are powerful, but the lure of their yes/no results can keep us from doing discovery experiments that may be complicated and messy but also more biologically realistic and practically relevant. Familiar examples include studies focusing on single genes rather than multigenic traits, model systems instead of natural hosts, sterile potting mix in place of natural soil, seedlings rather than mature plants, and controlled rather than field environments. Every researcher struggles to balance experimental feasibility with biological meaning, but a convenient and familiar assay can give deceptive results that hide a more interesting truth. For example, He and coworkers found that the phytotoxin coronatine facilitates pathogen entry into leaf mesophyll by causing stomates to open, but this effect was masked if leaves were infiltrated with bacteria and was only detectable when Pseudomonas syringae strains were inoculated onto leaf surfaces (Melotto et al., 2006). Similarly, a series of epidemiological studies of P. syringae as a bean epiphyte and pathogen by Hirano and Upper laid the foundation for elegant experiments showing that type III secreted effectors and the Gac regulon are each critical for epiphytic fitness in the field; these important phenotypes were invisible in the controlled environment of a growth chamber (Upper and Hirano, 1996; Hirano et al., 1997, 1999). Seedlings of the apple (Malus domestica) rootstock Budagovsky 9 appear to be susceptible to fireblight, but the mature woody trees are disease resistant in the field (Russo et al., 2008). WHAT ARE THEY UP TO BEHIND OUR BACKS? Pathologists have traditionally, and understandably, focused on discovering how bacteria incite disease during the early stages of acute pathogenesis. We know much less about the end stages of bacterial pathogenesis, how bacteria escape from dying plants, and the traits needed to grow or persist in free-living states in soil, drainage ditches, dead plant residues, on farm implements, or up in the sky. Some species must colonize seeds, vectors, or alternate hosts; others form lesions or other structures that foster bacterial spread in the environment. Although most plant pathogenic bacteria do not form spores, they often survive extremes of humidity and persist for years; how? There are fascinating biological questions in these understudied life stages, which can be found in the disease cycle of almost every plant pathogenic bacterium. Figure 1 Figure 1. Open in new tabDownload slide An example of the multiple relevant life stages in the disease cycle of plant pathogenic bacteria: soft rot of vegetables caused by pectinolytic enterobacteria in the genera Erwinia, Pectobacterium, and Dickeya. Reprinted with permission from Plant Pathology (4th edition), 1997, by George Agrios, APS Press, St. Paul. Figure 1. Open in new tabDownload slide An example of the multiple relevant life stages in the disease cycle of plant pathogenic bacteria: soft rot of vegetables caused by pectinolytic enterobacteria in the genera Erwinia, Pectobacterium, and Dickeya. Reprinted with permission from Plant Pathology (4th edition), 1997, by George Agrios, APS Press, St. Paul. provides only one of many illustrative examples. There are significant opportunities for improved disease control if any stage of the disease cycle can be disrupted. LESS-EXAMINED LIFE STAGES ARE NOW MORE ACCESSIBLE Detection methods such as real-time PCR, GFP tags, and immunofluorescence staining microscopy are sensitive enough to study small populations in situ in the rhizosphere, in water, in animals, and on soil particles. We do not know much about associations between plant pathogenic bacteria and native plants, especially the roles of these bacteria in natural ecosystems. The extent of our ignorance is exemplified by the recent discovery that the very well-studied soft-rot bacterium Erwinia chrysanthemi (now Dickeya dadantii) has a secret life as an insect pathogen (Grenier et al., 2006). Accumulating evidence suggests the ubiquity in plants of bacterial endophytes, most of which are currently unculturable (Zinniel et al., 2002). These largely unexplored communities likely affect disease development (Araújo et al., 2002) and endophytes are also potential biocontrol agents or delivery systems for antipathogenic compounds (Kobayashi and Palumbo, 2000). Further, important plant pathogens like Ralstonia solanacearum, Liberibacter asiaticus, X. fastidiosa, and Clavibacter sepidonicum cause long-term latent infections, effectively functioning as endophytes. What biological signals or conditions tip the balance and cause an innocuous endophyte to become a destructive pathogen? Metagenomic community analysis and in situ transcriptional studies can open windows into this previously inaccessible aspect of plant microbiology. The discovery that some plant pathogenic bacteria affect the weather when they are not living on plants (Christner et al., 2008; Morris et al., 2008) further demonstrates the importance of looking beyond the acute pathogenesis stage of the disease cycle and beyond crop hosts, as well as the power of cross-disciplinary collaborations. However, moving out of the one-gene/one-trait, plus-or-minus-assay comfort zone demands transdisciplinary approaches. For example, collaborations with epidemiologists who have expertise in the relevant statistical and modeling tools can reveal the mechanisms of subtle but crucial quantitative traits like competition, survival, and dispersal. GETTING LEVERAGE ON QUANTITATIVE PROBLEMS AND SYSTEMS BIOLOGY Biological interactions are dynamic, with balances sometimes tipping sharply when thresholds in signaling or population levels are reached. Environmental variability (humidity, temperature, drought stress) has significant but largely unknown effects on plant-bacterial interactions. Metagenomics have shown us that rhizospheres and leaf surfaces support complex communities of microbes that are mostly uncultured and undescribed (Riesenfeld et al., 2004). These communities almost certainly affect the behavior of plant pathogens, but most experiments ignore them. However, we now have computational tools and extensive genomic and gene expression data that allow us to model complex traits of interest to both phytobacteriologists and mathematicians. These instruments can add quantitative information for regulatory models; elucidate complex phenomena like the initiation of infection (Sepulchre et al., 2006) or bacterial cell differentiation (Craciun et al., 2006); identify emergent properties in bacteria and host plants (Long et al., 2008); and model the complex microbial communities that are important for both pathogenesis and biocontrol (Gilbert et al., 1993, 1996; Schloss and Handelsman, 2008). Many of the quantitative methods needed to design and analyze these kinds of experiments are familiar to ecologists and statisticians, who thus make excellent collaborators for molecular bacteriologists. THE UNDEREXPLOITED POWER OF THE MICROSCOPE Natural history remains a powerful form of biology, as everyone who is annotating genomes can testify. It is becoming clear that genomics and even gene expression studies cannot deliver the specific insights offered by using microscopy to follow bacterial colonization and pathogenesis in real time. High-quality histopathology is time consuming and technically demanding, but it is highly rewarding to use superior modern instruments to observe specifically labeled cells, structures, or proteins in situ. This approach has been very productive in studies of animal pathogenesis but is underused by plant microbiologists (but see Newman et al., 2003; Meng et al., 2005; Monier and Lindow, 2005; Melotto et al., 2006; Nakaho and Allen, 2009). Microscopy studies often suggest hypothesis-driven experiments using defined mutants, and this combination can produce especially rapid advances, such as the fascinating discovery that X. fastidiosa uses twitching motility to move against the transpirational flow in xylem and colonize below an infection point (Meng et al., 2005). Similarly, microscopy and fluorescent probes were combined with genetics to gauge the distance signal molecules can travel between cells on roots and leaves (Gantner et al., 2006; Dulla and Lindow, 2008). HYPOTHETICALLY SPEAKING Annotation reinforces the conventional wisdom because we can confidently identify only those genes that have been previously studied. The rapidly expanding set of genomes for plant pathogenic bacteria can be combined with powerful bioinformatics tools and a biologist's perspective to generate some fresh hypotheses about the many conserved hypothetical proteins crowding our genome databases. For example, straightforward experiments would be suggested by the discovery that a particular conserved gene of unknown function is present in genomes of all insect-transmitted bacteria, no matter how distantly related, but absent from genomes of closely related species transmitted by other means. Similar analyses can find conserved hypothetical proteins specific to epiphytes, xylem dwellers, bacteria attacking only monocots, etc. An analysis of plant pathogenic Xanthomonas genomes used this idea to identify genes potentially linked to infecting specific tissues (Lu et al., 2008). However, the next steps require a better understanding of the hidden lives of bacterial pathogens than we currently have, highlighting the need for a greater overlap with bioinformatics and natural history. A word of caution as we discuss bioinformatics and annotation: it is important to test the biological function of a putative gene before drawing too many conclusions. It would be unfortunate and ironic if, in our excitement about genomics, we allowed mutant construction and phenotypic testing to become an underexplored research niche. OTHER UNDEREXPLOITED METHODS Microarray-based profiling of pathogen gene expression under various conditions is advancing at a rapid pace, although more expression studies are needed in biologically relevant in planta settings. Gene expression profiling can be coupled with laser capture microdissection or other creative extraction methods so that specific microbial subpopulations (or specific host cells or tissues) can be analyzed with increased sensitivity. In vivo expression technology screens and their offshoots (Osbourn et al., 1987; Rainey and Preston, 2000; Boch et al., 2002; Brown and Allen, 2004) have still not been employed to detect genes that are expressed at key growth stages of many important plant pathogens. Biosensors offer a way to measure specific conditions as experienced by bacteria in planta (Wright and Beattie, 2004). Semirobotic sample processing, ordered gene knockout collections, and heavy isotope labeling are other examples of promising methods. These are only a few of the underused technologies; the broader microbial sciences offer a regular flow of new methods that beg for use in the study of plant pathogens. As noted above, an equally important source of new methods is collaboration with experts from other disciplines. Partnership with the right ecologist, computer scientist, microfluidics specialist, geologist, or others can break open a previously recalcitrant problem. Finally, getting out into the field to see pathogens under natural conditions often suggests fresh experimental methods or questions. MULTIFUNCTIONAL (CROSS-KINGDOM) SIGNALING Many of the same signal molecules are perceived by both plants and microbes. This is not surprising since angiosperms arose about 3 billion years after bacteria, and evolved in the constant presence of microbial signaling. Similar signal molecules are produced by a wide range of bacteria and all bacterial plant pathogens are likely to be exposed to plant signal molecules, yet the roles of these signals on plants and bacteria has only been explored in a handful of pathosystems and even fewer have been placed into signal networks (Brencic and Winans, 2005). In these cases, it is clear that bacteria are integrating signals from both plant and bacterial cells to regulate virulence genes at the transcriptional and posttranscriptional level, although the relative strength and timing of each signal remains obscure. For example, soft-rot pathogens use a combination of auxin, pectin metabolites, acyl-homoserine lactones, and organic acids to regulate pectate lyases and other virulence genes at both transcriptional and posttranscriptional stages and it is possible to interfere with soft-rot pathogenicity by disrupting these signaling cascades (Charkowski, 2009). Similarly, Agrobacterium responds to auxin, g-amino butyric acid, and salicylic acid (Yuan et al., 2008). It is also clear that signaling cascades are modular since closely related bacteria use identical signal molecules and receptors in different ways. Therefore, these cascades need to be examined in multiple species to understand how plants and bacteria are manipulating each other with small molecules. Aspects of small molecule signaling and defense studied in other areas of microbiology have not yet made large impacts on plant pathology. In some cases, signaling properties of classes of molecules, such as flavonoids, have been described in detail by those examining beneficial microbes such as Rhizobium (Gibson et al., 2008), but we have barely scratched the surface in determining if and how the same class of compounds affect signaling in pathogens. As an example, flavonoid glycosides induce SyrB, which is required for synthesis of syringomycin by P. syringae (Mo et al., 1995). Similarly, many plant compounds have been examined for their antimicrobial effects on human pathogens, but their effects on plant pathogens are unknown. For example, 5′-methoxyhydnocarpin, a plant compound that inhibits an ATP-binding cassette transporter, thereby making the plant-produced antibiotic berberine more effective, has been studied for its effect on a human pathogen, but not on plant pathogens (Stermitz et al., 2000). IDENTIFYING HIDDEN PARTNERSHIPS Plants face multiple pathogens and there are hints that some pathogens function best in pairs, but this area has been little explored. An almost completely unexamined example is soft-rot disease caused by Clostridium. Clostridium and Pectobacterium species are routinely found together in decaying vegetables and both can cause disease on their own (Pérombelon et al., 1979; Campos et al., 1982). These pathogens may work together to attack their plant hosts. Although potatoes (Solanum tuberosum) are mostly starch, Pectobacterium curiously cannot degrade starch, while Clostridium efficiently breaks down this polymer. Close relatives of Pectobacterium, such as Klebsiella, can metabolize starch (Holt, 1994), so the inability to use this abundant polymer is not inherent, but perhaps developed under the selection of the Pectobacterium-Clostridium partnership. This partnership, as well the role anaerobes play in the soil and on roots and the effects of low oxygen on bacterial pathogens in general, is little explored. This may be because plant pathologists are reluctant to work with anaerobes. Nevertheless, pathogens face low-oxygen environments in plants, in soil, and in waterways, suggesting that research on this topic could be fruitful. MICROBE-ASSOCIATED MOLECULAR PATTERNS AND EFFECTORS Two areas of phytobacteriology that are currently under intensive study are microbe-associated molecular patterns (MAMPs; also called PAMPs) and type III secretion system-dependent effector proteins. Our primary message is to encourage research beyond these heavily studied topics, but even these topics contain underexplored niches concerning the real-world relevance of MAMPs and effectors (Bent and Mackey, 2007). For example, in the biologically realistic setting of an intact plant infested with a reasonable population of living microorganisms, how much MAMP is present and needs to be present, and in what plant tissues, for effective defenses to be triggered? Are epidermal cells, which are routinely exposed to an extensive microbial flora, less sensitive to MAMPs? Learning how plant cell types differ in their responses would help us determine how MAMP detection systems work along natural routes of bacterial entry and spread, such as the vasculature. This could also move us toward understanding how many host processes need to be inhibited by effectors for any particular pathogen to succeed, and if different pathogen species commonly suppress the same host targets. The role of effectors in gene-for-gene systems is well studied, but are defense-suppressing effectors important for broad host-range pathogens as well? Why are plant resistance genes that work against necrotrophs so rare? It has been suggested that necrotrophs are less dependent on suppression of plant defenses and may even benefit from induction of some defense pathways (Glazebrook, 2005), but this hypothesis remains to be broadly tested. Similar unanswered questions remain about the real-world biology, especially in field settings, of other much-studied virulence factors like toxins, plant growth regulators, and macerating enzymes. Several major problems in the management of bacterial plant diseases could be solved with a better understanding of the underlying biology. A few examples are given below. COPPER AND ANTIBIOTIC RESISTANCE Some core methods for control of bacterial diseases, such as copper or streptomycin sprays, lose their utility because pathogens become resistant, often through acquisition of broad host-range transmissible plasmids from other bacteria. Can anything be done to prevent this? Alternatively, β-lactamase inhibitors like clavulanic acid are used clinically to make amoxicillin work against resistant strains (Payne et al., 1994); can this approach be adapted to agricultural ecosystems? Are there copper resistance protein inhibitors in the soil metagenome? ERADICATION FROM SEEDS, CUTTINGS, AND SEEDLINGS Seed treatments such as hot water treatments are one of the best interventions available to disrupt bacterial diseases, but they are only effective in some pathosystems (Leben and Sleesman, 1981). Why? Is it simply a pragmatic issue of accessing vulnerable bacteria without killing the seeds, or are there more interesting aspects of bacterial biology that underpin resistance to such treatments? What controls the ability of bacteria to colonize certain regions of seed or sanctuaries in other tissues, or the ability of plant seeds to tolerate antibacterial treatments? PATHOGEN DETECTION Research seems to have dwindled on the previous two problems, but ongoing efforts seek improved detection methods for bacteria on seeds and cuttings (Gitaitis and Walcott, 2007). The commercial and legal stakes are high. Much effort has been devoted to finding reliable targets for PCR-based detection, often drawing on genomic sequencing and high-thoughput resequencing of diverse strains. But if PCR, ELISA, selective culturing, and other tests remain insufficiently sensitive, is this due solely to the needle-in-a-haystack sampling challenges? Are there paradigm-shifting methods available from other subdisciplines like bioterrorism prevention that are waiting to be applied to phytobacteriology? We conclude by offering a few specific suggestions to increase exploration of new niches. SHAKE UP THE REVIEW PANELS Understandably, funding agencies often enlist researchers who currently receive funding from that agency to serve on their proposal review panels. However, this practice may reinforce a narrow vision of research excellence. To increase research on underexplored niches, some panel managers have successfully broadened their portfolio by recruiting panelists from outside of their funded community, including researchers from significantly different disciplines as well as recent applicants whose unfunded proposals were regarded as highly creative or novel. TRAIN FOR BREADTH Consciously multidisciplinary training will increase the likelihood that our students and postdoctoral researchers become scientists who think broadly and are eager to work with partners who have a very different perspective or toolset. Professors can encourage this by broadening coverage in their own courses and modeling broad collaboration in their research programs. Students and postdocs can generate breadth through their course selections, their reading, meeting, and seminar choices, and through active pursuit of collaborative research. SUPPORT OUTDOOR SCIENCE Finally, we will not succeed in these underexplored niches if molecular biology lab rats do not work with colleagues who spend time in the field. Scientists with a strong laboratory orientation can benefit enormously from the biological expertise and thoughtful perspectives of field pathologists. Find the time to chat regularly with these colleagues. Moreover, the ongoing loss of extension agents and applied plant pathologists through retirements and funding cuts imperils this entire field of study. Relevant insights from natural and agricultural environments will dry up if we do not give our strongest moral and practical support to scientists with expertise in field biology. ACKNOWLEDGMENTS The authors acknowledge helpful comments from Patricia McManus and support from the University of Wisconsin-Madison College of Agricultural and Life Sciences. 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The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Caitilyn Allen ([email protected]). www.plantphysiol.org/cgi/doi/10.1104/pp.109.140004 © 2009 American Society of Plant Biologists This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Nicaise, Valerie; Roux, Milena; Zipfel, Cyril

doi: 10.1104/pp.109.139709pmid: 19561123

In an environment rich in potentially harmful microbes, plant survival depends on efficient microbe perception and fast defense responses. Contrary to the mammalian immune system composed of cells specialized for defense (e.g. lymphocytes), plant immunity relies on the ability of each cell to recognize pathogens. A first level of microbe recognition is performed by membrane proteins termed pattern recognition receptors (PRRs), which perceive molecular signatures characteristic of a whole class of microbes, termed pathogen-associated (or microbe-associated) molecular patterns (PAMPs; Medzhitov and Janeway, 1997). Perception of PAMPs by PRRs is common to all multicellular organisms and leads to an array of defense responses and redeployment of cellular energy in a fast, efficient, and multiresponse manner, which prevents further pathogen ingress. PAMP recognition leads to a chain of signaling events, broadly referred to as general defense responses in plants. PAMP perception also results in plant systemic acquired resistance (Mishina and Zeier, 2007b). Faced with PAMP-triggered immunity (PTI), successful pathogens evolved secreted effectors targeting key PTI actors to interfere with plant defense. In turn, some plant cultivars have evolved resistance (R) proteins to directly or indirectly detect these effectors (previously termed avirulence or Avr proteins) according to the gene-to-gene theory and leading to effector-triggered immunity (ETI), which is often accompanied by the hypersensitive response, a form of programmed cell death. This model illustrates the dynamic coevolution between plants and pathogens (Chisholm et al., 2006; Jones and Dangl, 2006). Gaining knowledge related to recognition and signaling in PTI constitutes a challenge in plant pathology research, as many of the underlying molecular mechanisms remain largely unknown. In this review, we summarize our knowledge of PTI with a special focus on recognition of bacteria. PAMP RECOGNITION BY PRRS: THE PLANT SENTINELS ARE AT THE PLASMA MEMBRANE PAMPs are molecular components highly conserved within a class of microbes, where they carry out an essential function for fitness or survival (Medzhitov and Janeway, 1997). Plants recognize a wide range of bacterial PAMPs, most of which are derived from structural components of the bacterial cell. Although the number of identified bacterial PAMPs recognized by plants is increasing constantly, very few plant PRRs have been discovered. Flagellin and FLS2 The protein flagellin, the building block of the motility organ flagellum, is recognized by most plants, indicating that detection of flagellin is evolutionarily ancient (Boller and Felix, 2009). Synthetic peptides corresponding to a highly conserved part of the flagellin N terminus act as potent elicitors at subnanomolar concentrations (Felix et al., 1999), although a certain degree of recognition specificity exists. The peptide flg22 (corresponding to 22 amino acids localized in the conserved region) elicits responses in most plant species and is as active as the full-length flagellin. Interestingly, the flg22 region is required for bacterial virulence and motility, consistent with the fact that PAMP mutation has a fitness cost for microbes (Naito et al., 2008). Rice (Oryza sativa) was reported to be insensitive to flg22 (Felix et al., 1999). However, recent results showed that flg22 is recognized by rice but that this response is weaker than with full-length flagellin (Takai et al., 2008). A shorter version of the flg22 peptide, called flg15, acts as an antagonist in Arabidopsis (Arabidopsis thaliana), while it is fully active in tomato (Solanum lycopersicum; Meindl et al., 2000; Bauer et al., 2001; Robatzek et al., 2007). The recent finding that flg22, as well as flagellin, induces the hypersensitive response (Naito et al., 2008) revoked the apparent dogma that PAMPs generally do not induce this response. In addition, flagellins derived from nonadapted bacteria but having identical protein sequences differentially induce strong defense responses in nonhost plants, suggesting that other domains and/or posttranslational modifications of flagellin are also recognized (Taguchi et al., 2003, 2006; Takeuchi et al., 2003, 2007). In rare examples, some virulent phytopathogenic bacteria are able to mask recognition of a PAMP (e.g. flagellin) by mutating residues within the recognized epitope (Felix et al., 1999; Pfund et al., 2004; Sun et al., 2006). This reflects a virulence strategy evolved by successful pathogens complementary to effector secretion. In other words, while flagellins from nearly all bacteria are recognized by plants, only a few from plant pathogenic bacteria are not. The Leu-rich repeat receptor kinase (LRR-RK) FLS2 is the PRR for flagellin. It belongs to subfamily XII of LRR-RK and consists of an extracellular domain with 28 LRR motifs, a transmembrane domain, and a cytoplasmic Ser/Thr kinase domain (Gomez-Gomez and Boller, 2000). First identified in Arabidopsis, FLS2 orthologs have since been cloned in tomato, Nicotiana benthamiana, and rice (Hann and Rathjen, 2007; Robatzek et al., 2007; Takai et al., 2008). Although FLS2 directly binds flg22 and is responsible for recognition specificity, the flg22 binding site is still unknown (Chinchilla et al., 2006; Robatzek et al., 2007). Nevertheless, the AtFLS2 LRR domains 9 to 15 contribute significantly to flg22 binding (Dunning et al., 2007). In Arabidopsis, pretreatment with flg22 restricts the growth of the pathogenic bacterium Pseudomonas syringae pv tomato DC3000 (Pto DC3000), and fls2 mutants are more susceptible to this bacterium (Zipfel et al., 2004). In addition, lack of flagellin recognition allows more growth of the nonadapted bacteria P. syringae pv phaseolicola and P. syringae pv tabaci (Li et al., 2005; de Torres et al., 2006). These data demonstrate the importance of flagellin perception in innate immunity. EF-Tu and EFR The elongation factor Tu (EF-Tu) acts as a very potent bacterial PAMP in Arabidopsis and other members of the Brassicaceae family (Kunze et al., 2004). EF-Tu is one of the most abundant and conserved bacterial proteins. Similar to flg22 and flagellin, a synthetic peptide corresponding to the N-acetylated first 18 amino acids of EF-Tu, elf18, is sufficient for recognition, while the shorter peptide elf12 is an agonist in Arabidopsis (Kunze et al., 2004). Although EF-Tu is mostly intracellular, its release from lysis of dying bacteria during plant colonization should be sufficient to trigger its subnanomolar recognition. Moreover, EF-Tu is clearly present in the secretome of several bacteria and serves as an adhesion factor at the bacterial surface, in addition to its primary role in translation (Zipfel et al., 2006, and refs. therein). The LRR-RK EFR is the PRR for EF-Tu and, like FLS2, belongs to subfamily XII (Zipfel et al., 2006). Similar to FLS2, the exact elf18 binding site is still unknown. EFR structure is highly similar to FLS2, with a 21-LRR extracellular domain, a transmembrane domain, and a cytoplasmic Ser/Thr kinase domain. As for FLS2, EFR autophosphorylation has been reported (Xiang et al., 2008), suggesting that both FLS2 and EFR carry active kinase domains. EF-Tu responsiveness was found only in Brassicaceae species (Kunze et al., 2004), suggesting that EFR is an innovation of this family. Nevertheless, genes with high similarities with EFR exist in Arabidopsis and other plants; their function as PRRs needs to be determined. Arabidopsis plants lacking EFR are more amenable to transformation by Agrobacterium tumefaciens, revealing that plant transformation is normally restricted by plant defenses (Zipfel et al., 2006). In addition, efr mutant plants are more susceptible to colonization by weakly virulent mutant strains of Pto DC3000 (C. Zipfel, unpublished data). Interestingly, transient heterologous expression of AtEFR in N. benthamiana, a plant that normally lacks elf18 responsiveness, restores elf18 binding and responses (Zipfel et al., 2006), demonstrating that downstream signaling components are conserved between Brassicaceae and Solanaceae. AvrXa21 and Xa21 In rice, the LRR-RK Xa21 confers resistance to Xanthomonas oryzae pv oryzae strains carrying the Avr gene AvrXa21 (Song et al., 1995). Strikingly, Xa21 also belongs to subfamily XII of the LRR-RKs and is highly similar to EFR. Similar to FLS2 and EFR, Xa21 possesses a non-RD kinase, whose presence has been proposed to be correlated with a role in innate immunity across kingdoms (Dardick and Ronald, 2006). Although the identity of AvrXa21 was until now unknown, recent work identified AvrXa21 as a type I secreted sulfated peptide (da Silva et al., 2004; Lee et al., 2006, 2008; P. Ronald, personal communication). AvrXa21 is conserved among all Xanthomonas strains sequenced (P. Ronald, personal communication), suggesting that AvrXa21/Xa21 constitutes a PAMP/PRR perception system. Orphan PAMPs The bacterial cell wall is an important source of PAMPs. Peptidoglycans (PGNs) are polymers of alternating GlcNAc and N-acetyl-muramic acid residues in β-1-4 linkage that are cross-linked by short peptides. They constitute the major structural components of the gram-positive bacterial cell wall, while their presence is restricted to the periplasmic space in gram-negative bacteria. PGNs from both gram-positive and gram-negative bacteria are recognized by Arabidopsis (Gust et al., 2007; Erbs et al., 2008). However, while perception of gram-positive PGNs mostly depends on their sugar backbones (Gust et al., 2007), muropeptides derived from gram-negative PGNs are more potent elicitors than intact PGNs (Erbs et al., 2008). Lipopolysaccharide (LPS) is the principal component of the outer membrane of gram-negative bacteria and acts as a PAMP in dicots and monocots (Newman et al., 2007). It contains a long-chain polysaccharide, called O-antigen, which is highly variable with respect to composition, length, and the branching of its carbohydrate subunits. In contrast, the oligosaccharide core and the lipid A, which form the sheet of the membrane, are highly conserved in different bacteria. The lipid A part of LPS is as effective as intact LPS in inducing a defense response in Arabidopsis (Zeidler et al., 2004). Interestingly, the phosphorylation and acylation of the lipid A moiety seem to influence LPS elicitor activity (Silipo et al., 2008). In addition, synthetic oligorhamnans, which are common components of the otherwise highly variable O-chain in LPS, can trigger defense responses in Arabidopsis (Bedini et al., 2005). Intriguingly, in addition to activating defenses, LPS and other exopolysaccharides can suppress defense responses, for example by chelating calcium ions (Newman et al., 2007; Tellstrom et al., 2007; Aslam et al., 2008). Recently, rhamnolipids derived from Pseudomonas aeruginosa were identified as PAMPs recognized by grapevine (Vitis vinifera; Varnier et al., 2009). Cyclic lipopeptides derived from multiple strains of Bacillus subtilis have also been demonstrated to stimulate defense responses in tobacco (Nicotiana tabacum; Jourdan et al., 2009). Similar to flg22 and elf18, the highly conserved RNA-binding motif RNP-1 of bacterial cold shock proteins (CSPs) acts as a PAMP in Solanaceae via the recognition of the 22-amino acid core of RNP-1 (CSP22; Felix and Boller, 2003). Other proteinaceous PAMPs perceived by plants include the superoxide dismutase SodM (Watt et al., 2006), harpins (Engelhardt et al., 2009), and Nep1 (for necrosis- and ethylene-inducing peptide 1)-like proteins (Qutob et al., 2006). The bacterial siderophore pseudobactin is also a potential PAMP perceived by Arabidopsis (Meziane et al., 2005). Microbial nucleic acids are classical PAMPs recognized in mammals (Kawai and Akira, 2009). Very recently, bacterial nonmethylated CpG DNA was also shown to be recognized as a PAMP by Arabidopsis (Yakushiji et al., 2009). The PRRs for all of these PAMPs are still unknown. The LysM motif can bind PGN (Buist et al., 2008) and is present in several receptor kinases and transmembrane proteins in plants (Zhang et al., 2007b), suggesting that they might function as PRRs for carbohydrate PAMPs. Interestingly, the legume Nod factor receptors involved in the establishment of rhizobial nitrogen-fixing symbiosis carry LysM motifs (Radutoiu et al., 2007). In addition, the rice LysM-containing transmembrane protein CeBiP directly binds the fungal PAMP chitin (Kaku et al., 2006), while the LysM-RK CERK1 is required for chitin responses in Arabidopsis (Miya et al., 2007; Wan et al., 2008). PGN, Nod factors, and chitin all contain GlcNAc moieties. Unexpectedly, CERK1 was recently shown to be also involved in bacterial recognition, as cerk1 Arabidopsis mutants are more susceptible to Pto DC3000 (Gimenez-Ibanez et al., 2009). cerk1 mutants, however, were not impaired in their responsiveness to flg22, elf18, LPS, or PGN (Gimenez-Ibanez et al., 2009; J. Rathjen and S. Gimenez-Ibanez, personal communication), suggesting that CERK1 is involved in the recognition of yet unknown bacterial PAMP(s). Whether CERK1 is a dual-specificity PRR capable of binding different PAMPs or acts as a downstream signaling adaptor needs to be determined. IMMEDIATE EVENTS AT THE PLASMA MEMBRANE BAK1: A Signaling Facilitator? Formation of receptor complexes linking extracellular perception to intracellular signal transduction is a common theme in plant and animal signaling (Aker and de Vries, 2008). The LRR-RK BRI1 is the receptor for brassinosteroids (BRs), a class of phytohormones that control many aspects of growth and development (Vert et al., 2005). BRI1 forms a complex with the LRR-RK SERK1, SERK3/BAK1, and SERK4/BKK1 to ensure full BR signaling (Aker and de Vries, 2008). Unexpectedly, it has recently been demonstrated that FLS2 and BAK1 interact rapidly (less than 2 min) in a ligand-dependent manner (Chinchilla et al., 2007; Heese et al., 2007). The rapid FLS2-BAK1 association suggests that BAK1 may exist in a preformed complex at the membrane, weakly associated with FLS2. Conformational changes induced by flg22 binding would result in tighter interactions, possibly due to mutual transphosphorylation of the kinase domains. Although BAK1 is not required for flg22 binding, early and late flg22 responses are strongly impaired in bak1 mutants (Chinchilla et al., 2007; Heese et al., 2007). bak1 mutants also show reduced early elf18-triggered responses (Chinchilla et al., 2007), although a direct interaction between EFR and BAK1 has not yet been reported. In Arabidopsis and N. benthamiana, BAK1 is also required for responses triggered by the orphan PAMPs CSP22, HrpZ, PGN, and LPS (Heese et al., 2007; Shan et al., 2008). Together, these data demonstrate that BAK1 is a positive PTI regulator that acts downstream of several PRRs, independently of BR. The fact that BAK1 is involved in BR and PTI responses, as well as in cell death control (He et al., 2007; Kemmerling et al., 2007), suggests that BAK1 is a general signaling adaptor for RKs. Interestingly, a recent report showed that BRI1-BAK1 interaction leads to the transphosphorylation of their respective kinase domains and the subsequent enhancement of BRI1 signaling output (Wang et al., 2008b), suggesting that BAK1 is a signal “amplifier” rather than an integral component of downstream signaling pathways. It would be interesting to test if this model also applies for FLS2-BAK1 and whether CERK1 plays a similar role as BAK1 in BAK1-independent PTI responses. Bacterial Virulence Effectors Directly Target PRRs and Their Associated Proteins To infect plants, pathogens need to defeat PTI. Several recent studies clearly showed that one strategy to do so is to directly target PRRs and their associated proteins by virulence effectors. The model bacterium Pto DC3000 secretes more than 30 effectors (Jones and Dangl, 2006). Among them, AvrPto is a small triple helix protein that could act as a kinase inhibitor (Xing et al., 2007), and AvrPtoB contains an E3 ligase domain (Janjusevic et al., 2006). In resistant tomato plants, AvrPto and AvrPtoB are directly recognized by the cytoplasmic protein kinase Pto in plants carrying the nucleotide-binding site-LRR gene Prf, leading to ETI responses (Mucyn et al., 2006). Interestingly, ETI triggered by Pto and Prf in resistant tomato plants results from the inhibition of the AvrPtoB E3 ligase activity by Pto (Ntoukakis et al., 2009). In susceptible tomato and Arabidopsis plants, AvrPto contributes to virulence and inhibits all responses induced by several PAMPs (He et al., 2006; Hann and Rathjen, 2007; Xiao et al., 2007), suggesting that AvrPto might target very early PTI events. Indeed, based on homology between Pto, FLS2, and EFR kinase domains, Zhou and colleagues have shown that AvrPto interacts in vivo with FLS2 and EFR and inhibits their autophosphorylation in a dose-dependent manner (Xiang et al., 2008). In parallel, FLS2 and CERK1 have recently been identified as targets of AvrPtoB, leading to their degradation (Gohre et al., 2008; Shan et al., 2008; Gimenez-Ibanez et al., 2009). In addition, AvrPto and AvrPtoB are also able to target BAK1, thereby preventing the formation of PRR/BAK1 complexes (Shan et al., 2008). Together, these discoveries illustrate an effective strategy employed by pathogens to suppress PTI by directly targeting PRRs. PRR Endocytosis Analysis of FLS2-GFP fate using confocal microscopy revealed that FLS2-GFP is rapidly internalized into intracellular vesicles after flg22 treatment (Robatzek et al., 2006). This finding parallels the fact that other plant RKs are endocytosed (Shah et al., 2002; Russinova et al., 2004; Gifford et al., 2005; Geldner et al., 2007). FLS2 endocytosis depends on receptor activation, its PEST motif present in the cytoplasmic domain, the proteasome, cytoskeleton functions, and BAK1 (Robatzek et al., 2006; Chinchilla et al., 2007). Whether FLS2 internalization regulates its recycling, degradation, or signaling is still unclear. Negative Regulation by Phosphatases and E3 Ubiquitin Ligases Phosphorylation/dephosphorylation events are efficient regulatory mechanisms for signaling pathways involving kinases. The kinase-associated protein phosphatase (KAPP) is a member of the protein phosphatase 2C (PP2C) family. KAPP binds the kinase domain of FLS2 in yeast two-hybrid experiments (Gomez-Gomez et al., 2001), and transgenic Arabidopsis plants overexpressing KAPP are affected in flg22 binding and induced responses. Therefore, KAPP is a negative regulator of FLS2 (Gomez-Gomez et al., 2001). The fact that KAPP interacts with many plant RKs through their phosphorylated kinase domains (Chevalier et al., 2009) suggests that it is a general regulator of RKs. Another PP2C, the rice XB15 (for Xa21-binding protein 15), interacts both in vitro and in vivo with the kinase domain of Xa21 (Park et al., 2008). The dephosphorylation of Xa21 by XB15 inactivates the receptor and compromises Xa21-mediated bacterial resistance. In mammals, E3 ligases are known to act as mediators of immune responses, via the degradation of negative regulators of PRR pathways as well as the activation of mitogen-activated protein kinase (MAPKs) and transcription factors (Liu et al., 2005; Moynagh, 2009). Recently, a triplet of plant U-box E3 ligases (PUBs), PUB22, PUB23, and PUB24, was shown to act as a negative regulator of PTI in response to various PAMPs in Arabidopsis (Trujillo et al., 2008), probably through the degradation of positive regulators or conceivably by interaction with PRRs. Conversely, a reduced level of the E3 ubiquitin ligase XB3, a substrate of the Xa21 kinase, correlates with a reduction of Xa21 accumulation and compromises Xa21-mediated resistance (Wang et al., 2006). Heterotrimeric G Proteins In mammals, the heterotrimeric G protein complexes (composed of three subunits, α, β, and γ) are associated to the plasma membrane and interact with specific receptors to initiate intracellular signaling cascades (Luttrell, 2006). Heterotrimeric G proteins are involved in many diverse physiological processes in plants (Temple and Jones, 2007; Chen, 2008; Gao et al., 2008b; Oki et al., 2009). The gene AGB1, encoding the β-subunit in Arabidopsis, is highly induced after flg22 and elf18 treatment (Zipfel et al., 2006). Based on this observation, Ishikawa (2009) investigated whether AGB1 is involved in FLS2- and EFR-mediated responses and showed that agb1 mutants are impaired in the oxidative burst and seedling growth inhibition triggered by flg22 and elf18. Similarly, XLG2 (for extra-large protein G 2) gene expression is induced after bacterial infection, and xlg2 mutant plants are more susceptible to P. syringae (Zhu et al., 2009). How heterotrimeric G proteins regulate PTI responses is still unclear and requires further investigation. DOWNSTREAM SIGNALING: PLANTS COUNTERATTACK Ion Fluxes The first easily detectable physiological response to PAMPs in plant cell cultures is the alkalinization of the growth medium. Occurring 0.5 to 2 min after elicitation, this event relies on drastic changes in ion fluxes across the plasma membrane (Nurnberger et al., 2004; Garcia-Brugger et al., 2006). Fluxes of H+, K+, Cl−, and Ca2+ have been observed after PAMP treatment (Jabs et al., 1997; Pugin et al., 1997; Garcia-Brugger et al., 2006). Elevation of cytoplasmic calcium is a critical step in plant innate immunity and is mediated by an increase in Ca2+ influx (Ma and Berkowitz, 2007). The cyclic nucleotide-gated channel 2 mediates this influx after LPS perception (Ali et al., 2007). Changes in [Ca2+]cyt are perceived by calcium-binding proteins such as calmodulin, calcium-dependent protein kinases, and calcineurin B-like proteins (Reddy and Reddy, 2004). Some of these have a demonstrated role in plant defense, particularly in the control of reactive oxygen species (ROS) and salicylic acid (SA) production (Chiasson et al., 2005; Takabatake et al., 2007; Xing et al., 2007; Galon et al., 2008; Du et al., 2009; Wang et al., 2009). Interestingly, the Arabidopsis S-locus RK CBRLK1 interacts with calmodulin and acts as a negative regulator of plant defense against bacteria (Kim et al., 2009a, 2009b). The importance of Ca2+ in defense signal transduction is further supported by the demonstration that Ca2+ chelation by bacterial exopolysaccharides is a virulence strategy used by pathogens to overcome PTI (Aslam et al., 2008). Oxidative Burst PAMPs induce rapid and transient production of ROS in an oxidative burst within a few minutes after treatment. ROS are highly toxic intermediates corresponding to reduced oxygen forms, such as the superoxide anion and hydrogen peroxide. ROS produced during pathogen challenge are largely derived from the activity of membrane-localized NADPH oxidases (respiratory burst oxidase homologs [Rboh]; Torres et al., 2006), with AtRbohD being the most important for PAMP-triggered oxidative burst (Meszaros et al., 2006; Nuhse et al., 2007; Zhang et al., 2007a). The relative position of oxidative burst in the sequence of signaling events during PTI is still unclear. In Arabidopsis, RbohD-dependent ROS production seems to be downstream or independent of MAPK activation (Zhang et al., 2007a). From MAPKs to Defense Gene Expression Protein phosphorylation occurs in diverse cellular processes as a means of controlling protein activity. Signaling via the MAPK network relies on directional and sequential phosphorylation events between three elements, MAPK kinase kinases, MAPK kinases, and MAPKs. MAPKs are involved in various processes in eukaryote cells, including plant defense (Colcombet and Hirt, 2008). In Arabidopsis, a complete MAPK cascade including MEKK1-MKK4/5-MPK3/6 was initially proposed to be involved in PTI downstream of FLS2 (Asai et al., 2002). More recent work showed that MEKK1 does not regulate flg22-activated MPK3/6 but rather activates MPK4, known as a negative regulator of defense (Ichimura et al., 2006; Nakagami et al., 2006; Su et al., 2007; Suarez-Rodriguez et al., 2007; Gao et al., 2008a). At the MAPK kinase level, flg22-induced activation of MPK3/4/6 is dependent on MKK1, while MPK3 and MPK6 are also activated by MKK4 (Meszaros et al., 2006). Furthermore, MKK1 and MKK2 seem to act redundantly to control MPK4 (Gao et al., 2008a; Qiu et al., 2008). Thus, FLS2 activates two simultaneous MAPK cascades: one consists of an unknown MEKK-MKK4/5-MPK3/6 and acts positively on PTI, while the other, consisting of MEKK1-MKK1/2-MPK4, acts negatively on PTI. During PTI, MAPK cascade activation leads to the activation of WRKY-type transcription factors, key regulators of plant defenses (Eulgem and Somssich, 2007; for review, see Pandey and Somssich, 2009). For example, the positive regulators WRKY22 and WRKY29 act downstream of the MPK3/6 cascade (Asai et al., 2002), while MPK4 directly regulates gene expression by interaction with WRKY25 and WRKY33 and the MPK4-interacting protein MKS1 (Andreasson et al., 2005; Zheng et al., 2007; Qiu et al., 2008). Interestingly, MPK4 exists constitutively in nuclear complex with MKS1 and WRKY33. Pathogen challenge leads to MPK4 activation and MKS1 phosphorylation, leading to the release of MKS1 and WRKY33 and the activation of gene expression (Qiu et al., 2008). Although PAMPs trigger the simultaneous activation of positive (MPK3/6) and negative (MPK4) regulators of defense gene expression, these antagonistic pathways are regulated by the same PP2C phosphatase, AP2C1 (Schweighofer et al., 2007). It may seem counterintuitive, but in practice this may provide a sensitive mechanism for the control of defense responses by maintaining a careful balance of positive and negative regulators during signaling. Interestingly, MPK3 and MPK6 are directly targeted by the bacterial virulence effector HopAI1 via its phosphothreonine lyase activity (Zhang et al., 2007a). MAPKs are involved in many different aspects of plant physiology, including stomata patterning (Wang et al., 2007b), ovule and anther development (Bush and Krysan, 2007; Wang et al., 2008a), and leaf senescence (Zhou et al., 2009). This poses the question of signal specificity, which may occur through the action of as yet unidentified parallel signaling pathways or through time- and location-specific expression (Colcombet and Hirt, 2008). The latter example was recently nicely illustrated during embryonic patterning, where an upstream regulator of the MAPK3 YODA, the receptor-like cytoplasmic kinase SSP, is specifically transcribed in the mature pollen but is only translated in the ovule, where SSP protein transiently accumulates (Bayer et al., 2009). Callose Deposition The accumulation of callose, a plant β-1,3-glucan polymer synthesized between the cell wall and the plasma membrane, is a classical marker of PTI responses after treatment with PAMPs or noninfectious pathogens (Bestwick et al., 1995; Brown et al., 1998; Gomez-Gomez et al., 1999). The callose synthase GSL5/PMR4 is responsible for callose synthesis in response to PAMPs and fungal pathogens in Arabidopsis (Jacobs et al., 2003; Nishimura et al., 2003; Kim et al., 2005). pmr4 mutant plants allow 20-fold more growth than wild-type plants of the type 3 secretion system (TTSS)-deficient strain Pto DC3000 hrcC − (Kim et al., 2005), while the double mutant pmr4 pad4 (which also reduces SA levels) allows some growth of the nonadapted bacterium P. syringae pv phaseolicola in comparison with the respective single mutants (Ham et al., 2007). This indicates that PMR4-dependent callose deposition contributes to antibacterial immunity. Although the order of events in PTI is still not clear, callose deposition may be downstream of ROS production, as AtrbohD mutants exhibit fewer callose deposits after flg22 treatment (Zhang et al., 2007a). Interestingly, callose deposition was recently shown to depend on PAMP-induced glucosinolates (Clay et al., 2009), components that are linked to antimicrobial immunity (Brader et al., 2001; Mishina and Zeier, 2007a; Bednarek et al., 2009; Clay et al., 2009). Hormone Action SA, jasmonic acid (JA), and ethylene (ET) function as classical defense hormones (Bari and Jones, 2009). Bacterial PAMPs, such as flg22, induce the production of SA (Mishina and Zeier, 2007a; Tsuda et al., 2008) that is required for both local and systemic acquired resistances (Durrant and Dong, 2004), consistent with the induction of systemic acquired resistance by flg22 and LPS (Mishina and Zeier, 2007b). Moreover, bacterial PAMPs induce the production of ET (Felix et al., 1999). Interestingly, the Arabidopsis enzyme 1-aminocyclopropane-1-carboxylate synthase 6, involved in ET biosynthesis, is a substrate of PAMP-activated MPK6 (Liu and Zhang, 2004; Joo et al., 2008). Despite the flg22-triggered production of SA and ET, local resistance induced by flg22 does not strictly depend on SA, ET, or JA pathways (Zipfel et al., 2004; Ferrari et al., 2007; Tsuda et al., 2008). However, disruption of key SA signaling components affects the expression of a subset of PAMP-regulated genes (Tsuda et al., 2008). Flg22 up-regulates the expression of the Arabidopsis microRNA miRNA393, which reduces auxin receptor levels by targeting TIR1-like proteins (Navarro et al., 2006), and SA antagonizes auxin signaling by stabilizing auxin response repressors (Wang et al., 2007a). This suggests that PTI and auxin signaling pathways are antagonistic. Consistently, the phytopathogenic bacteria Xanthomonas campestris pv campestris and Pto DC3000 increase plant auxin levels (O'Donnell et al., 2003), potentially by up-regulating the expression of auxin biosynthetic genes (Schmelz et al., 2003). Recent excellent reviews summarize the role of phytohormones in plant disease resistance in more detail (Spoel and Dong, 2008; Bari and Jones, 2009). Stomatal Closure Gaseous exchange and water transpiration influenced by environmental conditions are controlled by pores present in the epidermis of aerial plant organs, called stomata. During plant-pathogen interactions, stomata constitute one entry point for bacteria, which need to reach apoplastic spaces to multiply and cause disease. PAMP treatments induce stomatal closure (Lee et al., 1999; Melotto et al., 2006) in a manner dependent on abscisic acid, SA, K+ fluxes, and heterotrimeric G proteins (Melotto et al., 2006; Zhang et al., 2008). Consistent with a role of stomatal closure to limit bacterial infection, the phytotoxin coronatine, a JA-Ile mimic secreted by the pathogenic bacterium Pto DC3000, reverts PAMP-induced stomatal closure (Melotto et al., 2006). Intriguingly, the Gly-rich binding protein 7, which plays a role in stomatal closure in response to abiotic stress (Kim et al., 2008), has been identified as a positive regulator of PTI against bacteria and is a target of the Pto DC3000 effector HopU1 (Fu et al., 2007). Gene Silencing Flg22 treatment leads to the rapid down-regulation of several primary auxin response genes (Navarro et al., 2004, 2006; Zipfel et al., 2004). This initial observation was later linked to the flg22-induced accumulation of the conserved microRNA miRNA393 that targets the auxin receptor (TIR1) and its close paralogs (Navarro et al., 2006). Constitutive overexpression of miRNA393 drastically restricts Pto DC3000 growth. Therefore, antibacterial immunity involves a rapid down-regulation of auxin responses mediated by RNA silencing. Consistently, Pto DC3000 effectors target the silencing machinery to achieve full virulence, with AvrPto interfering with the processing of miRNA393, while AvrPtoB leads to the degradation of miRNA393 precursors (Navarro et al., 2008). In addition to miRNA393, miRNA167 and miRNA160, which target auxin response factors/receptors to negatively regulate auxin signaling, are also induced after infection with Pto DC3000 TTSS − (Fahlgren et al., 2006). Hence, microRNA-deficient Arabidopsis mutants support the growth of Pto DC3000 TTSS − and the nonadapted bacterium P. syringae pv phaseolicola (Navarro et al., 2008), and Arabidopsis plants lacking Argonaute4, which is involved in RNA-directed DNA methylation, are more susceptible to Pto DC3000 and to the nonadapted strain P. syringae pv tabaci (Agorio and Vera, 2007). Thus, gene silencing appears as an inherent component of antibacterial immunity. PERSPECTIVES Perception of microbes by PRRs represents the first line of plant defense, relies on fast, efficient, and carefully coordinated reactions, and plays a major role in disease resistance. This is now clearly illustrated by the recent finding that bacterial virulence effectors directly target PRRs and downstream components to cause disease. However, only a few plant PRRs have been identified so far, and our knowledge of the molecular mechanisms underlying PTI is still limited. Therefore, we need to identify more bacterial PAMPs and their corresponding PRRs, and not only from the classical models Pto DC3000 and Arabidopsis. Crystallographic studies of PAMP/PRR complexes are required to define PAMP-binding sites and to understand receptor activation. From the numerous signaling outputs occurring after PAMP perception, the identity of the molecular players and the exact sequence of signaling events need to be deciphered. 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Nature 428 : 764 – 767 Crossref Search ADS PubMed Author notes 1 This work was supported by the Gatsby Charitable Foundation, the Biotechnology and Biological Sciences Research Council, European Research Area Network Plant Genomics, the Royal Society, and the Two Blades Foundation. 2 These authors contributed equally to the article. * Corresponding author; e-mail [email protected]. The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Cyril Zipfel ([email protected]). www.plantphysiol.org/cgi/doi/10.1104/pp.109.139709 © 2009 American Society of Plant Biologists This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Pandey, Shree P.; Somssich, Imre E.

doi: 10.1104/pp.109.138990pmid: 19420325

Plants constantly face a plethora of abiotic and biotic stresses in their natural habitat. Adapting to such changes requires a great degree of phenotypic plasticity that is mainly determined by the plant's genome. We currently do not know how plants are able to integrate the multitude of partly synergistic/partly antagonistic environmental signals that enable them to respond properly under any given condition. What has become apparent, however, is that plants are capable of extensive reprogramming of their transcriptome in a highly dynamic and temporal manner. This regulation in response, leading to adaptive plasticity of plants in highly variable environments, is mainly achieved by enforcement of a network of various transcription factors (TFs). WRKY TFs are a large family of regulatory proteins forming such a network (Eulgem and Somssich, 2007). They are involved in various plant processes but most notably in coping with diverse biotic and abiotic stresses. In this update, we will restrict our attention to the role of WRKY TFs in plant immunity. THE WRKY FACTORS The WRKY TF superfamily consists of 74 and 109 members in Arabidopsis (Arabidopsis thaliana) and rice (Oryza sativa), respectively (Eulgem and Somssich, 2007; Ross et al., 2007). Members of this family contain at least one conserved DNA-binding region, designated the WRKY domain, comprising the highly conserved WRKYGQK peptide sequence and a zinc finger motif (CX4–7CX22–23HXH/C). This domain generally binds to the DNA element termed the W box (C/TTGACT/C), although alternative binding sites have been identified (Sun et al., 2003; Cai et al., 2008; Ciolkowski et al., 2008; van Verk et al., 2008). WRKY family members are divided into three groups based on the number of WRKY domains and certain features of the zinc finger-like motifs (Eulgem et al., 2000). The NMR solution structure revealed that the C-terminal WRKY domain of Arabidopsis WRKY4 consists of a four-stranded β-sheet, with a zinc-binding pocket formed by the conserved Cys/His residues located at one end of the β-sheet, and the WRKYGQK residues, corresponding to the most N-terminal β-strand (strand β-1), kinked in the middle of the sequence by the Gly residue (Yamasaki et al., 2005). The concave curvature of strand β-1 induced by this kink is predicted to enable this strand to deeply enter the DNA groove and make contact with bases of the W box element. The crystal structure of the extended WRKY domain of Arabidopsis WRKY1 (AtWRKY1-C) revealed that this domain is composed of a globular structure with five β-strands forming an antiparallel β-sheet with an additional novel zinc-binding site at one end (Duan et al., 2007). One should note, however, that no crystal structure information exists of a WRKY domain associated with its DNA-binding site or for a full-length WRKY protein. WRKY factors were generally regarded as being plant specific, but their identification in the protist Giardia lamblia and the slime mold Dictyostelium discoideum imply an earlier origin (Ülker and Somssich, 2004; Pan et al., 2009). They may have evolutionary links with transposons such as Mutator-like elements and could have originated from a BED finger intermediate (an atypical zinc finger DNA-binding domain found both in cellular chromatin boundary element-binding proteins BEAF and DREF and in transposases from animals), although this is controversially debated (Babu et al., 2006; Yamasaki et al., 2008). Duplicated WRKY genes have been maintained in wild and cultivated plant species in the course of selection during domestication and polyploidization (Petitot et al., 2008). They have recently been associated with the viability of interploidy hybrids (Dilkes et al., 2008). Phylogenetic sequence analysis and comparative transcriptomics have revealed that they have retained their functions between monocots and dicots (Mangelsen et al., 2008). The majority of the analyzed WRKY genes respond to pathogen attack and to the endogenous signal molecule salicylic acid (SA; Eulgem and Somssich, 2007). WRKY FACTORS IN DISEASE RESISTANCE NETWORKS Plant innate immunity is composed of two interconnected branches: (1) PTI, or pathogen-associated molecular pattern (PAMP)-triggered immunity, which is initiated by the recognition of molecular signatures of many pathogens and often activates downstream mitogen-activated protein (MAP) kinase cascades and defense genes; and (2) ETI, or effector-triggered immunity, driven by plant disease resistance proteins (major R gene products) that recognize directly or indirectly specific pathogen-derived effectors (Chisholm et al., 2006). PTI and ETI activate local as well as systemic defense responses (called systemic acquired resistance [SAR]), which are modulated by phytohormones, especially jasmonic acid (JA) and SA (Durrant and Dong, 2004; Bostock, 2005). JA-dependent plant defenses are generally activated by necrotrophic pathogens and chewing insects, whereas SA-dependent defenses are often triggered by biotrophic pathogens. JA and SA signaling usually act antagonistically, but synergism between these two phytohormones has also been observed (Mur et al., 2006). These responses to pathogen attack require large-scale transcriptional reprogramming, including those of TF families such as WRKY genes (Eulgem, 2005; Ryu et al., 2006; Naoumkina et al., 2008). WRKY TFs in the Arabidopsis World Loss-of-function and gain-of-function studies in Arabidopsis have been pivotal in demonstrating that WRKY factors act in a complex defense response network as both positive and negative regulators (Eulgem and Somssich, 2007). AtWRKY52 (also designated RRS1) is a novel protein comprising structural features of nucleotide binding-Leu-rich repeat-type R gene products and a WRKY domain that confers wide-ranging resistance toward the bacterial wilt Ralstonia solanacearum (Deslandes et al., 2002). The discovery that AtWRKY52 physically interacts with its cognate bacterial effector PopP2 within the plant cell nucleus (Deslandes et al., 2003) helped to stimulate subsequent research clearly demonstrating the importance of nuclear trafficking for plant immunity (Caplan et al., 2008; Liu and Coaker, 2008). AtWRKY70 acts at a convergence point determining the balance between SA- and JA-dependent defense pathways as well as being required for R gene-mediated resistance (Li et al., 2006; Knoth et al., 2007). The indispensability of AtWRKY70 for JA and SA signaling, however, has recently been questioned (Ren et al., 2008). Similarly, AtWRKY33 functions as a positive regulator of resistance toward the necrotrophic fungi Alternaria brassicicola and Botrytis cinerea (Zheng et al., 2006), and AtWRKY53 and AtWRKY70 both positively modulate SAR (Wang et al., 2006). Moreover, SA biosynthesis and expression of NONEXPRESSOR OF PR1 (NPR1), a key central regulator of SA-dependent defenses and SAR, also appear to be regulated by WRKY TFs (Yu et al., 2001). Two closely related WRKY TFs, AtWRKY3 and AtWRKY4, play a positive role in plant resistance toward necrotrophic pathogens, as Atwrky4, Atwrky3, and Atwrky3 wrky4 mutants showed increasing susceptibility toward the fungus B. cinerea, whereas overexpression of AtWRKY4 enhanced susceptibility toward the biotrophic bacterium Pseudomonas syringae (Lai et al., 2008). Many WRKY TFs act as negative regulators of defense signaling, including AtWRKY7, -11, -17, -18, -23, -25, -27, -38, -40, -41, -48, -53, -58, -60, and -62. Showing functional redundancy, Atwrky7 along with Atwrky11 and Atwrky17 mutants were susceptible to virulent P. syringae (Journot-Catalino et al., 2006; Kim et al., 2006). Similarly, AtWRKY38 and AtWRKY62 also contribute negatively to basal resistance toward this bacterial pathogen (Kim et al., 2008). AtWRKY62 expression is induced by SA and JA in a NPR1-dependent manner. How AtWRKY62 alters JA/SA signaling remains unclear, since one study has shown that loss of AtWRKY62 function resulted in enhanced expression of JA-response genes, whereas AtWRKY62 overexpressor lines inhibited JA-response gene expression (Mao et al., 2007), while in a second study, elevated transcript levels of the SA-response gene PR1 were observed in the Atwrky62 mutant, whereas WRKY62 overexpression led to suppression of PR1 (Kim et al., 2008). AtWRKY48 also negatively influences basal resistance toward virulent P. syringae (Xing et al., 2008). Reduced bacterial growth in Atwrky48 mutants was associated with increased induction of PR1, whereas AtWRKY48 overexpressors showed the opposite phenotypes. AtWRKY58 acts downstream of NPR1, negatively regulating SAR (Wang et al., 2006). Recently, knockdown of AtWRKY23 expression was shown to decrease susceptibility toward the parasitic cyst nematode Heterodera schachtii (Grunewald et al., 2008). Mutation in AtWRKY27 resulted in delayed symptom development against R. solanacearum, possibly by affecting nitric oxide signaling, and vascular trafficking (Mukhtar et al., 2008). The closely related WRKY TFs AtWRKY18, -40, and -60 have partly redundant functions in negatively regulating resistance to P. syringae (Xu et al., 2006). Interestingly, Atwrky18 wrky40 double mutants also displayed enhanced resistance to the powdery mildew pathogen Golovonomyces orontii (Shen et al., 2007). In contrast, Atwrky18 wrky40 and Atwrky18 wrky60 double mutants were more susceptible to B. cinerea (Xu et al., 2006), and AtWRKY18 alone appears also to have positive regulatory functions in SAR (Wang et al., 2006). Dual functionality in defense signaling was also observed for AtWRKY53. While Atwrky53 mutants showed delayed symptom development against R. solanacearum, such plants displayed increased susceptibility toward P. syringae (Murray et al., 2007; Hu et al., 2008). Dual functionality was also suggested for AtWRKY41. Arabidopsis plants overexpressing AtWRKY41 showed enhanced resistance toward virulent Pseudomonas but decreased resistance toward Erwinia carotovora (Higashi et al., 2008). However, Atwrky41 mutants did not display a differential phenotype. Intriguingly, expression of AtWRKY41 is specifically suppressed by a compatible strain of P. syringae in an effector-dependent manner. Finally, overexpression of AtWRKY25 resulted in increased disease symptoms to P. syringae infections, possibly by negatively regulating SA-mediated defense responses. However, Atwrky25 mutants supported normal growth of a virulent P. syringae strain (Zheng et al., 2007). Thus, as in the case for AtWRKY41, the in vivo relevance of such findings remains to be critically assessed. Recent Developments in Rice An increasing number of studies in other plants, particularly in rice, have strongly confirmed the importance of WRKY TFs in plant defense signaling. The rice genome contains more than 100 WRKY genes, often present in duplicated chromosomal regions, suggesting genome duplications as one of the mechanisms for the expansion of this family in this plant species (Ross et al., 2007; Ramamoorthy et al., 2008). The majority of these genes respond to (a)biotic stresses and various phytohormones (Ryu et al., 2006; Ramamoorthy et al., 2008). Individual WRKY members have been associated with pathogen defense, albeit with the caveat that the majority of such studies have employed strong ecotopic overexpressor lines. For example, overexpression of OsWRKY13 enhances resistance to the bacterial blight Xanthomonas oryzae pv oryzae (Xoo) and the fungal blast Magnaportha grisea. It exerts its function by activating SA-biosynthesis and SA-response genes while suppressing JA signaling (Qiu et al., 2007, 2008a). Similarly, OsWRKY53 overexpressor lines are more resistant to M. grisea and may act as a positive regulator of basal defense (Chujo et al., 2007). Expression of OsWRKY03 and OsWRKY71 is strongly induced by pathogen-mimicking stimuli, and these genes function upstream of OsNH1 (the rice ortholog of NPR1) in defense signaling (Liu et al., 2005, 2007). In the case of OsWRKY71, overexpressor lines display enhanced resistance to virulent Xoo (Liu et al., 2007). Ectopic expression of OsWRKY31 resulted in enhanced resistance to fungal blast, altered lateral root formation, and constitutive expression of two early auxin-response genes (Zhang et al., 2008a). Whether these two phenotypes are functionally linked remains to be determined. Moreover, corresponding OsWRKY31 RNA interference lines showed no altered disease phenotype. Enhanced resistance to M. grisea was observed with OsWRKY45 overexpressor lines but not with plants overexpressing OsWRKY19, -62, and -76 (Shimono et al., 2007). In this case, OsWRKY45 knockdown lines decreased resistance to this fungal blast. OsWRKY45 appears to act in SA signaling independent of NH1. Notably, ecotopic expression of OsWRKY45 in Arabidopsis resulted in plants with enhanced resistance to virulent P. syringae, increased PR1 expression, elevated tolerance to salt and drought stress, but decreased sensitivity toward abscisic acid signaling (Qiu and Yu, 2009). OsWRKY89 overexpression seems to positively contribute to resistance against fungal blast and the white-backed plant hopper Sogatella furcifera by regulating the wax content/deposition on the leaf surface. OsWRKY89 knockdown lines showed reduced wax content and increased susceptibility to M. grisea (Wang et al., 2007). Finally, OsWRKY62 was recently shown to be a negative regulator of both PTI and ETI. The rice gene Xa21 confers race-specific resistance to Xoo. Xa21 was shown to bind to OsWRKY62, and overexpression of one splice variant, OsWRKY62-1, compromised basal defense and Xa21-mediated resistance to Xoo and suppressed defense gene activation (Peng et al., 2008). WRKY TFs in Other Plant Species The number of WRKY genes identified in other recently sequenced plant genomes are 66 in papaya (Carica papaya), 104 in poplar (Populus spp.), 68 in sorghum (Sorghum bicolor), and 38 in the moss Physcomitrella patens. Currently, no data exist on the role of these factors in mediating plant immunity. Some isolated studies in other plant species, however, have been reported. Overexpression of grapevine (Vitis vinifera) VvWRKY1 in tobacco (Nicotiana tabacum) rendered plants susceptible toward a variety of fungi (Marchive et al., 2007), whereas ectopic expression of grapevine VvWRKY2 resulted in enhanced resistance to the necrotrophic fungi Alternaria tenuis, B. cinerea, and Pythium (Mzid et al., 2007). Similarly, CaWRKY1 from chili pepper (Capsicum annuum) appears to act as a negative regulator of defense, as virus-induced gene silencing of this gene decreased growth of Xanthomonas, whereas its overexpression resulted in enhanced hypersensitive cell death to P. syringae and Tobacco mosaic virus (Oh et al., 2008). In barley (Hordeum vulgare), MLA confers isolate-specific resistance to the powdery mildew Blumeria graminis. MLA was shown to physically interact in the nucleus with HvWRKY1 and -2, two repressors of PAMP-triggered basal defense, thereby interfering with WRKY repressor functions and leading to resistance against the powdery mildew fungus (Shen et al., 2007). In the native tobacco Nicotiana attenuata, two WRKY genes, NaWRKY3 and -6, were identified that coordinate JA-mediated defense responses to native herbivory. Silencing of NaWRKY3, NaWRKY6, or both rendered plants highly vulnerable to Manduca sexta attack (Skibbe et al., 2008). Finally, elicitor-triggered reprogramming of secondary metabolites in Medicago truncatula seems to involve several WRKY factors: overexpression of four WRKY genes in tobacco demonstrated their regulatory roles in lignin deposition, PR gene expression, and systemic defense responses against Tobacco mosaic virus (Naoumkina et al., 2008). Overall, these findings highlight the importance of WRKY factors in transcriptionally reprogramming plant responses toward different invading pathogens (Supplemental Table S1). While some appear to positively influence the outcome of such plant-pathogen interactions, others actually appear to negatively influence it. This negative influence may be due to active targeting of the WRKY genes/factors, or products under their control, by certain pathogens. Manipulation of WRKY proteins by pathogen effectors may partly explain the existence of redundancy within the WRKY TF family as a reinforcement measure for essential regulatory functions. Coordinated modulation of positive- and negative-acting factors could also enable the proper amplitude and duration of the plant response during pathogen attack. Some key questions that need to be addressed in future WRKY research are as follows. (1) How are the WRKY genes themselves regulated? (2) With which cellular/nuclear components do they interact during defense signaling and during recruitment at specific target gene sites? (3) What are the exact targets of individual WRKY factors within the genome? WHAT REGULATES THE WRKY NETWORK? The last decade of research has clearly revealed that WRKY factors form a complex and highly interconnected regulatory network (Eulgem and Somssich, 2007). Such a network needs to be controlled at several levels. Auto/Cross-Regulation by WRKY Genes The majority of the Arabidopsis WRKY genes are themselves responsive to pathogenic stimuli and many contain numerous W box elements within their promoters (Eulgem and Somssich, 2007). This suggests that several WRKY genes are under direct positive or negative control by WRKY factors via specific feedback mechanisms (auto/cross-regulation). Studies in parsley (Petroselinum crispum) protoplast showed that a specific arrangement of W boxes within the promoter of PcWRKY1 determines its temporal expression upon PAMP treatment (Eulgem et al., 1999). Moreover, chromatin immunoprecipitation (ChIP) analysis confirmed PAMP-dependent in vivo binding of PcWRKY1 to its own promoter as well as to the defense-response gene PcPR10 (Turck et al., 2004). Additional cotransfection experiments have substantiated such a mode of regulation (Eulgem and Somssich, 2007; Lippok et al., 2007). Upon herbivore attack, NaWRKY6 transcript accumulation was shown to be dependent on NaWRKY3 expression (Skibbe et al., 2008). Moreover, physical interaction of related WRKY TFs may also be necessary for their efficient function, as evidenced by homodimer and heterodimer complex formation of Arabidopsis WRKY18, -40, and -60 in response to P. syringae (Xu et al., 2006). Regulation via Other TFs and Proteins Six distinct proteins, including OsWRKY13, were identified in a yeast one-hybrid screen that bind to functionally important cis-regulatory DNA elements within the rice OsWRKY13 promoter (Cai et al., 2008). Similar screens employing the AtWRKY53 promoter led to the identification of a MAP kinase kinase kinase (MEKK1). Interestingly, MEKK1 was also shown to interact with and to phosphorylate AtWRKY53 (Miao et al., 2007). The in vivo relevance of these interactions with respect to plant defense, however, remains to be tested. In Arabidopsis, expression of the key defense regulator NPR1 is controlled by unknown WRKY TFs (Yu et al., 2001). NPR1 does not bind DNA on its own but associates with TGA TFs to modulate SA-dependent genes and SAR (Durrant and Dong, 2004). Expression of at least nine WRKY genes, AtWRKY18, -38, -53, -54, -58, -59, -62, -66, and -70, is dependent on NPR1, suggesting that they may be under TGA factor control (Wang et al., 2006; Mao et al., 2007). In the case of AtWRKY51, ChIP and whole-genome arrays identified its promoter to be targeted by TGA2 in an SA-dependent manner (Thibaud-Nissen et al., 2006). PTI involves tightly regulated MAP kinase signaling cascades. The D motif within several WRKY TFs contains consensus phosphorylation sites for MAP kinases, and several WRKY TFs have been shown to be phosphorylated in vitro (Kim and Zhang, 2004; Menke et al., 2005; Eulgem and Somssich, 2007; Popescu et al., 2009). Recently, the association of MAP Kinase4 (MPK4) with AtWRKY33 and a coupling factor, MKS1, within the plant cell nucleus was demonstrated (Qiu et al., 2008b). Upon virulent P. syringae infection, MPK4 is phosphorylated, thereby releasing MKS1 and WRKY33 and thus allowing recruitment of WRKY33 to target promoters. Chromatin structure can locally and globally regulate gene expression. Interestingly, AtWRKY38 and -62 were found to interact with Histone Deacetylase19 (HDA19), a chromatin-remodeling factor that contributes to global transcriptional repression (Kim et al., 2008). Overexpression of HDA19 enhanced resistance to P. syringae, whereas the hda19 mutant was compromised in resistance. These are the opposite phenotypes obtained from similar studies with AtWRKY38 and -62, revealing yet another level of WRKY network regulation in fine-tuning the plant basal defense response (Kim et al., 2008). The Small RNA-WRKY Interactome Small RNAs (smRNAs) have emerged as a fundamental layer of regulation of gene expression. Plant smRNAs are broadly classified into micro RNAs (miRNAs) and small interfering RNAs (siRNAs). miRNAs are approximately 21 nucleotides and derived from the precursor-stem-loop structures encoded by distinguished miRNA genes (Voinnet, 2009); siRNAs are derived from double-stranded RNAs, in an RNA-directed RNA polymerase-dependent manner, and may be further classified as trans-acting siRNAs, repeat-associated siRNAs, and natural antisense transcript-derived siRNAs. High-throughput sequencing of the smRNA portion of the transcriptome revealed that a multitude of smRNAs accumulate in plants (Lu et al., 2005; Kasschau et al., 2007; Pandey et al., 2008). These 18- to 40-nucleotide-long smRNAs regulate gene expression posttranscriptionally in a process often called RNA interference, RNA silencing, or posttranscriptional gene silencing. The importance of smRNAs in plant processes related to adaptation to (a)biotic stresses is increasingly becoming evident, and the endogenous plant-derived smRNAs probably have broad implications in posttranscriptionally regulating plant responses to pathogen attack (Navarro et al., 2006; Pandey and Baldwin, 2007; Voinnet, 2008). Phytohormone treatments induced the expression of several miRNAs in rice (Liu et al., 2009). Predicted targets for several miRNAs encode WRKY factors (Zhang et al., 2008b; S.P. Pandey and I.T. Baldwin, unpublished data), suggesting smRNA-mediated regulation of WRKY TFs. Conversely, several miRNA gene promoters are highly abundant in W box sequences, implicating WRKY TFs in their activation/repression (Zhou et al., 2008). Further evidence of a WRKY-smRNA interactome comes from our studies on AtWRKY18 and -40 in modulating responses to powdery mildew (S.P. Pandey, M. Roccaro, E. Logemann, and I.E. Somssich, unpublished data). Atwrky18 wrky40 double mutants are resistant to powdery mildew infection and strongly up-regulate the expression of SIMILAR TO RCD ONE5 (SRO5) upon infection, suggesting WRKY-dependent suppression of siRNA-generating loci. SRO5 along with PYRROLINE -5-CARBOXYLATE DEHYDROGENASE (an overlapping gene in the antisense orientation) generate 24- and 21-nucleotide siRNAs, which together are components of a regulatory loop controlling reactive oxygen species production and stress response (Borsani et al., 2005). Similar suppression of the host miRNA machinery by bacterially derived effector proteins has recently been demonstrated in Arabidopsis (Navarro et al., 2008). The current data point toward the existence of a WRKY-smRNA interactome, where on the one hand, pathogen attack triggers the expression of WRKY genes that regulate cellular smRNA populations, and on the other hand, several differentially regulated smRNAs modulate WRKY TF levels by targeting their transcripts (Fig. 1 Figure 1. Open in new tabDownload slide Modeling the WRKY-smRNA interactome during reprogramming of defense responses. During pathogen attack, smRNA-generating loci may be under the control of WRKY TFs; at the same time, WRKY abundance may be regulated by smRNAs. RdR, RNA-directed RNA polymerase. Figure 1. Open in new tabDownload slide Modeling the WRKY-smRNA interactome during reprogramming of defense responses. During pathogen attack, smRNA-generating loci may be under the control of WRKY TFs; at the same time, WRKY abundance may be regulated by smRNAs. RdR, RNA-directed RNA polymerase. ). This model certainly warrants further investigation. PATHOGEN-DEPENDENT IN VIVO WRKY TF TARGETS IN THE POSTGENOMIC ERA As with other large TF families, identification of all in vivo downstream targets of specific WRKY TFs is a highly challenging endeavor. Sequenced genomes reveal a widespread distribution of W box-like elements, but the biological relevance of these potential WRKY-binding sites remains unclear. Earlier target identification was limited to selected candidates on a gene-for-gene basis and rested mostly on ectopic expression of the respective WRKY gene in transient cotransfection assays. Development of the ChIP technology was a major step forward, allowing DNA-protein and protein-protein interactions to be studied under in vivo conditions (Massie and Mills, 2008). ChIP studies in parsley identified two PcWRKY1 target genes activated upon PAMP treatment (Turck et al., 2004). Similarly, PAD3, a gene encoding a key enzyme of camalexin biosynthesis, was detected as a direct target of AtWRKY33 following pathogen infection (Qiu et al., 2008b). Recently, using information derived from whole-genome microarrays followed by ChIP analyses, we identified two key regulators of plant defense as being direct targets of AtWRKY40 during powdery mildew infection (S.P. Pandey, M. Roccaro, E. Logemann, and I.E. Somssich, unpublished data). A major limitation of previous studies was that the number of target genes that could be assayed was restricted. Recent developments expanding the use of ChIP-enriched DNA for hybridization to genomic microarrays (ChIP-chip) or for direct sequencing (ChIP-Seq) using second-generation high-throughput sequencing technology are opening the door to identify WRKY TF binding sites on a global level (Massie and Mills, 2008). Nevertheless, despite such progress, the task remains daunting both technically, starting with the quality of various specific antibodies and proper evaluation of the gigabits of sequencing information obtained, and because such in vivo interactions can be highly dynamic in both temporal and spatial terms. CONCLUSION WRKY TFs are indeed global regulators of host responses following challenge by phytopathogenic organisms. They participate in regulating defense gene expression at various levels, partly by directly modulating immediate downstream target genes, by activating or repressing other TF genes, and by regulating WRKY genes by means of feed-forward and feedback regulatory loops. Moreover, they also appear to interact with key chromatin-remodeling factors, thereby adding another layer of complexity to the WRKY network. WRKY factors can associate with MAP kinases in the nucleus, and MAP kinase cascades constitute key components of plant defense signaling. In yeast, the majority of terminal MAP kinases appear to be within the nucleus, associated with transcriptional complexes at target genes (Pokholok et al., 2006). Hence, one can expect that future studies will reveal additional nuclear functions of such WRKY-MAP kinase associations involving chromatin remodeling at target DNA sites. In addition, the involvement of WRKY TFs in modulating the expression of several miRNAs while at the same time their transcription is possibly partly under smRNA surveillance adds yet another dimension to the regulatory complexity that must be sorted out. Nevertheless, to fully understand regulation, we need to gain access to the full set of proteins associated with WRKY TFs at specific genomic loci. Indeed, promising technological advances combining DNA probes and mass spectrometry, such as proteomics of isolated chromatin segments and stable isotope labeling with amino acids, are starting to demonstrate that identification of TFs and associated proteins in vivo at given promoters may become feasible in the near future (Dèjardin and Kingston, 2009; Mittler et al., 2009). The WRKY transcriptional network may provide the proper balance to respond quickly and efficiently to deter pathogens but at the same time to restrict defense responses that can be detrimental for plant growth and development. Elucidation of how WRKY TFs help to exert these functions will certainly be assisted in the near future by the ability to monitor specific WRKY TF interactions with DNA/chromatin on a global basis. This will allow us to construct testable hypotheses regarding how WRKY factors can influence diverse metabolic pathways and overall cellular physiology. 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[W] The online version of this article contains Web-only data. www.plantphysiol.org/cgi/doi/10.1104/pp.109.138990 © 2009 American Society of Plant Biologists This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Büttner, Daniela; He, Sheng Yang

doi: 10.1104/pp.109.139089pmid: 19458111

Many gram-negative plant and animal pathogenic bacteria employ a type III secretion system (T3SS) to subvert and colonize their respective host organisms. The T3SS injects effector proteins directly into the cytosol of eukaryotic cells and thus allows the manipulation of host cellular activities to the benefit of the pathogen. In plant pathogenic bacteria, T3SSs are encoded by hrp (for hypersensitive response and pathogenicity) genes, which are so named because they are required for bacteria to cause disease in susceptible plants and to elicit the hypersensitive response in resistant plants (Lindgren et al., 1986). The hypersensitive response is a rapid local cell death at the infection site that restricts bacterial multiplication and is triggered by individual effector proteins in plants carrying a corresponding resistance gene (Dangl and Jones, 2001). hrp genes were found in almost all major gram-negative bacterial plant pathogens (e.g. Pseudomonas syringae, Xanthomonas spp., Ralstonia solanacearum, and Erwinia spp.), illustrating a central role of the T3SS in mediating diverse plant-bacteria interactions (Alfano and Collmer, 2004; He et al., 2004; Büttner and Bonas, 2006). In this Update, we highlight some basic as well as recent experiments that have collectively yielded molecular insights into general principles and unique properties of T3SSs in plant pathogenic bacteria. Environmental conditions that influence the level of hrp gene expression during the infection have been reviewed recently (Tang et al., 2006) and will not be discussed here. T3SSs: FILAMENTOUS SUPRAMOLECULAR STRUCTURES Delivery of effector proteins from the cytoplasm of gram-negative bacteria to the plant cell interior requires the T3SS to transport proteins across multiple physical barriers: the two bacterial membranes separated by a peptidoglycan layer and the plasma membrane of the plant cell, which is surrounded by a thick cell wall (Fig. 1A Figure 1. Open in new tabDownload slide Schematic representation of the T3SS from plant (A) and animal (B) pathogenic bacteria. The secretion apparatus spans both bacterial membranes and is associated with a cytoplasmic ATPase. The T3SS from plant pathogenic bacteria is connected to an extracellular pilus that presumably spans the plant cell wall. The T3SS system from animal pathogenic bacteria is associated with a short extracellular needle, which serves as a transport channel for secreted proteins. The needle is linked via the so-called tip complex to the translocon, which forms a proteinaceous channel in the host plasma membrane and allows transport of effector proteins into the host cell cytosol. Evidence for the presence of a tip complex in plant pathogenic bacteria is still missing. IM, Inner membrane; OM, outer membrane; PM, plasma membrane. Figure 1. Open in new tabDownload slide Schematic representation of the T3SS from plant (A) and animal (B) pathogenic bacteria. The secretion apparatus spans both bacterial membranes and is associated with a cytoplasmic ATPase. The T3SS from plant pathogenic bacteria is connected to an extracellular pilus that presumably spans the plant cell wall. The T3SS system from animal pathogenic bacteria is associated with a short extracellular needle, which serves as a transport channel for secreted proteins. The needle is linked via the so-called tip complex to the translocon, which forms a proteinaceous channel in the host plasma membrane and allows transport of effector proteins into the host cell cytosol. Evidence for the presence of a tip complex in plant pathogenic bacteria is still missing. IM, Inner membrane; OM, outer membrane; PM, plasma membrane. ). It is widely believed that the T3SS provides a continuous channel for effector proteins to travel from the bacterial cytoplasm directly into the cytoplasm of eukaryotic cells. Central to this belief is the observation that T3SSs in different bacteria invariably assemble filamentous supramolecular structures (He et al., 2004). Although the first T3SS-associated filamentous structure was discovered in the plant pathogen P. syringae (Roine et al., 1997), the most elegant and seminal work in the characterization of T3SS supramolecular structures was carried out in the mammalian pathogen Salmonella enterica. Work of Kubori and colleagues (1998) revealed that the T3SS of S. enterica consists of two pairs of rings that interact with the cytoplasmic and the outer membrane, respectively, and an extracellular filamentous extension that is 8 nm in diameter and 80 nm in length, resembling a needle, which is also present in other animal pathogenic bacteria (Kubori et al., 1998; Marlovits et al., 2004, 2006; Galán and Wolf-Watz, 2006; Fig. 1B). The needle presumably serves as conduit for secreted proteins to the eukaryotic plasma membrane. Notably, in enteropathogenic Escherichia coli, the T3SS needle is connected to an additional filament that consists of the EspA protein and reaches a length of up to 600 nm (Daniell et al., 2001; Sekiya et al., 2001). It was therefore suggested that the EspA filament helps to penetrate the glycocalyx on the surface of the intestinal epithelium. A similar structure is associated with the needle of one T3SS from Salmonella typhimurium (Chakravortty et al., 2005). So far, a complete T3SS supramolecular structure has not been purified from plant pathogenic bacteria. However, as mentioned above, T3SS filaments called Hrp pili have been found and characterized in all major plant pathogens that contain an active T3SS (Roine et al., 1997; Van Gijsegem et al., 2000; Jin et al., 2001; Weber et al., 2005; Table I Table I. Contribution of secreted and cytoplasmic control proteins from plant pathogenic bacteria to virulence Bacterial Species/Predicted Protein Function . Proteina . Localizationb . Contribution to T3S and Virulence . Reference . E. amylovora:     Pilus protein HrpA S Essential for disease and T3S Jin et al. (2001)     Translocon proteins HrpK S Homologous to HrpK1 from P. syringae, dispensable for virulence Nissinen et al. (2007)     Control protein HrpN S+T Harpin, contributes to translocation of the effector DspA/E Bocsanczy et al. (2008) HrpJ S Required for virulence, contributes to secretion of harpins and translocation of the effector DspA/E Nissinen et al. (2007); Bocsanczy et al. (2008) P. syringae pv tomato:     Pilus protein HrpA S Essential for disease and T3S Roine et al. (1997)     Translocon protein HrpK1 S+T Contributes to disease and effector protein translocation Petnicki-Ocwieja et al. (2005)     Control proteins HrpZ1 S+T Harpin, forms ion channels Lee et al. (2001); Kvitko et al. (2007); Engelhardt et al. (2009) HrpW1 S+T Harpin, C-terminal pectate lyase domain HopAK1 S+T Harpin, C-terminal pectate lyase domain HrpZ1, HrpW, and HopAK1 contribute to effector protein translocation and disease HrpH T HrpH, HopP1, and HopAJ1 are predicted lytic transglycosylases and contribute to effector protein translocation and disease Oh et al. (2007) HopP1 T HopAJ1 NT HrpJ S+T Required for disease, contributes to T3S Fu et al. (2006) HrpP T Predicted T3S4 protein, essential for disease Morello and Collmer (2009) R. solanacearum:     Pilus protein HrpY S Essential for disease and T3S Van Gijsegem et al. (2000)     Translocon protein PopF1 S Essential for disease Meyer et al. (2006) PopF2 S Dispensable for disease Meyer et al. (2006)     Control proteins HpaB n.a. Required for disease Mukaihara et al. (2004) Xanthomonas spp.:     Pilus protein HrpEXcv S Essential for disease and T3S Weber et al. (2005)     Translocon protein HrpFXcv S Essential for disease and effector protein translocation Büttner et al. (2002) HrpFXoo S Contributes to disease Sugio et al. (2005)     Control proteins HpaAXcv S+T Contributes to disease, secreted regulator of HpaB Lorenz et al. (2008a) HpaBXcv C Global T3S chaperone, essential for disease Büttner et al. (2004) HpaCXcv C T3S4 protein, contributes to disease Lorenz et al. (2008a) HpaHXcv n.a. Predicted lytic transglycosylase, contributes to effector protein translocation and disease Büttner et al. (2007) Bacterial Species/Predicted Protein Function . Proteina . Localizationb . Contribution to T3S and Virulence . Reference . E. amylovora:     Pilus protein HrpA S Essential for disease and T3S Jin et al. (2001)     Translocon proteins HrpK S Homologous to HrpK1 from P. syringae, dispensable for virulence Nissinen et al. (2007)     Control protein HrpN S+T Harpin, contributes to translocation of the effector DspA/E Bocsanczy et al. (2008) HrpJ S Required for virulence, contributes to secretion of harpins and translocation of the effector DspA/E Nissinen et al. (2007); Bocsanczy et al. (2008) P. syringae pv tomato:     Pilus protein HrpA S Essential for disease and T3S Roine et al. (1997)     Translocon protein HrpK1 S+T Contributes to disease and effector protein translocation Petnicki-Ocwieja et al. (2005)     Control proteins HrpZ1 S+T Harpin, forms ion channels Lee et al. (2001); Kvitko et al. (2007); Engelhardt et al. (2009) HrpW1 S+T Harpin, C-terminal pectate lyase domain HopAK1 S+T Harpin, C-terminal pectate lyase domain HrpZ1, HrpW, and HopAK1 contribute to effector protein translocation and disease HrpH T HrpH, HopP1, and HopAJ1 are predicted lytic transglycosylases and contribute to effector protein translocation and disease Oh et al. (2007) HopP1 T HopAJ1 NT HrpJ S+T Required for disease, contributes to T3S Fu et al. (2006) HrpP T Predicted T3S4 protein, essential for disease Morello and Collmer (2009) R. solanacearum:     Pilus protein HrpY S Essential for disease and T3S Van Gijsegem et al. (2000)     Translocon protein PopF1 S Essential for disease Meyer et al. (2006) PopF2 S Dispensable for disease Meyer et al. (2006)     Control proteins HpaB n.a. Required for disease Mukaihara et al. (2004) Xanthomonas spp.:     Pilus protein HrpEXcv S Essential for disease and T3S Weber et al. (2005)     Translocon protein HrpFXcv S Essential for disease and effector protein translocation Büttner et al. (2002) HrpFXoo S Contributes to disease Sugio et al. (2005)     Control proteins HpaAXcv S+T Contributes to disease, secreted regulator of HpaB Lorenz et al. (2008a) HpaBXcv C Global T3S chaperone, essential for disease Büttner et al. (2004) HpaCXcv C T3S4 protein, contributes to disease Lorenz et al. (2008a) HpaHXcv n.a. Predicted lytic transglycosylase, contributes to effector protein translocation and disease Büttner et al. (2007) a Xcv, X. campestris pv vesicatoria; Xoo, X. oryzae pv oryzae. b Localization: C, cytoplasmic; n.a., not analyzed; NT, not translocated; S, secreted; T, translocated. Open in new tab Table I. Contribution of secreted and cytoplasmic control proteins from plant pathogenic bacteria to virulence Bacterial Species/Predicted Protein Function . Proteina . Localizationb . Contribution to T3S and Virulence . Reference . E. amylovora:     Pilus protein HrpA S Essential for disease and T3S Jin et al. (2001)     Translocon proteins HrpK S Homologous to HrpK1 from P. syringae, dispensable for virulence Nissinen et al. (2007)     Control protein HrpN S+T Harpin, contributes to translocation of the effector DspA/E Bocsanczy et al. (2008) HrpJ S Required for virulence, contributes to secretion of harpins and translocation of the effector DspA/E Nissinen et al. (2007); Bocsanczy et al. (2008) P. syringae pv tomato:     Pilus protein HrpA S Essential for disease and T3S Roine et al. (1997)     Translocon protein HrpK1 S+T Contributes to disease and effector protein translocation Petnicki-Ocwieja et al. (2005)     Control proteins HrpZ1 S+T Harpin, forms ion channels Lee et al. (2001); Kvitko et al. (2007); Engelhardt et al. (2009) HrpW1 S+T Harpin, C-terminal pectate lyase domain HopAK1 S+T Harpin, C-terminal pectate lyase domain HrpZ1, HrpW, and HopAK1 contribute to effector protein translocation and disease HrpH T HrpH, HopP1, and HopAJ1 are predicted lytic transglycosylases and contribute to effector protein translocation and disease Oh et al. (2007) HopP1 T HopAJ1 NT HrpJ S+T Required for disease, contributes to T3S Fu et al. (2006) HrpP T Predicted T3S4 protein, essential for disease Morello and Collmer (2009) R. solanacearum:     Pilus protein HrpY S Essential for disease and T3S Van Gijsegem et al. (2000)     Translocon protein PopF1 S Essential for disease Meyer et al. (2006) PopF2 S Dispensable for disease Meyer et al. (2006)     Control proteins HpaB n.a. Required for disease Mukaihara et al. (2004) Xanthomonas spp.:     Pilus protein HrpEXcv S Essential for disease and T3S Weber et al. (2005)     Translocon protein HrpFXcv S Essential for disease and effector protein translocation Büttner et al. (2002) HrpFXoo S Contributes to disease Sugio et al. (2005)     Control proteins HpaAXcv S+T Contributes to disease, secreted regulator of HpaB Lorenz et al. (2008a) HpaBXcv C Global T3S chaperone, essential for disease Büttner et al. (2004) HpaCXcv C T3S4 protein, contributes to disease Lorenz et al. (2008a) HpaHXcv n.a. Predicted lytic transglycosylase, contributes to effector protein translocation and disease Büttner et al. (2007) Bacterial Species/Predicted Protein Function . Proteina . Localizationb . Contribution to T3S and Virulence . Reference . E. amylovora:     Pilus protein HrpA S Essential for disease and T3S Jin et al. (2001)     Translocon proteins HrpK S Homologous to HrpK1 from P. syringae, dispensable for virulence Nissinen et al. (2007)     Control protein HrpN S+T Harpin, contributes to translocation of the effector DspA/E Bocsanczy et al. (2008) HrpJ S Required for virulence, contributes to secretion of harpins and translocation of the effector DspA/E Nissinen et al. (2007); Bocsanczy et al. (2008) P. syringae pv tomato:     Pilus protein HrpA S Essential for disease and T3S Roine et al. (1997)     Translocon protein HrpK1 S+T Contributes to disease and effector protein translocation Petnicki-Ocwieja et al. (2005)     Control proteins HrpZ1 S+T Harpin, forms ion channels Lee et al. (2001); Kvitko et al. (2007); Engelhardt et al. (2009) HrpW1 S+T Harpin, C-terminal pectate lyase domain HopAK1 S+T Harpin, C-terminal pectate lyase domain HrpZ1, HrpW, and HopAK1 contribute to effector protein translocation and disease HrpH T HrpH, HopP1, and HopAJ1 are predicted lytic transglycosylases and contribute to effector protein translocation and disease Oh et al. (2007) HopP1 T HopAJ1 NT HrpJ S+T Required for disease, contributes to T3S Fu et al. (2006) HrpP T Predicted T3S4 protein, essential for disease Morello and Collmer (2009) R. solanacearum:     Pilus protein HrpY S Essential for disease and T3S Van Gijsegem et al. (2000)     Translocon protein PopF1 S Essential for disease Meyer et al. (2006) PopF2 S Dispensable for disease Meyer et al. (2006)     Control proteins HpaB n.a. Required for disease Mukaihara et al. (2004) Xanthomonas spp.:     Pilus protein HrpEXcv S Essential for disease and T3S Weber et al. (2005)     Translocon protein HrpFXcv S Essential for disease and effector protein translocation Büttner et al. (2002) HrpFXoo S Contributes to disease Sugio et al. (2005)     Control proteins HpaAXcv S+T Contributes to disease, secreted regulator of HpaB Lorenz et al. (2008a) HpaBXcv C Global T3S chaperone, essential for disease Büttner et al. (2004) HpaCXcv C T3S4 protein, contributes to disease Lorenz et al. (2008a) HpaHXcv n.a. Predicted lytic transglycosylase, contributes to effector protein translocation and disease Büttner et al. (2007) a Xcv, X. campestris pv vesicatoria; Xoo, X. oryzae pv oryzae. b Localization: C, cytoplasmic; n.a., not analyzed; NT, not translocated; S, secreted; T, translocated. Open in new tab ). The essential contribution of Hrp pili to T3S and the results of in situ immunogold labeling experiments suggest that Hrp pili similarly to T3SS needles provide a protein transport channel for effector proteins to the host-pathogen interface (Jin and He, 2001; Li et al., 2002). Since Hrp pili are much longer (in the micrometer range) than the needle extension from animal pathogenic bacteria, they presumably span the thick plant cell wall, which is a major obstacle in the interkingdom protein transport between plant pathogenic bacteria and their host cells (Fig. 1A). Interestingly, the amino acid sequences of the major subunits of Hrp pili are hypervariable in different subspecies of bacterial pathogens, although the predicted secondary structures of these proteins are remarkably similar, consisting almost exclusively of α-helices (Lee et al., 2005; Weber and Koebnik, 2005). This observation led to the speculation that extracellular Hrp pili in plant pathogens may be rapidly evolving to avoid recognition by the plant defense surveillance system. Indeed, two studies have provided evidence for strong positive selection (generation of new beneficial alleles) or diversifying selection (generation of multiple different alleles) of Hrp pilus protein sequences in Xanthomonas spp. and P. syringae, respectively (Guttman et al., 2006; Weber and Koebnik, 2006). HOW ARE SUBSTRATE PROTEINS RECOGNIZED BY THE T3SS? T3S substrate proteins possess noncleavable secretion signals in the N-terminal protein regions, but no discernible amino acid or peptide similarities can be found (Michiels and Cornelis, 1991; Arnold et al., 2009; Samudrala et al., 2009). In fact, there has been some debate as to whether it is the amino acid or mRNA sequence that is recognized by the T3SS. The prevailing view is that it is the amphipathic nature and the amino acid composition of the N-terminal region of T3S substrate proteins that serves as a secretion signal (Galán and Wolf-Watz, 2006; Arnold et al., 2009; Samudrala et al., 2009). This view is consistent with the finding that some specific biophysical features are present in the first 50 amino acids of effector proteins from P. syringae: (1) solvent-exposed amino acids in the first five amino acids, (2) the lack of Asp or Glu residues in the first 12 amino acids, and (3) the amphipathicity and the enrichment of polar residues in the first 50 amino acids (Guttman et al., 2002; Petnicki-Ocwieja et al., 2002). Furthermore, recent bioinformatic analyses of effector proteins from plant and animal pathogenic bacteria revealed that the N-terminal 25 amino acids are enriched in Ser and coiled regions but lack Leu (Arnold et al., 2009; Samudrala et al., 2009). Taken together, these features may make the N-terminal regions of T3S substrate proteins structurally flexible and probably unfolded, an important prerequisite for their transport through the narrow inner channel of the T3SS, which is presumably only 2.8 nm in diameter as was shown for the T3SS from animal pathogenic bacteria (Marlovits et al., 2004, 2006; Galán and Wolf-Watz, 2006). For some T3S substrate proteins, however, the presence of an N-terminal secretion signal may not be sufficient for maximal secretion. In these cases, specific T3S chaperone proteins are needed. T3S chaperones are generally small (<170 amino acids), acidic (pI < 5.5), and often contain an amphipathic α-helix near the C terminus (Parsot et al., 2003). Interestingly, the chaperone-encoding genes are often located adjacent to the cognate effector genes, suggesting strong selection for their coexistence in the genome. Although many T3S chaperones are specifically required for the secretion of a cognate effector protein, some seem to be more general and are involved in the secretion of many substrate proteins, as reported for HpaB from Xanthomonas campestris pv vesicatoria (Parsot et al., 2003; Büttner et al., 2004, 2006; Table I). T3S chaperones presumably target their substrates to conserved components of the T3SS, such as the cytoplasmic ATPase that is associated with the secretion apparatus (see below; Fig. 1). There is also some evidence that T3S chaperones may contribute to the stability of at least some effector proteins inside bacteria. In the plant pathogen Erwinia amylovora, for example, the T3S chaperone DspF seems to be required for the stability of the effector protein DspA/E, a major virulence factor in this devastating pathogen (Gaudriault et al., 2002). Since the normal folding environment for effector proteins is inside the eukaryotic cell, it was proposed that some T3S chaperones prevent misfolding and thus subsequent degradation of their interaction partners in the bacterial cytoplasm. TRANSLOCATION OF EFFECTOR PROTEINS INTO THE HOST CELL How the T3SS penetrates the host plasma membrane is very much an open question. In principle, one could imagine that the T3SS needle/pilus may physically penetrate the membrane and/or cell wall of the eukaryotic cell, as suggested for the Yersinia needle (Hoiczyk and Blobel, 2001). Alternatively, the needle/pilus may connect to additional T3SS-associated protein complexes in the eukaryotic cell membrane and/or cell wall to provide a continuous conduit for the delivery of effector proteins into the eukaryotic cell. This hypothesis is supported by the finding that the T3SS secretes several translocator proteins. The function of these proteins is to facilitate the translocation of effector proteins across the eukaryotic cell membrane (i.e. they are not required for secretion of effector proteins across the bacterial envelope). In the mammalian pathogen Yersinia spp., three proteins (YopB, YopD, and LcrV) were shown to be involved in the formation of a protein complex (called a translocon) that inserts into the eukaryotic cell membrane. YopB and YopD form a proteinaceous transmembrane channel that is connected to the needle via a so-called tip complex consisting of LcrV (Håkansson et al., 1996; Neyt and Cornelis, 1999; Mueller et al., 2008). The tip complex presumably facilitates the assembly of the translocon and thus allows the continuous passage of effector proteins into the eukaryotic cell cytosol (Fig. 1B). YopB, YopD, and LcrV are not conserved among animal and plant pathogenic bacteria, suggesting that the mechanisms underlying effector protein translocation vary among different pathogens. In plant pathogenic bacteria, several putative translocator proteins of the T3SS have been identified, including HrpF from X. campestris pv vesicatoria, PopF1 and PopF2 from R. solanacearum, and HrpK proteins from P. syringae and E. amylovora (Table I; Fig. 2 Figure 2. Open in new tabDownload slide Protein identities among selected putative translocators and T3S4 proteins, respectively, from plant pathogenic bacteria and the animal pathogen Yersinia enterocolitica. Full-length protein sequences were compared using the BLASTP program (http://blast.ncbi.nlm.nih.gov). Numbers refer to the percentage of protein identity. Boxes with protein identities of 75% to 100% are shaded in dark gray, and boxes with protein identities of 25% to 50% are in light gray. The following protein sequences were used: HrpF (X. campestris pv vesicatoria strain 85-10, AAB86527), PopF1 (R. solanacearum GMI1000, CAD18706), PopF2 (R. solanacearum GMI1000, CAD18051), HrpK1 (P. syringae pv tomato DC3000, AAO54927), HrpK (E. amylovora, AAX39435), YopB (Y. enterocolitica, AAK69211), HpaC (X. campestris pv vesicatoria strain 85-10, CAJ22055), HpaP (R. solanacearum GMI1000, CAB58249), HrpP (P. syringae pv tomato DC3000, AAG33881), and YscP (Y. enterocolitica, AAK69225). n.s., Not significant (protein identity among full-length proteins was defined as not significant when regions with identical residues were smaller than 100 amino acids); Eam, E. amylovora; Psyr, P. syringae pv tomato DC3000; Rsol, R. solanacearum GMI1000; Xcv, X. campestris pv vesicatoria strain 85-10; Xoo, X. oryzae pv oryzae PXO99A; Yent, Y. enterocolitica. a, The region with 32% protein identity is restricted to 137 amino acids; b, the region with 24% protein identity is restricted to 205 amino acids; c, the region with 26% protein identity is restricted to 105 amino acids; d, the region with 27% protein identity is restricted to 147 amino acids. Figure 2. Open in new tabDownload slide Protein identities among selected putative translocators and T3S4 proteins, respectively, from plant pathogenic bacteria and the animal pathogen Yersinia enterocolitica. Full-length protein sequences were compared using the BLASTP program (http://blast.ncbi.nlm.nih.gov). Numbers refer to the percentage of protein identity. Boxes with protein identities of 75% to 100% are shaded in dark gray, and boxes with protein identities of 25% to 50% are in light gray. The following protein sequences were used: HrpF (X. campestris pv vesicatoria strain 85-10, AAB86527), PopF1 (R. solanacearum GMI1000, CAD18706), PopF2 (R. solanacearum GMI1000, CAD18051), HrpK1 (P. syringae pv tomato DC3000, AAO54927), HrpK (E. amylovora, AAX39435), YopB (Y. enterocolitica, AAK69211), HpaC (X. campestris pv vesicatoria strain 85-10, CAJ22055), HpaP (R. solanacearum GMI1000, CAB58249), HrpP (P. syringae pv tomato DC3000, AAG33881), and YscP (Y. enterocolitica, AAK69225). n.s., Not significant (protein identity among full-length proteins was defined as not significant when regions with identical residues were smaller than 100 amino acids); Eam, E. amylovora; Psyr, P. syringae pv tomato DC3000; Rsol, R. solanacearum GMI1000; Xcv, X. campestris pv vesicatoria strain 85-10; Xoo, X. oryzae pv oryzae PXO99A; Yent, Y. enterocolitica. a, The region with 32% protein identity is restricted to 137 amino acids; b, the region with 24% protein identity is restricted to 205 amino acids; c, the region with 26% protein identity is restricted to 105 amino acids; d, the region with 27% protein identity is restricted to 147 amino acids. ). It was shown that the secreted HrpF protein from X. campestris pv vesicatoria is essential for effector protein translocation and induces the formation of ion channels in artificial lipid bilayers, suggesting that it is a component of the predicted translocation channel (Büttner et al., 2002). HrpF is homologous to PopF1 and PopF2 from R. solanacearum (Fig. 2), which contribute to bacterial pathogenicity and effector protein translocation but are not required for efficient T3S (Table I). Interestingly, the phenotype of a popF1 popF2 double mutant can partially be restored upon ectopic expression of hrpF from X. campestris pv campestris, suggesting a functional similarity among putative translocon proteins from Xanthomonas spp. and R. solanacearum (Meyer et al., 2006). Notably, HrpF, PopF1, and PopF2 do not share significant protein identity with the predicted translocator HrpK1 from P. syringae (Fig. 2). HrpK1 is secreted and translocated by the T3SS and contributes to bacterial pathogenicity and effector protein translocation (Petnicki-Ocwieja et al., 2005; Table I). By contrast, the homologous HrpK protein from E. amylovora is dispensable for the bacterial interaction with the plant, suggesting the presence of additional accessory proteins (Table I). What could these other accessory proteins be? Many T3SSs in plant pathogenic bacteria secrete a family of extracellular proteins called harpins (Wei et al., 1992; He et al., 1993; Arlat et al., 1994). Harpins share the general properties of being Gly rich and heat stable and are able to induce a suite of plant defense responses when infiltrated into the plant apoplast at high concentrations (for review, see He et al., 2004). However, the physiological functions of harpins in promoting disease have been enigmatic ever since their discovery. Recent experimental evidence reported for P. syringae and E. amylovora suggests an exciting possibility that harpins are important for the translocation of effector proteins into the plant cell. For example, deletion of all four harpin-encoding genes (hrpZ1, hrpW1, hopAK1, and hopP1; Table I) in P. syringae pv tomato strain DC3000 leads to reduced effector protein translocation (Kvitko et al., 2007). This deleterious effect is severely enhanced upon additional deletion of the putative translocator gene hrpK1 (Kvitko et al., 2007). Importantly, effector secretion in culture is not affected in this polymutant deleted in harpin genes and hrpK1, suggesting additive effects of harpins and HrpK1 in effector translocation. Notably, HrpN, a major harpin secreted by E. amylovora, was recently shown to be essential for translocation of the DspA/E effector protein into the plant cell (Bocsanczy et al., 2008; Table I). It is currently unclear how harpins facilitate effector translocation. Several harpins contain intriguing motifs that suggest potential interactions with plant cell wall components. For example, the P. syringae harpins HrpW1 and HopAK1 contain a C-terminal pectate lyase-like domain (Kvitko et al., 2007). This domain in HrpW1 binds to calcium pectate, a major plant cell wall component (Charkowski et al., 1998). Even HrpZ1, which lacks an obvious plant cell wall-interacting domain, also seems to bind to the plant cell wall (Hoyos et al., 1996). These observations suggest that harpins may be involved in modifying (loosening?) the plant cell wall to facilitate the initial penetration of the T3SS pilus. Although very attractive, this possibility is not consistent with results from further genetic analyses of hrpK1 and harpin genes in P. syringae. Most strikingly, the polymutant lacking both hrpK1 and the four harpin genes could be complemented by either hrpK1 or individual harpin genes (Kvitko et al., 2007). These observations suggest that HrpK1 and harpins are functionally redundant and act at the same step of effector translocation. Notably, harpins are found to be associated with synthetic lipid membranes and to form pores (Lee et al., 2001; Racapé et al., 2005; Engelhardt et al., 2009). Further studies are needed to determine whether harpins and HrpK proteins form functional complexes in the plant plasma membrane and/or the plant cell wall in vivo. If such complexes are found and shown to be important for effector translocation, the next step would be to study a possible physical connection with the T3SS pilus in planta. HOW DO BACTERIAL PATHOGENS COORDINATE T3SS ASSEMBLY AND EFFECTOR TRANSLOCATION? T3S is presumably a hierarchical process. As the T3SS appears to be dedicated to delivering effector proteins, which function inside the eukaryotic cell, it would make sense if secretion and translocation of effector proteins occur after the T3SS is fully assembled to prevent excessive leakage of effector proteins into the extracellular milieu. If this is indeed the case, how does the T3SS prevent secretion of effector proteins before the extracellular parts (pilus and translocon) of the T3SS are assembled? Studies of the T3SS in animal pathogens suggest a fascinating substrate specificity switch process from secretion of the needle structural proteins to secretion of translocators and effector proteins (Cornelis et al., 2006; Ferris and Minamino, 2006). As a consequence, the needle extension in mammalian pathogens has a relatively well-defined length (e.g. 80 nm in Salmonella; Cornelis et al., 2006; Galán and Wolf-Watz, 2006). The substrate specificity switch is presumably mediated by T3S substrate specificity switch (T3S4) proteins that are secreted by the T3SS and interact with the C-terminal cytoplasmic domain of a member of the conserved YscU/FlhB protein family. YscU and homologous proteins are inner membrane components of the T3SS and possess a cytoplasmic domain that was proposed to be involved in T3S substrate recognition (Cornelis et al., 2006; Ferris and Minamino, 2006). In animal pathogenic bacteria, T3S4 proteins presumably induce a conformational change in the cytoplasmic domain of YscU or homologous proteins and thus alter the substrate specificity of the T3SS. According to a model proposed for the T3S4 protein YscP from Yersinia spp., the N terminus of YscP is attached to the growing needle, whereas the C terminus interacts with YscU and triggers the substrate specificity switch once the T3S4 protein is stretched (Journet et al., 2003; Cornelis et al., 2006; Ferris and Minamino, 2006). This hypothesis implies that T3S4 proteins act as molecular rulers that determine needle length. Notably, however, this molecular ruler model was challenged by the finding that T3S4 proteins control the formation of an inner rod structure inside the base of the T3SS (i.e. beneath the needle extension), suggesting that it is the formation of the inner rod that induces the switch in T3S substrate specificity (Marlovits et al., 2004, 2006; Wood et al., 2008). It is currently unknown whether plant and animal pathogenic bacteria employ similar mechanisms to control substrate specificity and length of extracellular appendages of the T3SS. In plant pathogenic bacteria, it is not yet clear whether or not T3SS pili have a defined length in vivo. In vitro sample preparation often shears long Hrp pili into shorter fragments, making it impossible to accurately estimate the full length of Hrp pili. Nevertheless, putative T3S4 proteins have also been identified in plant pathogenic bacteria. In X. campestris pv vesicatoria, the predicted T3S4 protein HpaC was shown to switch the substrate specificity of the T3SS from secretion of the putative inner rod protein HrpB2 to secretion of translocators and effector proteins. HpaC interacts with and presumably induces a conformational change in the C-terminal domain of HrcU, which is a member of the YscU/FlhB protein family (Lorenz et al., 2008b; Fig. 3 Figure 3. Open in new tabDownload slide Model of T3S in the plant pathogenic bacterium X. campestris pv vesicatoria. The secretion apparatus consists of approximately 20 components, nine of which, labeled with single letters here, are designated Hrc (Hrp conserved) because they are conserved among plant and animal pathogenic bacteria. The pilus protein HrpE and the putative inner rod protein HrpB2 are the first substrates that travel the T3SS. A yet unidentified signal activates a switch in the T3S substrate specificity that depends on the cytoplasmic T3S4 protein HpaC and an inner membrane component of the T3SS, HrcU. HrcU consists of four transmembrane helices and a C-terminal cytoplasmic domain (UC), which is proteolytically cleaved. HpaC presumably induces a conformational change in the C-terminal cytoplasmic domain of HrcU and activates secretion of translocon and effector proteins. Targeting of effector proteins to the secretion apparatus depends on the global T3S chaperone HpaB, which binds to multiple effector proteins. The activity of HpaB is normally inhibited by HpaA, which binds to HpaB in the bacterial cytoplasm. Secretion and translocation of HpaA after assembly of the T3SS liberates HpaB and is thus a prerequisite for the efficient translocation of effector proteins (Lorenz et al., 2008a). Secretion of all known T3S substrates depends on the ATPase HrcN, which was shown to disassemble HpaB-effector complexes. IM, Inner membrane; OM, outer membrane; PM, plasma membrane. Figure 3. Open in new tabDownload slide Model of T3S in the plant pathogenic bacterium X. campestris pv vesicatoria. The secretion apparatus consists of approximately 20 components, nine of which, labeled with single letters here, are designated Hrc (Hrp conserved) because they are conserved among plant and animal pathogenic bacteria. The pilus protein HrpE and the putative inner rod protein HrpB2 are the first substrates that travel the T3SS. A yet unidentified signal activates a switch in the T3S substrate specificity that depends on the cytoplasmic T3S4 protein HpaC and an inner membrane component of the T3SS, HrcU. HrcU consists of four transmembrane helices and a C-terminal cytoplasmic domain (UC), which is proteolytically cleaved. HpaC presumably induces a conformational change in the C-terminal cytoplasmic domain of HrcU and activates secretion of translocon and effector proteins. Targeting of effector proteins to the secretion apparatus depends on the global T3S chaperone HpaB, which binds to multiple effector proteins. The activity of HpaB is normally inhibited by HpaA, which binds to HpaB in the bacterial cytoplasm. Secretion and translocation of HpaA after assembly of the T3SS liberates HpaB and is thus a prerequisite for the efficient translocation of effector proteins (Lorenz et al., 2008a). Secretion of all known T3S substrates depends on the ATPase HrcN, which was shown to disassemble HpaB-effector complexes. IM, Inner membrane; OM, outer membrane; PM, plasma membrane. ). In contrast to T3S4 proteins from animal pathogenic bacteria, however, HpaC is not secreted by the T3SS (Table I). Furthermore, HpaC does not control secretion of the pilus protein HrpE, suggesting that HpaC is not involved in length determination of the pilus (Lorenz et al., 2008b). T3S4 proteins are not highly conserved among different plant and animal pathogenic bacteria, and no significant sequence identity is detected between HpaC from X. campestris pv vesicatoria and the predicted T3S4 protein HrpP from P. syringae (Fig. 2). HrpP is secreted and translocated by the T3SS from P. syringae and is required for efficient secretion of all T3S substrates that were tested, including the pilus protein HrpA and the predicted inner rod component HrpB (Morello and Collmer, 2009). Thus, HrpP presumably does not act similarly to known T3S4 proteins from animal pathogenic bacteria that switch the substrate specificity of the T3SS from early (inner rod proteins and needle proteins) to late (translocators and effector proteins) T3S substrates. In addition to T3S4 proteins, T3S in plant pathogenic bacteria is controlled by other accessory proteins that act in the bacterial cytoplasm or are secreted by the T3SS (Table I). For instance, it was shown that the secreted HrpJ proteins from E. amylovora and P. syringae are required for efficient T3S (Fu et al., 2006; Bocsanczy et al., 2008). Both proteins contribute to the secretion of harpin proteins and thus may indirectly affect effector protein translocation (Fu et al., 2006; Nissinen et al., 2007; Bocsanczy et al., 2008; see above). In X. campestris pv vesicatoria, translocation of effector proteins is differentially regulated by the global T3S chaperone HpaB, which specifically promotes the translocation of a certain class of effector proteins (Büttner et al., 2006). The activity of HpaB is presumably controlled by the secreted regulator HpaA that binds to HpaB in the bacterial cytoplasm and allows secretion of extracellular components of the T3SS. After assembly of the T3SS, secretion of HpaA liberates HpaB and thus activates secretion and translocation of effector proteins (Lorenz et al., 2008a). In addition to HpaB, translocation of a certain set of effectors from X. campestris pv vesicatoria requires HpaH, which is a predicted lytic transglycosylase that might facilitate assembly of the T3SS (Büttner et al., 2007). Interestingly, experimental evidence for a role of lytic transglycosylases in the regulation of effector protein translocation was also reported for P. syringae (Oh et al., 2007; Table I). However, the precise role of lytic transglycosylases from plant pathogenic bacteria during T3S is not yet understood. ENERGY SOURCE FOR POWERING T3S The final aspect of T3SSs from plant pathogenic bacteria we would like to highlight concerns the energy source for T3S. All characterized T3SSs contain a cytoplasmic/inner membrane ATPase (HrcN in plant pathogens; Fig. 1) that bears sequence similarity to the catalytic β-subunit of the mitochondrial F1 ATPase. The F1 ATPase is a heterohexamer consisting of alternating α- and β-subunits with a central channel (Abrahams et al., 1994). However, the α-subunit equivalent is not found in T3SSs. Using hydrodynamic, cross-linking, and ultrastructural analyses, Pozidis et al. (2003) found that the P. syringae HrcN ATPase is activated by homo-oligomerization and is associated peripherally at the plasma membrane. The dodecamer oligomer has the highest ATPase activity. When viewed by electron microscopy, the dodecamer appears as an organized round particle with an o.d. of 13 nm (Pozidis et al., 2003). The dodecameric HrcN ATPase seems to form double hexameric stacks, as was found for other dodecameric traffic ATPases (Müller et al., 2006). Analysis of the HrcN ATPase from X. campestris pv vesicatoria revealed multiple protein-protein interactions between HrcN and cytoplasmic and inner membrane components of the T3SS, including the T3S4 protein HpaC and the global T3S chaperone HpaB (Lorenz and Büttner, 2009; Fig. 3). It is likely that the ATPase is required to release and unfold chaperone-bound effectors (Akeda and Galán, 2005; Lorenz and Büttner, 2009; Fig. 3). Furthermore, the HrcN ATPase presumably also serves as part of a docking site for accepting T3S substrates without accompanying T3SS chaperones (Lorenz and Büttner, 2009). Another potential energy source for T3S is the proton motive force, as reported for the assembly of bacterial flagella (Minamino and Namba, 2008; Paul et al., 2008). However, there is so far no experimental evidence that T3S in plant pathogens can occur in the absence of a functional ATPase (Lorenz and Büttner, 2009). It therefore remains to be determined experimentally whether proton motive force is also an energy source for powering virulence-associated T3SSs in plant pathogens. CONCLUDING REMARKS With this update, we hope to give readers an impression of the substantial progress made in the understanding of the T3SS in plant pathogenic bacteria following the initial discovery of the enigmatic hrp genes in the 1980s. Many questions remain to be answered: Can we visualize the real-time assembly and action of the T3SS in translocating effectors in planta? How does a bacterium make sure that all necessary effector proteins, which can be as many as several dozens, are injected into a host cell and in a timely manner? Does the assembly of the T3SS activate plant defense responses? If so, what is the nature of such defenses? Can we find chemicals and genetic engineering methods that could effectively and safely inhibit the T3SS during infection? Can we purify the complete T3SS from plant pathogens? Answering these fundamental questions should further advance our basic understanding of the T3S mechanism in plant-bacteria interactions and provide possible solutions to bacterial disease control. ACKNOWLEDGMENTS We thank Karen Bird and Christy Mecey, who helped us in the preparation of this review. Work in S.Y.H.'s laboratory is supported by funds from the Chemical Sciences, Geosciences, and Biosciences Division, Office of Basic Energy Sciences, Office of Science, U.S. Department of Energy (Award DE–FG02–91ER20021), the National Science Foundation, the U.S. Department of Agriculture, and the National Institutes of Health. Work in D.B.'s laboratory is supported by grants from the Deutsche Forschungsgemeinschaft (BU 2145/1–1) and the Sonderforschungsbereich SFB 648 “Molekulare Mechanismen der Informationsverarbeitung in Pflanzen.” LITERATURE CITED Abrahams JP, Leslie AG, Lutter R, Walker JE ( 1994 ) Structure at 2.8 Å resolution of F1-ATPase from bovine heart mitochondria. 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Gelvin, Stanton B.

doi: 10.1104/pp.109.139873pmid: 19439569

Members of the genus Agrobacterium cause the neoplastic diseases crown gall, hairy root, and cane gall on numerous plant species. Extensive genetic analyses conducted in the 1980s identified key bacterial genes involved in virulence. During the past decade, however, genomic technologies have revealed numerous additional bacterial genes that more subtly influence transformation. The results of these genomic analyses allowed scientists to develop a more integrated view of how Agrobacterium interacts with host plants. In a similar manner, genomic technologies have identified numerous plant genes important for Agrobacterium-mediated genetic transformation. Knowledge of these genes and their roles in transformation has revealed how Agrobacterium manipulates its hosts to increase the probability of a successful transformation outcome. In this article, I review our current knowledge of Agrobacterium-plant interactions and how genomic and proteomic technologies have increased our understanding of this unique plant-microbe interaction. Agrobacterium species are phytopathogens that cause a variety of neoplastic diseases, including crown gall (Agrobacterium tumefaciens and Agrobacterium vitis), hairy root (Agrobacterium rhizogenes), and cane gall (Agrobacterium rubi). Virulent strains of Agrobacterium contain tumor-inducing (Ti) or root-inducing (Ri) plasmids. During infection, enzymes encoded by plasmid-localized virulence (vir) genes process the T-DNA region of these plasmids. The resulting single-strand DNA (T-strand) linked to VirD2 protein exits the bacterium via a type IV protein secretion system and enters the plant cell. Within the plant, T-strands likely form complexes with other secreted virulence effector proteins, including VirE2, VirE3, VirD5, and VirF, and supercomplexes with plant proteins as they traverse the cytoplasm and target the nucleus. Once inside the nucleus, T-strands integrate randomly into the plant genome and express T-DNA-encoded transgenes. Two classes of T-DNA genes mediate the pathology of Agrobacterium infection. The first group, the oncogenes, either effect phytohormone production (iaa and ipt; Akiyoshi et al., 1984; Schroder et al., 1984), sensitize the plant to endogenous hormone levels (rol and other genes of pRi, gene5 and gene6 of pTi; Shen et al., 1988; Spanier et al., 1989; Tinland et al., 1990; Korber et al., 1991), or may be involved in chromatin remodeling (gene6b; Terakura et al., 2007). Expression of these genes results in tumorigenic or rhizogenic growth. A second set of genes directs the synthesis of various low M r compounds, opines, that can serve as energy sources for the inciting bacterial strain and can perhaps affect virulence (Veluthambi et al., 1989). For reviews, the reader should see Gelvin (2000, 2003), Tzfira and Citovsky (2001, 2003), McCullen and Binns (2006), and Citovsky et al. (2007). In addition, the reader is directed to an excellent new book on Agrobacterium biology (Tzfira and Citovsky, 2008). Most plant biologists, however, best know Agrobacterium as an agent of horizontal gene transfer that plays an essential role in basic scientific research and in agricultural biotechnology. In the 1980s, scientists learned to disarm (delete the oncogenes and, usually, the opine synthase genes) virulent Agrobacterium strains such that tissues infected by the bacteria could regenerate into normal plants (Bevan et al., 1983; Fraley et al., 1983; Herrera-Estrella et al., 1983). Substituting genes of interest for oncogenes and opine synthase genes resulted in plants expressing these novel transgenes and, thus, novel phenotypes. Although transgene substitution for oncogenes within T-DNA was initially conducted in cis (i.e. novel transgenes were placed within T-DNA of Ti-plasmids; Caplan et al., 1983; Fraley et al., 1985), the development of binary systems, in which T-DNA and virulence helper plasmids were separated into two different replicons (de Framond et al., 1983; Hoekema et al., 1983), greatly increased the utility of Agrobacterium as a vehicle for gene transfer in plant biology laboratories. Throughout its development as a gene jockeying tool, genomic studies on Agrobacterium and its plant hosts guided scientists in basic science and agricultural biotechnology developments. In this article, I review some of the key genomic methodologies and findings that have contributed to our knowledge of how Agrobacterium works and will contribute in the future better to utilize Agrobacterium's amazing gene transfer abilities in the laboratory and in the agricultural biotechnology industry. GENOMICS OF AGROBACTERIUM Whole-Genome Mutagenesis Although not frequently considered genomics, important early studies on A. tumefaciens and A. rhizogenes utilized whole-genome mutagenesis and mass phenotypic screening to define Agrobacterium genes important for transformation (i.e. T-DNA and Vir protein transfer) and tumorigenesis. Transposon mutagenesis was generally the method of choice because of the relatively random integration pattern of transposons in the bacterial genome and because the positions of transposon insertions could easily be determined by restriction endonuclease mapping. Thus, scientists localized genes involved in opine catabolism to a specific region of the Ti-plasmid and identified genes involved in crown gall tumorigenesis in the T-DNA, in regions of the Ti-plasmid not within the T-DNA (later to be identified as the virulence region), and in the bacterial chromosome (chromosomal virulence [chv] genes; Garfinkel and Nester, 1980; Holsters et al., 1980; Ooms et al., 1980; De Greve et al., 1981). More directed mutagenesis studies identified opine synthase genes and oncogenes in T-DNA regions of Ti- and Ri-plasmids and specific virulence genes within the vir gene region (Garfinkel et al., 1981; Leemans et al., 1981; Ooms et al., 1981; Ream et al., 1983; Inze et al., 1984; White et al., 1985; Stachel and Nester, 1986; Stachel and Zambryski, 1986). These studies involved testing of hundreds or thousands of individually mutagenized Agrobacterium strains for virulence, opine catabolism and synthesis, and tumor morphology and may thus be categorized as early Agrobacterium genomic studies. Rong et al. (1990) conducted a second type of genetic screening, using the promoter-less lacZ-containing transposon MuDI-1681, to identify plant-inducible Agrobacterium genes on the chromosome of A. tumefaciens A136 (C58 chromosomal background lacking a Ti-plasmid). These authors assayed several thousand randomly mutagenized Agrobacterium strains for induced gene expression on plates containing carrot root extract and X-gal. Insertion of the transposon into the picA gene revealed that this gene was >10-fold inducible by the root extract. The picA gene, currently identified as encoding a polygalacturonase-like protein (Atu3129), was the first identified plant-inducible Agrobacterium chromosomal gene. Agrobacterium Whole-Genome Sequencing Scientists had sequenced large portions of the Agrobacterium genome, including entire Ti-plasmids, by the late 1990s and the following years (Barker et al., 1983; Gielen et al., 1984; Slightom et al., 1986; Thompson et al., 1988; Ward et al., 1988; Rogowsky et al., 1990; Suzuki et al., 2000; Moriguchi et al., 2001; Oger et al., NC_010929; Kalogeroki and Winans, NC_002377). Generation of a complete nucleotide sequence of the nopaline-type strain A. tumefaciens C58 (Goodner et al., 2001; Wood et al., 2001), from which many Agrobacterium strains commonly used for plant genetic engineering are derived [e.g. GV3010::pMP90, C58-Z707, NT1(pKPSF2), EHA101/105, AGL-0/-1; see Lee and Gelvin (2008) for characteristics of these strains], opened the door for more extensive analyses of this important phytopathogen. A. tumefaciens C58 contains four replicons: a circular and a linear chromosome and two plasmids (pTiC58 and pAtC58). The genome contains approximately 30 insertion sequence elements and encodes an unusually large number of transporters (at least 153) and two-component regulatory systems (at least 25). Recently, Ulker et al. (2008) described the surprising observation that Agrobacterium can transfer its chromosomal DNA to plants. Interestingly, particular insertion sequence elements and transporter gene sequences are hot spots for chromosomal DNA transfer. The preferential appearance of these chromosomal sequences associated with T-DNA in Arabidopsis (Arabidopsis thaliana) and rice (Oryza sativa) T-DNA/plant DNA junctions suggests multiple mechanisms for chromosomal DNA mobilization during T-DNA transfer (Gelvin, 2008). The complete genome sequence and annotation of A. tumefaciens C58 is posted on http://depts.washington.edu/agro/. In addition to this biovar I A. tumefaciens strain, DNA sequence analysis of the Agrobacterium radiobacter biovar II strain K84 and the A. vitis biovar III strain S4 has recently appeared (Slater et al., 2009). A. radiobacter K84 is an especially important strain because it and its derivatives are widely used as biocontrol agents against tumorigenic Agrobacterium strains (Kerr and Panagopoulos, 1977; Jones et al., 1988). Comparative analysis of the sequences of the three Agrobacterium strains and several other species of the family Rhizobiaceae indicate a complex genome evolution, including the migration of gene blocks among replicons within and between species. Sequencing of other Agrobacterium strains (the biovar III strain A. vitis F5R19 and the biovar II strain A. rhizogenes A4) is in progress. Agrobacterium Transcriptional Profiling Based upon the A. tumefaciens C58 sequence, scientists have generated microarrays to probe the response of bacterial genes to environmental and chemical conditions important for Agrobacterium virulence and plant defense. The first such study investigated genes on the octopine-type Ti-plasmid pTiA6 and the nopaline-type Ti-plasmid pTiC58. Cho and Winans (2005) incubated bacteria individually containing these Ti-plasmids with acetosyringone (AS), a potent inducer of the vir gene regulon synthesized by wounded plant cells (Stachel et al., 1985; Stachel and Nester, 1986; Stachel and Zambryski, 1986). They used RNA extracted from induced and noninduced cells as probes of microarrays containing all Ti-plasmid genes. As expected, they observed an increase in all previously identified vir genes, along with several other Ti-plasmid genes previously not identified as part of the vir regulon. Most interestingly, they noted an increase in expression of all Ti-plasmid-encoded genes, suggesting that AS induction of the vir regulon increases the copy number of the Ti-plasmid relative to that of the bacterial chromosomes. Veluthambi et al. (1988) previously observed a similar increase in Ti-plasmid copy number in bacteria cocultivated with plant cells. Further investigation by Cho and Winans (2005) demonstrated that the repABC operon, essential for replication of these Ti-plasmids, was induced by AS. Induction was under the control of the two-component VirA/VirG regulatory system also responsible for vir gene induction. Thus, when Agrobacterium is in the environment of a wounded plant cell, the Ti-plasmid overreplicates, perhaps increasing the probability of T-DNA transfer to the plant. The plant wound environment in which Agrobacterium effects horizontal gene transfer is acidic (Fierer and Jackson, 2006), and the bacterium must maintain pH homeostasis. An acidic environment is also essential for efficient vir gene induction (Stachel et al., 1986), and two promoters regulate virG, one of which is acid inducible (Mantis and Winans, 1992; Chang and Winans, 1996). In addition, two chromosomal genes important for vir gene induction and transformation, chvG and chvI, are acid inducible (Charles and Nester, 1993; Li et al., 2002). With these facts in mind, Yuan et al. (2008a) conducted a microarray-based, whole-genome transcriptional profiling study of all Agrobacterium genes responding to acidic conditions. These authors identified 152 acid-responsive genes. These included previously identified acid-induced genes, genes involved in cell envelope synthesis, genes involved in exopolysaccharide (succinoglycan) synthesis and metabolism, several newly recognized acid-inducible vir genes (virE0, virE1, virH1, and virH2), and genes encoding a recently described type VI secretion system (Wu et al., 2008). Acidic conditions repressed a number of genes, including some involved in motility, chemotaxis, and cellular metabolism. Salicylic acid (SA) is a major signaling molecule that is important for plant defense responses. Although induction of SA and downstream plant defense genes by bacterial elicitation is well studied, fewer reports have investigated the effect of plant-derived SA on pathogen gene expression. Two groups used microarray analysis to investigate the effect of SA on the accumulation of Agrobacterium transcripts. Yuan et al. (2007) showed that SA, at concentrations that do not influence bacterial growth (2–8 μ m), inhibits vir gene expression in acidified medium containing AS. At higher concentrations (>10 μ m), SA inhibits bacterial growth in acidic medium. Transcriptional profiling of RNA from bacteria incubated for 6 h with AS and 6 μ m SA indicated that expression of Ti-plasmid-localized vir genes and the repABC genes was repressed. However, SA induced a number of Agrobacterium genes, including attKLM, which encodes a quormone degradation system. Because plants deficient in SA production are hypersusceptible to Agrobacterium transformation, whereas elicitation with SA decreased virulence (Yuan et al., 2007; Anand et al., 2008; Veena and S.B. Gelvin, unpublished data), these data suggest that the plant signaling molecule SA may inhibit transformation by shutting down vir gene expression and consequently T-DNA transfer. Anand et al. (2008) noted another effect of SA on Agrobacterium. Bacteria treated with 100 μ m SA did not efficiently attach to plant cells. Thus, bacterial attachment may be yet another process that is disrupted by this plant hormone. To explore further the effects of plant-released signal molecules on Agrobacterium gene expression, Yuan et al. (2008b) incubated bacteria with physiological levels of SA, indole-3-acetic acid (IAA), and γ-amino butyric acid (GABA) that do not inhibit Agrobacterium growth. Previous data had indicated that each of these compounds affect Agrobacterium virulence (Chevrot et al., 2006; Liu and Nester, 2006; Yuan et al., 2007; Anand et al., 2008). Incubation of Agrobacterium at acid pH with each of these compounds, followed by microarray analysis, revealed 100 to 200 genes for each treatment whose expression was modulated. In some instances, different compounds affected the same genes, whereas numerous Agrobacterium genes showed differential regulation by one compound only. IAA inhibited expression of the entire Ti-plasmid-localized vir regulon but did not have appreciable effects on expression of chromosomal virulence genes. Thus, the effect of IAA on vir regulon induction was similar to that of SA. However, the effects of SA and GABA on Agrobacterium gene expression were generally very different. In a most interesting exception, SA and GABA both induced the attKLM operon, which is involved in destroying the quorum sensing homoserine lactone that serves as a signaling molecule between Agrobacterium cells. In addition, seven genes were coregulated by IAA, SA, and GABA. Most of these were transporters, and mutation of some of these resulted in altered AttM lactonase activity. Taken together, these data suggest that at later times during Agrobacterium infection, plant signal molecules shut down vir gene expression (which is no longer needed once infection has been established) and may destroy quorum sensing signals. Agrobacterium Proteomics Engstrom et al. (1987) conducted the first proteomic study of Agrobacterium. Using one-dimensional SDS-PAGE, they identified 10 to 15 protein bands that appeared in various Agrobacterium strains following incubation with the vir gene inducer AS. Several of these bands corresponded to VirB membrane proteins comprising the type IV secretion system that transfers T-DNA and virulence effector proteins to plants. They also identified VirF and VirE2, two proteins of the vir regulon. In addition, they detected a number of other AS-induced proteins encoded by the Ti-plasmid or by the Agrobacterium chromosome. Similarly, Rong et al. (1990) detected by one-dimensional SDS-PAGE 10 plant-induced Agrobacterium chromosomal protein bands. Rosen et al. (2004) made the first attempt at experimentally defining the Agrobacterium proteome. Using two-dimensional gel electrophoresis, they detected approximately 300 proteins from exponentially growing bacteria. Interestingly, approximately 10% of the proteins were represented by multiple spots on the gel. The authors suggested that a high level of protein modification of the proteome occurs. Similar studies by this group (Rosen et al., 2001, 2002) investigated stress (high temperature, oxidative, and mild acid conditions) and heat shock-induced proteins of Agrobacterium. This group also identified proteins induced when the bacteria were incubated with and bound to cut tomato (Solanum lycopersicum) root segments, simulating plant infection conditions (Rosen et al., 2003). As controls, they examined, by two-dimensional gel electrophoresis, proteins from unbound bacteria and bacteria not incubated with root segments. Incubation of bacteria with roots induced approximately 30 proteins, regardless of whether the bacteria bound to the root segments or not. Although incubation with root segments induced ChvE, AttK, and AttM (all proteins involved in virulence), their experiments detected no induced Ti-plasmid-encoded virulence proteins. Because, for example, VirE2 is a major virulence protein induced by phenolic molecules such as AS (Engstrom et al., 1987; Lai et al., 2006), the results of this study indicate either that vir gene induction did not efficiently occur or that it occurred in only a small percentage of the bacteria. More recently, Lai et al. (2006) investigated Agrobacterium proteins induced by the phenolic vir regulon inducer AS. Using two-dimensional gel electrophoresis coupled with mass spectrometry, they identified 11 AS-induced proteins. Nine of these proteins were well-known Ti-plasmid-encoded Vir proteins (VirE2, several VirB proteins, Tzs, VirH1, and VirK), thus verifying their vir regulon induction conditions. In addition, they identified two proteins encoded by chromosomal genes, HspL (a small heat shock protein) and Y4mC (a protein of unknown function). Reverse transcription-PCR analysis indicated that transcripts of the genes encoding these proteins also were AS inducible and that induction was dependent upon the two-component sensing system VirA/VirG that mediates induction of the vir regulon. All vir regulon genes previously identified contain a vir box in their promoter regions (Das et al., 1986). The y4mC gene promoter similarly contains a vir box, but, interestingly, the hspL promoter does not. Thus, hspL activation by AS may be an indirect consequence of expression of Vir proteins. In addition, Wu et al. (2008) analyzed proteins secreted by Agrobacterium into the medium. They identified 12 proteins, including VirB1* (a cleaved fragment of VirB1 protein) and Hcp (hemolysin-coregulated protein). Hcp is secreted by a newly discovered type VI secretion system. GENOMICS OF PLANT GENES IMPORTANT FOR AGROBACTERIUM-MEDIATED GENETIC TRANSFORMATION Scientists have used a variety of genomic techniques to investigate plant genes important for Agrobacterium-mediated transformation. These include forward genetic screens to identify mutant plants with altered transformation susceptibility, yeast two-hybrid studies to detect plant proteins that interact with Virulence effector proteins, and transcriptional profiling to discover plant genes whose expression is altered following Agrobacterium infection. In addition, reverse genetic analyses have been used to probe the importance of candidate genes in the transformation process. Forward Genetic Screens for Plant Mutants with Altered Transformation Characteristics Plant species, and even different cultivars/genotypes of the same species, are notoriously varied in their transformation susceptibility (DeCleene and DeLey, 1976; Anderson and Moore, 1979; Conner and Commisse, 1992; van Wordragen and Dons, 1992; Bliss et al., 1999; Pena and Seguin, 2001; Somers et al., 2003; Shrawat and Lorz, 2006). In addition, Agrobacterium can transform Streptomyces, yeast, and other fungal species (Bundock et al., 1995, 2002; Piers et al., 1996; de Groot et al., 1998; Abuodeh et al., 2000; Kelly and Kado, 2002; Roberts et al., 2003; Schrammeijer et al., 2003; van Attikum and Hooykaas, 2003; Michielse et al., 2004), HeLa cells (Kunik et al., 2001), and sea urchin embryos (Bulgakov et al., 2006). Thus, Agrobacterium is incredibly promiscuous in its ability to mediate horizontal gene flow among numerous species of different phylogenetic kingdoms. A genetic basis for susceptibility to Agrobacterium exists in many crop species (Owens and Cress, 1984; Szegedi and Kozma, 1984; Smarrelli et al., 1986; Robbs et al., 1991; Bailey et al., 1994; Mauro et al., 1995), and Nam et al. (1997) also described a genetic basis for various degrees of susceptibility among approximately 40 Arabidopsis ecotypes. Large-scale forward genetic screening of approximately 20,000 T-DNA mutagenized Arabidopsis lines resulted in the first identification of plant genes involved in Agrobacterium-mediated transformation (Nam et al., 1999; Zhu et al., 2003b). These forward genetic analyses revealed >120 genes encoding proteins involved in transformation and, because the screen was not saturating (e.g. no gene was discovered more than once), the authors suggested that >200 Arabidopsis genes likely influence plant transformation susceptibility (Zhu et al., 2003b). The authors termed mutants with greatly decreased susceptibility to transformation rat (for resistant to Agrobacterium transformation) mutants and the corresponding mutant genes, rat genes. The identified genes represent most of the proposed transformation events that occur in the plant (bacterial attachment/biofilm formation, T-DNA and Virulence protein transfer to the plant, cytoplasmic trafficking and targeting of the proposed T-complex to the nucleus, virulence protein removal from the T-strand, T-DNA integration into the plant genome, and transgene expression). Examples of plant proteins identified in these initial genetic screens and mediating transformation include those involved in cell wall structure and biosynthesis (Rat1 and Rat4, and arabinogalactan and cellulose synthase-like [CslA9] proteins, respectively; Zhu et al., 2003a; Gaspar et al., 2004), cytoskeleton proteins potentially involved in cytoplasmic trafficking of T-complex components (actins and a kinesin; Zhu et al., 2003b), importin α and β proteins that may mediate nuclear targeting of T-complex components (Ballas and Citovsky, 1997; Bakó et al., 2003; Bhattacharjee et al., 2008), chromatin proteins such as various histones, histone acetyltransferases, histone deacetylases, and histone chaperones that may facilitate T-DNA integration into the plant genome (Nam et al., 1999; Mysore et al., 2000; Yi et al., 2002, 2006; Tian et al., 2003; Zhu et al., 2003b; Gelvin and Kim, 2007), and histone proteins that can increase transgene expression (G. Tenea and S.B. Gelvin, unpublished data). The nature of these rat genes has stimulated reverse genetic experiments to determine the potential roles of candidate genes in the transformation process (see below). Recently, the Gelvin laboratory further identified several Arabidopsis mutants that are hypersusceptible to Agrobacterium transformation (hat mutants and, therefore, hat genes; Fig. 1 Figure 1. Open in new tabDownload slide Activation tagging identifies Arabidopsis mutants that are hypersusceptible to Agrobacterium transformation (hat mutants). Root segments of wild-type (ecotype Wassilewskija) and T-DNA activation-tagged mutants (Weigel et al., 2000) were inoculated with the tumorigenic strain A. tumefaciens A208 at low inoculum density (106 colony forming units/mL). After 2 d of cocultivation, the root segments were transferred to Murashige and Skoog medium lacking phytohormones and tumors were allowed to develop (Zhu et al., 2003b). The plates were photographed after 4 weeks. Note the larger and more numerous tumors formed on root segments of the hat1 mutant line, compared to the tumors formed on wild-type roots. The hat1 mutant has a T-DNA activation tag inserted into a cellulose synthase-like gene. Expression of a neighboring UGT gene is greatly enhanced in the hat1 mutant. Figure 1. Open in new tabDownload slide Activation tagging identifies Arabidopsis mutants that are hypersusceptible to Agrobacterium transformation (hat mutants). Root segments of wild-type (ecotype Wassilewskija) and T-DNA activation-tagged mutants (Weigel et al., 2000) were inoculated with the tumorigenic strain A. tumefaciens A208 at low inoculum density (106 colony forming units/mL). After 2 d of cocultivation, the root segments were transferred to Murashige and Skoog medium lacking phytohormones and tumors were allowed to develop (Zhu et al., 2003b). The plates were photographed after 4 weeks. Note the larger and more numerous tumors formed on root segments of the hat1 mutant line, compared to the tumors formed on wild-type roots. The hat1 mutant has a T-DNA activation tag inserted into a cellulose synthase-like gene. Expression of a neighboring UGT gene is greatly enhanced in the hat1 mutant. ; N. Sardesai and S.B. Gelvin, unpublished data). Arabidopsis lines containing T-DNA activation tags (Weigel et al., 2000) provide a resource for overexpressed genes that may influence transformation susceptibility. When roots of these mutagenized plants were assayed at low bacterial inoculum conditions (102- to 103-fold lower than that usually used to screen for rat mutants), we identified seven independent lines that displayed increased levels of transformation relative to that of wild-type control plants. T-DNA/plant DNA junction sequences from five hat mutants identified several new genes involved in transformation susceptibility, including a cellulose synthase-like protein (CslB5), a potassium transporter family protein (two independent T-DNA insertion lines), a UDP-glucosyltransferase (UGT), and a myb transcription factor (MTF). Overexpression of the UGT cDNA in wild-type plants confirmed that this gene is a hat gene. Interestingly, metabolic profiling of roots from UGT overexpressing plants indicated alterations in the levels of key defense compounds, and microarray analyses of these plants revealed decreased expression of most genes in the phenypropanoid biosynthetic and SA signaling pathways (N. Sardesai, A. Perera, R. Doerge, and S.B. Gelvin, unpublished data). These results further indicate that plant defense response signaling pathways are involved in susceptibility to Agrobacterium-mediated transformation (see the discussion of transcriptional profiling below). The hat3 mutant has a T-DNA activation tag inserted into the 5′ untranslated region of an MTF gene. Although we could not isolate any homozygous hat3 mutants (suggesting that this MTF is essential for normal plant growth and development), heterozygous hat3 mutants are approximately 10-fold more susceptible to Agrobacterium-mediated transformation than are wild-type control plants (Fig. 2A; Figure 2. Open in new tabDownload slide An MTF negatively affects transformation susceptibility. A, Transformation efficiency of the hat3 (MTF) mutant and its wild-type control (ecotype Columbia-7 [Col-7]) and three independent T-DNA insertion mutants in the MTF gene (mtf1, -2, and -3) and its wild-type control (ecotype Columbia-0 [Col-0]). Root segments were inoculated with the tumorigenic strain A. tumefaciens A208 at low inoculum density (105 colony forming units/mL). After 2 d of cocultivation, the root segments were transferred to Murashige and Skoog medium lacking phytohormones and tumors were allowed to develop (Zhu et al., 2003b). The plates were photographed after 4 weeks. Note the more numerous tumors formed on root segments of MTF mutant lines compared to the tumors formed on wild-type roots. B, Map of the MTF gene mutated in the hat3 mutant. Numbers below the bar indicate nucleotides (+1 is the start site of translation). hat3, mtf1, mtf2, and mtf3 indicate the positions of three independent T-DNA insertions into the gene. Figure 2. Open in new tabDownload slide An MTF negatively affects transformation susceptibility. A, Transformation efficiency of the hat3 (MTF) mutant and its wild-type control (ecotype Columbia-7 [Col-7]) and three independent T-DNA insertion mutants in the MTF gene (mtf1, -2, and -3) and its wild-type control (ecotype Columbia-0 [Col-0]). Root segments were inoculated with the tumorigenic strain A. tumefaciens A208 at low inoculum density (105 colony forming units/mL). After 2 d of cocultivation, the root segments were transferred to Murashige and Skoog medium lacking phytohormones and tumors were allowed to develop (Zhu et al., 2003b). The plates were photographed after 4 weeks. Note the more numerous tumors formed on root segments of MTF mutant lines compared to the tumors formed on wild-type roots. B, Map of the MTF gene mutated in the hat3 mutant. Numbers below the bar indicate nucleotides (+1 is the start site of translation). hat3, mtf1, mtf2, and mtf3 indicate the positions of three independent T-DNA insertions into the gene. N. Sardesai and S.B. Gelvin, unpublished data). Three additional independent T-DNA insertions in this gene are also hat mutants (Fig. 2B), indicating that this MTF is a negative regulator of Agrobacterium-mediated transformation. Microarray analysis of RNA isolated from roots of mtf mutant plants indicated that a WRKY transcription factor gene was expressed to a lower level in the mutant. A homozygous T-DNA insertion into this WRKY transcription factor gene also resulted in a hat phenotype. This WRKY transcription factor is involved in regulating plant defense responses, once again implicating plant defense responses as a component of transformation susceptibility. As an alternative to screening T-DNA insertion mutants for hat and rat phenotypes, Anand et al. (2007b) used virus-induced gene silencing to investigate Nicotiana benthamiana genes important for Agrobacterium-mediated transformation. The authors identified 21 genes whose expression, when lowered, resulted in an altered crown gall phenotype. Proteins encoded by these genes include a nodulin-like protein, α-expansin,VIP1, importin-α, and histones H2A and H3. Identification of rat and hat mutants emphasizes the utility of large-scale forward genetic screens to understand the plant contribution to the Agrobacterium-mediated transformation process. Yeast Two-Hybrid Screening for Plant Proteins That Interact with Virulence Effector Proteins A. tumefaciens transfers at least five Virulence effector proteins to plants (VirD2 attached to the T-strand, VirD5, VirE2, VirE3, and VirF; Otten et al., 1984; Stahl et al., 1998; Vergunst et al., 2000, 2003, 2005; Schrammeijer et al., 2003). In addition, A. rhizogenes transfers GALLS-FL (full-length) and GALLS-CT (C-terminal) to plant cells (Hodges et al., 2006). Several laboratories have used yeast two-hybrid systems to search for plant proteins that interact with these effector proteins or with other proteins that appear on the bacterial surface. The rationale for these experiments is that if a plant protein interacts with an Agrobacterium protein, it is likely that this plant protein is involved in the transformation process. VirB2 is the major constituent protein of the Agrobacterium T-pilus (Lai and Kado, 1998). The T-pilus is an important bacterial structure that may come into contact with the plant during T-DNA and Vir protein transfer. Although proteins on the plant cell surface had previously been implicated in bacterial adhesion (Neff and Binns, 1985; Gurlitz et al., 1987; Neff et al., 1987; Wagner and Matthysse, 1992; Swart et al., 1994; Clauce-Coupel et al., 2008), no previously identified plant surface protein directly influenced bacterial virulence. Hwang and Gelvin (2004) used the processed form of VirB2 (Lai and Kado, 1998) as a bait protein to screen in yeast for Arabidopsis VirB2 interacting proteins. In addition to a RAB8 GTPase, they identified three reticulon domain proteins termed BTI1, -2, and -3 (for VirB2 Interacting proteins 1, 2, and 3). Decreasing expression of the Arabidopsis BTI genes by T-DNA mutagenesis or RNA interference (RNAi) resulted in reduced susceptibility to Agrobacterium-mediated transformation, whereas overexpression of BTI1 made the plant hypersusceptible to transformation. As would be expected of a protein that interacts with the T-pilus, the BTI proteins localize to the plant surface. Although more experiments need to be conducted, the BTI proteins may serve as receptors for VirB2 protein on the T-pilus. In addition to VirB2, the role of VirB5 (a minor T-pilus constituent) needs further exploration. In animal pathogens that have type IV secretion systems, VirB5 orthologs, such as CagL, may serve as specialized adhesins that interacts with human integrin β1 and fibronectin during bacterial/animal cell contact (Backert et al., 2008). It would be interesting to determine, using yeast two-hybrid systems, whether Agrobacterium VirB5 interacts with a specific plant surface protein. VirD2 is the pilot protein that guides the T-strand through the type IV secretion system into the plant cell, through the plant cytoplasm, and into the nucleus. VirD2 may also influence T-DNA integration into the plant genome (Tinland et al., 1995; Mysore et al., 1998). It is therefore likely that VirD2 interacts with plant proteins during this journey, and yeast two-hybrid analyses have identified a number of these proteins. The first of these was the nuclear transfer importin α protein AtKAPα (Ballas and Citovsky, 1997), now known as IMPa-1 (Bhattacharjee et al., 2008). The Arabidopsis genome encodes nine importin α proteins, and VirD2 interacts in yeast with all tested importin α isoforms (Ballas and Citovsky, 1997; Bakó et al., 2003; Bhattacharjee et al., 2008). Additionally, bimolecular fluorescence complementation studies in planta indicated that each of these isoforms interacts with VirD2 and localizes the complex to the nucleus (Bhattacharjee et al., 2008). Yeast two-hybrid screening additionally identified several other plant proteins that interact with VirD2. These include several cyclophilins (Deng et al., 1998; Bakó et al., 2003), the kinase CAK2Ms (Bakó et al., 2003), and a protein phosphatase PP2C (Tao et al., 2004). Interaction with these latter two proteins suggested that VirD2 may be a phosphoprotein. Bakó et al. (2003) confirmed this hypothesis, and Tao et al. (2004) showed that PP2C can regulate nuclear entry of VirD2. The single-strand DNA binding protein VirE2 is important for transformation. Agrobacterium strains mutant for virE2 are highly attenuated in virulence (Stachel and Nester, 1986). VirE2 plays numerous important roles within the plant cell (Citovsky et al., 1992; Ward and Zambryski, 2001) and therefore likely interacts with numerous plant proteins. Yeast two-hybrid analyses have confirmed these interactions. VirE2 interacts with numerous importin α isoforms; however, only interaction with IMPa-4 results in nuclear localization (Bhattacharjee et al., 2008; Lee et al., 2008). VirE2 also interacts in yeast with the VirE2 interacting proteins VIP1 and VIP2 (Tzfira et al., 2001; Anand et al., 2007a). Interaction of VirE2 with these proteins likely contributes to nuclear targeting and genomic integration of T-strands (Tzfira et al., 2001; Citovsky et al., 2004; Li et al., 2005; Loyter et al., 2005; Anand et al., 2007a; Bhattacharjee et al., 2008; Lacroix et al., 2008) VirF is a nonessential virulence protein for infection of most plant species. However, it is required for efficient transformation of a few species (Melchers et al., 1990; Regensburg-Tuink and Hooykaas, 1993). Schrammeijer et al. (2001) screened for VirF interacting proteins in yeast and identified a plant Skp1 ortholog. Skp1 (ASK1) is a component of the SCF ubiquitin ligase complex that identifies and marks proteins for degradation via the 26S proteosome. Indeed, experiments in both yeast and in planta indicated the importance of VirF in proteolysis of VirE2, suggesting that VirF plays a role in stripping VirE2 from T-strands prior to integration (Tzfira et al., 2004). VirE3 is a nuclear-localized Agrobacterium effector protein that may serve as a plant transcription factor (Schrammeijer et al., 2003; Garcia-Rodriguez et al., 2006). VirE3 may also substitute for plant-encoded VIP1 when this latter protein is limiting (Lacroix et al., 2005). In yeast, VirE3 interacts with several importin α isoforms, with pCsn5-1 (also known as AJH1), a component of the COP9 signalosome involved in protein degradation, and with pBrp, a plant transcriptional activator. An intriguing potential function for VirE3 may be as a molecular bridge to transport plant transcription factors to the nucleus where they may activate plant genes involved in tumorigenesis or transformation (Garcia-Rodriguez et al., 2006). GALLS-FL and GALLS-CT are two effector proteins encoded by some A. rhizogenes Ri-plasmids (Hodges et al., 2006). Although these proteins do not share sequence homology with A. tumefaciens VirE2, they can substitute for this essential A. tumefaciens virulence effector protein (Hodges et al., 2004, 2009). Recent yeast two-hybrid analysis using GALLS-FL as the bait identified a specific interacting plant protein (GALLS interacting protein [GIP]; Y. Wang and S.B. Gelvin, unpublished data). Bimolecular fluorescence complementation experiments confirmed GALLS-FL and GALLS-CT interaction with GIP in planta (L.-Y. Lee and S.B. Gelvin, unpublished data). GIP is encoded by one of an eight-member multigene family whose functions are unknown. Research in this author's laboratory is aimed at defining the role of GIP in both A. rhizogenes- and A. tumefaciens-mediated plant transformation. Host Transcriptional Profiling and Agrobacterium Infection Several recent studies have investigated host transcriptional responses to Agrobacterium infection or to crown gall tumorigenesis. Veena et al. (2003) used suppressive subtractive hybridization and DNA macroarrays to investigate the transcriptional response of tobacco BY-2 cells to infection by several nontumorigenic Agrobacterium strains. The authors used nontumorigenic strains to avoid complications resulting from phytohormone overproduction by expression of oncogenes and studied the initial plant response by limiting sampling to times <36 h after infection. The results of these experiments indicated that Agrobacterium exquisitely manipulates expression of the plant genome to facilitate transformation: plant genes important for transformation, such as those encoding histone proteins, were induced by the bacterium, whereas expression of genes involved in host defense responses was suppressed. Interestingly, Anand et al. (2007a) later showed that expression of numerous Arabidopsis histone genes was higher in wild-type Arabidopsis plants than in vip2 (VirE2 interacting protein 2) mutant plants. Arabidopsis vip2 mutant plants are highly recalcitrant to Agrobacterium-mediated transformation. These authors suggested that VIP2, a putative transcription factor, may play a role in maintaining high-level expression of histone genes important for transformation. Ditt et al. (2001) used a disarmed Agrobacterium strain to infect Ageratum cell cultures. Using cDNA/amplified fragment length polymorphism analyses of RNA extracted at relatively long times after infection (48 h), the authors identified a few genes whose expression was either repressed or induced by cocultivation with Agrobacterium. Whereas expression of most of the identified genes was similarly affected by cocultivation with Escherichia coli, expression of two genes (encoding a nodulin-like protein and a lectin-like protein kinase) was specifically induced by Agrobacterium infection. Ditt et al. (2006) also used Arabidopsis Affymetrix ATH1 microarrays to investigate host gene expression changes following infection of Arabidopsis suspension cell cultures with a tumorigenic Agrobacterium strain. Interestingly, the authors were only able to detect transcriptional changes 48 h after infection. In contrast to the results of Veena et al. (2003), the study of Ditt et al. (2006) indicated that Agrobacterium infection induced, rather than repressed, defense gene expression and that infection repressed expression of genes encoding proteins involved in cell proliferation. This latter observation was rather surprising considering that growth of the cell cultures was not slowed by bacterial infection. The seemingly opposing results of Ditt et al. (2006) and those of Veena et al. (2003) may be explained by the different plant culture systems used (tobacco and Arabidopsis) and the fact that one group used disarmed strains, whereas the other used tumorigenic strains that would result in the overproduction of phytohormones by the host and, eventually, production of tumors. Differential gene expression occurs in Arabidopsis crown gall tumors (Deeken et al., 2006). The expression of numerous genes, including those involved in cell wall biosynthesis, Suc degradation, transport, and glycolysis, is up-regulated in tumors, whereas expression of genes involved in photosynthesis, nitrogen metabolism, lipid metabolism, and amino acid synthesis is down-regulated. Differential gene expression in crown gall tumors correlated with altered solute profiles, leading the authors to speculate that metabolism in mature crown gall tumors occurs mainly anaerobically (Deeken et al., 2006). In addition to examining host transcriptional responses following Agrobacterium-mediated transformation or the development of crown gall tumors, Kim et al. (2007) used Arabidopsis Affymetrix ATH1 microarrays and custom macroarrays to investigate the transcriptional and methylation status of host T-DNA integration sites. The results of these assays indicated that, in the absence of selection, T-DNA target sites were not preferentially transcribed to a greater extent than were Arabidopsis genes in general and that T-DNA integration occurred without regard to the methylation status of the target DNA. Reverse Genetic Screening for Genes Required for Agrobacterium-Mediated Transformation A large number of studies have employed reverse genetic strategies to determine the role of candidate genes in the transformation process. Candidate genes include those identified by yeast two-hybrid and transcriptional profiling analyses, as well as additional members of multigene families when one family member clearly plays a role in virulence. Gene/expression disruption techniques have included T-DNA insertional mutagenesis and RNAi and antisense inhibition of gene expression. Overexpression of several plant genes has also resulted in a hat phenotype (Mysore et al., 2000; Tzfira et al., 2002; Hwang and Gelvin, 2004; Yi et al., 2006; G. Tenea, J. Spantzel, and S.B. Gelvin, unpublished data). Some genes confirmed as rat genes by these studies include those encoding BTI proteins (Hwang and Gelvin, 2004), various importin α family members (Bhattacharjee et al., 2008), VIP1 (Tzfira et al., 2001, 2002; Li et al., 2005), VIP2 (Anand et al., 2007a), Ku80 (West et al., 2002; Friesner and Britt, 2003; Li et al., 2005), DNA ligase IV (Friesner and Britt, 2003; van Attikum et al., 2003), SGA1 (G. Tenea and S.B. Gelvin, unpublished data), and various histones (Mysore et al., 2000; Yi et al., 2006; Anand et al., 2007b; G. Tenea and S.B. Gelvin, unpublished data). In addition, mutation of several genes involved in plant defense responses and signal transduction results in altered susceptibility to Agrobacterium-mediated transformation (Veena, N. Sardesai, and S.B. Gelvin, unpublished data). Crane and Gelvin (2007) conducted a large-scale reverse genetic screen for Arabidopsis rat mutants. Using 340 independent mutant lines containing RNAi constructions targeted against 109 chromatin genes, they identified 24 genes important, to various extents, for transformation. These genes encoded histone acetyltransferases, histone deacetylases, chromatin remodeling proteins, DNA methyltransferases, global transcription factors, histone H1, nucleosome assembly factors, SET domain proteins, and antisilencing group proteins. Some of these genes, such as HDT19, were previously implicated in the transformation process (Tian et al., 2003). Most interesting were three genes whose expression is important for T-DNA integration: HDT1, HDT2, and SGA1. HDT1 and HDT2 encode histone deacetylases, whereas SGA encodes a histone H3 chaperone/chromatin assembly protein also known as ASF1 in yeast and animals. CONCLUSIONS AND PROSPECTIVE In addition to its long-described history as a plant pathogen (Smith and Townsend, 1907), Agrobacterium is a natural genetic engineer that scientists have used for gene transfer experiments for the past 25 years. Whole-genome saturation mutagenesis studies in the early 1980s defined many Agrobacterium genes important for transformation, but only during the past decade have scientists applied modern genomic technologies to unravel the full complement of bacterial and host proteins important for transformation. In the near future, advances in molecular biology combined with novel imaging (e.g. Lee et al., 2008) and genetic (e.g. House et al., 2004) techniques will give scientists a considerably more refined view of bacterial and host proteins involved in transformation. This knowledge will likely result in improved transformation technologies, both to increase our ability to control Agrobacterium host range and to improve the quality (e.g. single-copy T-DNA insertions that result in predictable and stable transgene expression) of transformation events. Numerous important questions need to be answered to understand Agrobacterium-mediated plant genetic transformation more fully. Many of these questions beg genome-wide answers: (1) What roles do plant defense responses, and Agrobacterium's ability to overcome these responses, play in transformation? (2) How does the transferred VirD2/T-strand assemble with other virulence effector proteins and host proteins to traverse the plant cytoplasm and nucleus? (3) What roles do plant proteins play in T-strand targeting to plant chromatin and in T-DNA integration into the genome? Can we manipulate Agrobacterium for gene targeting (site-directed integration) purposes? (4) How can we best manipulate both the bacterium and the host to obtain high-quality transformation events? (5) How does Agrobacterium manipulate host metabolism for its advantage? (6) How does Agrobacterium interact with other organisms in the rhizosphere? (7) To what extent do the lessons we have learned about transformation using laboratory conditions apply to transformation in nature? (8) Has horizontal gene transfer effected by Agrobacterium species influenced plant evolution? These and other questions will likely be answered using genomic, proteomic, and metabolomic approaches. ACKNOWLEDGMENTS I thank Drs. Erh-Min Lai and Walt Ream for critical reading of this manuscript and for helpful suggestions. Research in the author's laboratory is funded by the National Science Foundation, the Department of Energy through the Corporation for Plant Biotechnology Research, the Biotechnology Research and Development Corporation, and Dow Agrosciences. 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Gelvin ([email protected]). www.plantphysiol.org/cgi/doi/10.1104/pp.109.139873 © 2009 American Society of Plant Biologists This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

White, Frank F.; Yang, Bing

doi: 10.1104/pp.109.139360pmid: 19458115

Rice (Oryza sativa) cultivation represents a world laboratory for investigation into bacterial diseases of rice, in particular, and host-parasite interactions, in general, at the molecular, genetic, and genomic levels. The crop, in its various forms, has been under intense cultivation for more than 6,000 years, resulting in the selection of a wide variety of traits and germplasm (Khush, 1997). Tremendous molecular and genetic resources have also been developed in recent years. The species can be genetically manipulated by Agrobacterium tumefaciens-mediated transformation (Krishnan et al., 2009). Against this backdrop, bacterial blight is an important disease of rice and present throughout much of the rice-growing regions (Ou, 1985; Mew et al., 1993; Fig. 1 Figure 1. Open in new tabDownload slide Bacterial blight and bacterial streak of rice. Left, Bacterial blight in an experimental field plot in Korea, 2006. Right, Bacterial streak from a test inoculation (courtesy of Dr. A. Bogdanove). Figure 1. Open in new tabDownload slide Bacterial blight and bacterial streak of rice. Left, Bacterial blight in an experimental field plot in Korea, 2006. Right, Bacterial streak from a test inoculation (courtesy of Dr. A. Bogdanove). ). Xanthomonas oryzae pv oryzae (Xoo) is the causal agent and a member of the γ-proteobacteria and, like many other proteobacteria, depends, in part, on a type III secretion system for pathogenicity (Swings et al., 1990; Zhu et al., 2000). The bacterium invades the xylem tissue, either through wounds or water pores, leading to systemic infection. Bacterial streak of rice is caused by the closely related pathovar X. oryzae pv oryzicola (Xoc), which, in contrast to Xoo, is limited to the intervascular regions, giving the disease the characteristic leaf streaking (Niño-Liu et al., 2006). Bacterial streak can also cause severe losses, although the disease tends to be tropical and is more sporadic than blight (Niño-Liu et al., 2006). This review will highlight recent advances in our knowledge of bacterial blight and, to a lesser extent, bacterial streak with regard to the host-bacteria interaction and the peculiar exploitation of a family of type III effectors. We will also provide speculation and thought as to how these findings might relate to classical analyses of disease resistance and susceptibility as well as a discussion of prospects for future research. RESISTANCE AND SUSCEPTIBILITY GENES TO XOO Bacterial blight is subject to control by genetic resistance, and rice has representatives of the two major classes of resistance genes (R genes) against the disease, which are given the prefix Xa for Xanthomonas (Table I Table I. Cloned R genes of rice Gene . Class . Comments (Product Localization; Resistance Profile; Function) . Cognate Elicitor/Effector . References . Xa21 RLK Extracellular, membrane, and intracellular domains; kinase; broad resistance Unknown small extracellular molecule Song et al. (1995), da Silva et al. (2004) Xa26 RLK Similar to Xa21; same locus as Xa3; broad resistance Unknown Sun et al. (2004), Xiang et al. (2006) Xa1 NBS-LRR Cytoplasm; narrow resistance Unknown Yoshimura et al. (1998) Rxo1 NBS-LRR (maize) Cytoplasm; broad resistance; transferred to rice from maize AvrRxo1 Zhao et al. (2004, 2005) Xa27 T3 TAL effector-inducible Membrane and cell wall; novel protein; broad resistance AvrXa27 Gu et al. (2005) xa5 Missense mutant of TFIIAγ5; small subunit of TFIIA transcription factor complex Nuclear; broad resistance Unknown Iyer and McCouch (2004), Jiang et al. (2006) xa13 Promoter mutants of Os8N3; nodulin 3 family Membrane; unresponsive to S gene to T3 effector PthXo1 Chu et al. (2006), Yang et al. (2006) Gene . Class . Comments (Product Localization; Resistance Profile; Function) . Cognate Elicitor/Effector . References . Xa21 RLK Extracellular, membrane, and intracellular domains; kinase; broad resistance Unknown small extracellular molecule Song et al. (1995), da Silva et al. (2004) Xa26 RLK Similar to Xa21; same locus as Xa3; broad resistance Unknown Sun et al. (2004), Xiang et al. (2006) Xa1 NBS-LRR Cytoplasm; narrow resistance Unknown Yoshimura et al. (1998) Rxo1 NBS-LRR (maize) Cytoplasm; broad resistance; transferred to rice from maize AvrRxo1 Zhao et al. (2004, 2005) Xa27 T3 TAL effector-inducible Membrane and cell wall; novel protein; broad resistance AvrXa27 Gu et al. (2005) xa5 Missense mutant of TFIIAγ5; small subunit of TFIIA transcription factor complex Nuclear; broad resistance Unknown Iyer and McCouch (2004), Jiang et al. (2006) xa13 Promoter mutants of Os8N3; nodulin 3 family Membrane; unresponsive to S gene to T3 effector PthXo1 Chu et al. (2006), Yang et al. (2006) Open in new tab Table I. Cloned R genes of rice Gene . Class . Comments (Product Localization; Resistance Profile; Function) . Cognate Elicitor/Effector . References . Xa21 RLK Extracellular, membrane, and intracellular domains; kinase; broad resistance Unknown small extracellular molecule Song et al. (1995), da Silva et al. (2004) Xa26 RLK Similar to Xa21; same locus as Xa3; broad resistance Unknown Sun et al. (2004), Xiang et al. (2006) Xa1 NBS-LRR Cytoplasm; narrow resistance Unknown Yoshimura et al. (1998) Rxo1 NBS-LRR (maize) Cytoplasm; broad resistance; transferred to rice from maize AvrRxo1 Zhao et al. (2004, 2005) Xa27 T3 TAL effector-inducible Membrane and cell wall; novel protein; broad resistance AvrXa27 Gu et al. (2005) xa5 Missense mutant of TFIIAγ5; small subunit of TFIIA transcription factor complex Nuclear; broad resistance Unknown Iyer and McCouch (2004), Jiang et al. (2006) xa13 Promoter mutants of Os8N3; nodulin 3 family Membrane; unresponsive to S gene to T3 effector PthXo1 Chu et al. (2006), Yang et al. (2006) Gene . Class . Comments (Product Localization; Resistance Profile; Function) . Cognate Elicitor/Effector . References . Xa21 RLK Extracellular, membrane, and intracellular domains; kinase; broad resistance Unknown small extracellular molecule Song et al. (1995), da Silva et al. (2004) Xa26 RLK Similar to Xa21; same locus as Xa3; broad resistance Unknown Sun et al. (2004), Xiang et al. (2006) Xa1 NBS-LRR Cytoplasm; narrow resistance Unknown Yoshimura et al. (1998) Rxo1 NBS-LRR (maize) Cytoplasm; broad resistance; transferred to rice from maize AvrRxo1 Zhao et al. (2004, 2005) Xa27 T3 TAL effector-inducible Membrane and cell wall; novel protein; broad resistance AvrXa27 Gu et al. (2005) xa5 Missense mutant of TFIIAγ5; small subunit of TFIIA transcription factor complex Nuclear; broad resistance Unknown Iyer and McCouch (2004), Jiang et al. (2006) xa13 Promoter mutants of Os8N3; nodulin 3 family Membrane; unresponsive to S gene to T3 effector PthXo1 Chu et al. (2006), Yang et al. (2006) Open in new tab ). Xa21 is an R gene introgressed into rice from the related species Oryza longistaminata and was the first R gene for bacterial blight as well as the first R gene of the receptor kinase (RLK) class to be cloned (Song et al., 1995). Xa21 confers resistance to many strains of Xoo, which is known as broad resistance (Ronald et al., 1992; Wang et al., 1996). RLKs have subsequently been shown to play a major role in a number of signaling pathways in plants, including innate immunity (Morillo and Tax, 2006). The prototypic RLK has a multidomain structure and includes an extracellular leucine-rich repeat (LRR) domain, which serves as the receptor for specific extracellular molecules, a transmembrane domain, and an intracellular kinase domain. Xa21D, a member of the multicopy locus that includes Xa21 and confers partial resistance, has only portions of the prototypic protein and likely functions through intermolecular interactions (Wang et al., 1998). Other, more recently cloned, members of the RLK family include Xa26 (and Xa3, the same locus by a different name), which also provides broad resistance to a somewhat different strain profile (Sun et al., 2004). RLK receptors, as a class, respond to a variety of molecules, both exogenous elicitors and endogenous signaling factors. With regard to innate immunity in rice, the molecules produced by Xoo and recognized by XA21 or XA26 have not been fully characterized. However, genetic analyses indicated that the elicitor is likely to be a sulfunated peptide that is secreted through a type I system (da Silva et al., 2004). In Arabidopsis, RLK receptors that signal innate immunity have been shown to respond to bacterial flagellin (FLS2) and elongation factor Tu (EFR; Gomez-Gomez and Boller, 2000; Zipfel et al., 2006). FLS2 and EFR were not formally identified as R genes but as components of the pathogen-associated molecular pattern-triggered immunity (PTI) response (Gomez-Gomez and Boller, 2002; Zipfel et al., 2004). Thus, Xa21, Xa26, and other RLKs are likely to represent genetic components of the PTI surveillance pathway of rice and related species. Xa1 represents the second major class of R genes, the nucleotide-binding site (NBS)-LRR group (Yoshimura et al., 1998). Xa1 expression is elevated upon bacterial infection and, as such, represents one of the cases in which R gene regulation is coordinated with other pathways for defense responses (Yoshimura et al., 1998; Zhang and Gassmann, 2007). Rxo1 from maize (Zea mays) is a member of the NBS-LRR gene family and confers broad resistance to Xoc isolates upon transfer to rice (Zhao et al., 2005). Rxo1-mediated resistance is triggered by strains containing the type III effector gene avrRxo1 from Xoc (Zhao et al., 2004). Although rice has relatively large numbers of NBS-LRR genes, Xa1 remains the only endogenous NBS-LRR gene identified for bacterial resistance (Monosi et al., 2004). Xa1 is effective against some Xoo isolates in Japan but is not effective against most strains from the Philippines. The gene for the elicitor signal from the pathogen has also not been identified for Xa1. Xa27 confers broad resistance and is representative of an unusual class of dominant R genes in rice (Gu et al., 2004). Upon analysis, Xa27 differed remarkably from other R genes in that specificity is based on differential gene expression (Gu et al., 2005). Identical open reading frames are present in resistant and susceptible cultivars but are only expressed in resistant cultivars upon bacterial infection. No function or conditional expression for the susceptible allele of Xa27 has been determined. The product of Xa27 is unrelated to any other class of R protein and has two related open reading frames on chromosome 6, although whether these related genes are functional is unknown. Xa27 is only expressed upon inoculation with Xoo strains harboring the type III effector gene avrXa27 (Gu et al., 2005). XA27 protein appears to be toxic to rice and initiates a resistant reaction upon expression (Gu et al., 2005; Tian and Yin, 2009). When fused to a pathogen-nonspecific-inducible rice OsPR1 promoter, Xa27 conferred resistance to compatible and incompatible strains of the pathogen alike. Localization studies of XA27 indicate that the protein is secreted into apoplastic space and associated with both the plant membrane and cell walls upon gene induction during resistance (Wu et al., 2008; Fig. 2 Figure 2. Open in new tabDownload slide Localization of XA27∷GFP. XA27 fused to GFP is found associated with cell plasmalemma and cell wall when cells are plasmolyzed (top). A GFP control is shown at bottom. A complete description of the approach is provided by Wu et al. (2008). P, Plasmolyzed plant cell. Arrows indicate adjoining plant cell wall. Photographs are courtesy of Z. Yin. Figure 2. Open in new tabDownload slide Localization of XA27∷GFP. XA27 fused to GFP is found associated with cell plasmalemma and cell wall when cells are plasmolyzed (top). A GFP control is shown at bottom. A complete description of the approach is provided by Wu et al. (2008). P, Plasmolyzed plant cell. Arrows indicate adjoining plant cell wall. Photographs are courtesy of Z. Yin. ). Two additional, as yet uncharacterized, R genes of rice have similar requirements for the transcription activation domain and nuclear localization signal motifs of the corresponding transcription activation-like (TAL) effectors as required for AvrXa27-mediated induction of Xa27 (Hopkins et al., 1992; Zhu et al., 1998; Yang et al., 2000). Cloning and analysis of these R genes will reveal whether the effectors and R gene expression have similar features to Xa27 (Porter et al., 2003; Chen et al., 2008; Gu et al., 2008). Two recessive R genes have been characterized from rice. The R gene xa5 confers broad resistance and, oddly, is an allele of the gene for the γ or small subunit of the transcription factor TFIIA (Iyer and McCouch, 2004; Jiang et al., 2006). The recessive allele is not a null allele but differs from the susceptible form by a single codon substitution of Val at position 39 to Glu. TFIIA, consisting of α-, β-, and γ-subunits, is involved in stabilizing the binding of the TATA box-binding protein complex (TFIID) to the TATA box of gene promoters (Hieb et al., 2007). The TFIIA components are highly conserved across the eukaryotes but are not required for transcription of all genes. Rice is unusual in comparison with human and yeast in having two loci for TFIIAγ (Iyer and McCouch, 2004). One gene is on chromosome 5 (TFIIAγ5, xa5) and the other is on chromosome 1 (TFIIAγ1). The proteins are closely related but not identical. The fact that the TFIIAγ5 message is present at greater levels based on hybridization analyses of leaf tissue indicates that TFIIAγ5 is likely the predominant form of the proteins in rice (Iyer and McCouch, 2004; Jiang et al., 2006; Sugio et al., 2007). The presence of two genes raises the question of whether or not rice or an ancestral relative adapted to bacterial infection by duplication of an ancestral gene for TFIIAγ. xa5 may provide effective resistance to bacterial infection while maintaining the necessary functionality for normal rice gene expression under most circumstances. TFIIAγ1 may be needed under some other, as yet unknown, conditions or developmental state, and maintenance of a second gene may have a fitness advantage. However, no evidence has been reported that indicates that xa5 has a detrimental effect under any field or experimental conditions. Additional recessive rice R genes await characterization and may prove as equally fascinating as xa5 (Iyer-Pascuzzi and McCouch, 2007). Another recessive resistance gene, xa13, has been identified by map-based cloning (Chu et al., 2005, 2006). Like xa5, the recessive allele is not a null mutant and encodes a protein related to MtN3, encoding nodulin 3 (N3) protein of Medicago truncatula. The MtN3 gene was first identified in EST libraries of M. truncatula root nodules (Gamas et al., 1996). The dominant allele Xa13 was also named Os8N3 due to the location on rice chromosome 8 and the similarity to MtN3, and that name is used throughout this review (Yang et al., 2006). Os8N3 is a member of a moderately large gene family consisting of 17 to 20 members in rice. Arabidopsis (Arabidopsis thaliana) also has an 18-member MtN3 family, and the family is also conserved across kingdom boundaries, with related genes in mammals, insects, nematodes, and filamentous fungi, although not to the same degree as TFIIAγ (Chu et al., 2006; Yang et al., 2006; Guan et al., 2008; B. Yang and F.F. White, unpublished data). The critical difference between resistant (xa13/xa13) and susceptible plants is the elevated expression of Os8N3 during bacterial infection in susceptible plant genotypes and the absence of Os8N3 induction during bacterial infection in the resistant genotypes (Yang et al., 2006; Yuan et al., 2009). Knockdown of Os8N3 expression by inhibitory RNA also results in resistance to some bacterial strains, but, unlike xa13 genotypes, silenced plants have low pollen viability and reduced numbers of viable seeds (Chu et al., 2006; Yang et al., 2006). Thus, xa13 confers resistance due to a lack of elevated Os8N3 expression during bacterial infection. Os8N3, therefore, is a susceptibility (S) gene that is exploited by Xoo, and the xa13 resistance is a naturally occurring allele, actually a series of alleles that “fix” a genetic disease vulnerability in the plant developmental pathways (Yang et al., 2006). Bacterial strains that induce Os8N3 expression are virulent, and those strains that normally induce Os8N3 but not the xa13 allele are incompatible. The xa13 gene appears to have been a valuable allele in rice cultivation, as the resistance has occurred in various forms in a variety of cultivars (Chu et al., 2005). The simplest variant is present in ‘BJ1’, where the difference between the resistant and susceptible alleles is a single nucleotide change in the promoter region of the gene. Other variants have more apparent complex alterations, including multiple base changes and/or insertions in the Os8N3 promoter region (Chu et al., 2006). Further expression analysis of Os8N3 and the various xa13 alleles will provide important insights into the intricacy of host genes that have adapted for disease resistance while maintaining normal developmental processes. Despite the number of alleles, xa13 is not as broad as the resistance provided by Xa21, Xa27, and xa5, and many strains from China, the Philippines, Japan, and Korea are compatible on xa13 lines (Yang et al., 2006). The number of strains that are compatible on plants containing xa13 illustrates that Xoo can and has adapted to xa13 alleles as well as other R genes in the host. Studies of the basis of compatibility and incompatibility between rice genotypes and strains of the pathogen have been conducted in the hope that insight may be gained into the factors that affect R gene breadth and durability, which here is defined loosely as the time the R gene is effective against extant populations of pathogens. TYPE III EFFECTORS IN XOO Despite all the resources, only two pathogen genes for elicitors that correspond to cloned R genes for bacterial blight or streak of rice have been cloned: avrXa27 from Xoo as the cognate elicitor gene for Xa27 and, if the heterologous Rxo1 gene from maize is included, avrRxo1 from Xoc (Table II Table II. Type III effectors of Xoo and Xoc with phenotypic effects in rice T3 Effector . Strain . Cognate Host Gene(s) . References . Additional Commentsa b . AvrRxo1 (Xoc) BLS256 Rxo1 (maize) Zhao et al. (2004) No virulence activity AvrXa7 PXO86 Xa7 Hopkins et al. (1992) Major TAL effector; present also in T7174 and KXO85; Xa7-dependent avirulence activity AvrXa7-Δ38 PXO99a Xa7 Yang et al. (2005) Derivative of AvrXa7 with no virulence activity AvrXa7-sacB50 PXO99a New unnamed R gene Yang et al. (2005) Derivative of AvrXa7 with no virulence or Xa7-dependent avirulence activity AvrXa10 PXO86 Xa10 Hopkins et al. (1992) No virulence activity AvrXa27 PXO99a Xa27 Gu et al. (2005) No virulence activity PthXo1 PXO99a Os8N3 Yang and White (2004), Yang et al. (2006) Present also in PXO71 PthXo2 JOX1 Unknown Yang and White (2004) Major TAL effector; also present in PXO71 and T7174 PthXo3 PXO61 Unknown Yang and White (2004) Major TAL effector PthXo4 PXO99a Unknown Yang et al. (2005) Derivative of AvrXa7 PthXo5 PXO99a Unknown Yang et al. (2005) Derivative of AvrXa7 PthXo6 PXO99a OsTFX1 Sugio et al. (2007) Virulence activity in all strains tested PthXo7 PXO99a OsTFIIAγ1 Sugio et al. (2007) Unique to PXO99a; weak virulence activity in host with xa5 recessive resistance PthXo8 PXO99a Unknown B. Yang and F.F. White (unpublished data) Virulence activity T3 Effector . Strain . Cognate Host Gene(s) . References . Additional Commentsa b . AvrRxo1 (Xoc) BLS256 Rxo1 (maize) Zhao et al. (2004) No virulence activity AvrXa7 PXO86 Xa7 Hopkins et al. (1992) Major TAL effector; present also in T7174 and KXO85; Xa7-dependent avirulence activity AvrXa7-Δ38 PXO99a Xa7 Yang et al. (2005) Derivative of AvrXa7 with no virulence activity AvrXa7-sacB50 PXO99a New unnamed R gene Yang et al. (2005) Derivative of AvrXa7 with no virulence or Xa7-dependent avirulence activity AvrXa10 PXO86 Xa10 Hopkins et al. (1992) No virulence activity AvrXa27 PXO99a Xa27 Gu et al. (2005) No virulence activity PthXo1 PXO99a Os8N3 Yang and White (2004), Yang et al. (2006) Present also in PXO71 PthXo2 JOX1 Unknown Yang and White (2004) Major TAL effector; also present in PXO71 and T7174 PthXo3 PXO61 Unknown Yang and White (2004) Major TAL effector PthXo4 PXO99a Unknown Yang et al. (2005) Derivative of AvrXa7 PthXo5 PXO99a Unknown Yang et al. (2005) Derivative of AvrXa7 PthXo6 PXO99a OsTFX1 Sugio et al. (2007) Virulence activity in all strains tested PthXo7 PXO99a OsTFIIAγ1 Sugio et al. (2007) Unique to PXO99a; weak virulence activity in host with xa5 recessive resistance PthXo8 PXO99a Unknown B. Yang and F.F. White (unpublished data) Virulence activity a Virulence activity indicates contribution to virulence based on lesion length and bacterial population on the host plant. Avirulence activity indicates ability to elicit a resistance reaction on appropriate genotype of the host. b Major TAL effectors are defined by their loss due to mutation. Loss of major TAL effector gene, if the only one present in a strain, results in more than 80% virulence as measured by lesion length when compared with the wild type. Open in new tab Table II. Type III effectors of Xoo and Xoc with phenotypic effects in rice T3 Effector . Strain . Cognate Host Gene(s) . References . Additional Commentsa b . AvrRxo1 (Xoc) BLS256 Rxo1 (maize) Zhao et al. (2004) No virulence activity AvrXa7 PXO86 Xa7 Hopkins et al. (1992) Major TAL effector; present also in T7174 and KXO85; Xa7-dependent avirulence activity AvrXa7-Δ38 PXO99a Xa7 Yang et al. (2005) Derivative of AvrXa7 with no virulence activity AvrXa7-sacB50 PXO99a New unnamed R gene Yang et al. (2005) Derivative of AvrXa7 with no virulence or Xa7-dependent avirulence activity AvrXa10 PXO86 Xa10 Hopkins et al. (1992) No virulence activity AvrXa27 PXO99a Xa27 Gu et al. (2005) No virulence activity PthXo1 PXO99a Os8N3 Yang and White (2004), Yang et al. (2006) Present also in PXO71 PthXo2 JOX1 Unknown Yang and White (2004) Major TAL effector; also present in PXO71 and T7174 PthXo3 PXO61 Unknown Yang and White (2004) Major TAL effector PthXo4 PXO99a Unknown Yang et al. (2005) Derivative of AvrXa7 PthXo5 PXO99a Unknown Yang et al. (2005) Derivative of AvrXa7 PthXo6 PXO99a OsTFX1 Sugio et al. (2007) Virulence activity in all strains tested PthXo7 PXO99a OsTFIIAγ1 Sugio et al. (2007) Unique to PXO99a; weak virulence activity in host with xa5 recessive resistance PthXo8 PXO99a Unknown B. Yang and F.F. White (unpublished data) Virulence activity T3 Effector . Strain . Cognate Host Gene(s) . References . Additional Commentsa b . AvrRxo1 (Xoc) BLS256 Rxo1 (maize) Zhao et al. (2004) No virulence activity AvrXa7 PXO86 Xa7 Hopkins et al. (1992) Major TAL effector; present also in T7174 and KXO85; Xa7-dependent avirulence activity AvrXa7-Δ38 PXO99a Xa7 Yang et al. (2005) Derivative of AvrXa7 with no virulence activity AvrXa7-sacB50 PXO99a New unnamed R gene Yang et al. (2005) Derivative of AvrXa7 with no virulence or Xa7-dependent avirulence activity AvrXa10 PXO86 Xa10 Hopkins et al. (1992) No virulence activity AvrXa27 PXO99a Xa27 Gu et al. (2005) No virulence activity PthXo1 PXO99a Os8N3 Yang and White (2004), Yang et al. (2006) Present also in PXO71 PthXo2 JOX1 Unknown Yang and White (2004) Major TAL effector; also present in PXO71 and T7174 PthXo3 PXO61 Unknown Yang and White (2004) Major TAL effector PthXo4 PXO99a Unknown Yang et al. (2005) Derivative of AvrXa7 PthXo5 PXO99a Unknown Yang et al. (2005) Derivative of AvrXa7 PthXo6 PXO99a OsTFX1 Sugio et al. (2007) Virulence activity in all strains tested PthXo7 PXO99a OsTFIIAγ1 Sugio et al. (2007) Unique to PXO99a; weak virulence activity in host with xa5 recessive resistance PthXo8 PXO99a Unknown B. Yang and F.F. White (unpublished data) Virulence activity a Virulence activity indicates contribution to virulence based on lesion length and bacterial population on the host plant. Avirulence activity indicates ability to elicit a resistance reaction on appropriate genotype of the host. b Major TAL effectors are defined by their loss due to mutation. Loss of major TAL effector gene, if the only one present in a strain, results in more than 80% virulence as measured by lesion length when compared with the wild type. Open in new tab ). Both genes encode substrate proteins that are secreted through the bacterial type III secretion system (T3SS) and are generally known as type III (T3) effectors, which are fully reviewed in this issue. A functioning T3SS is required for pathogenicity of both Xoo and Xoc and serves to secrete multiple T3 effectors into the host cells. Although not all are associated with phenotypic effects for virulence, T3 effectors function, in general, as virulence factors in pathogenicity. A subset of the T3 effectors, including AvrXa27 and AvrRxo1, can serve as the cognate elicitors (avirulence proteins) for specific R genes in many plant systems. Two additional cognate T3 effector (avirulence) genes that have been cloned are avrXa7 and avrXa10, corresponding to the R genes Xa7 and Xa10. Five additional T3 effectors from Xoo have known contributions to virulence under the appropriate conditions (Table II). These genes are pthXo1, pthXo2, pthXo3, pthXo6, and pthXo7. The gene avrXa7, in addition to Xa7-dependent elicitor activity, is also a virulence factor. Four additional T3 effectors were derived under a laboratory setting from avrXa7 (Table II; Yang et al., 2005). Two genes, named pthXo4 and pthXo5, have lost Xa7-dependent elicitor activity while maintaining virulence activity. A third gene, avrXa7-Δ38, retained Xa7-dependent activity without the accompanying virulence activity, and a fourth, avrXa7-sacB50, appears to have lost both avirulence and virulence activity but acquired avirulence activity on the otherwise susceptible rice line IR24. The new incompatible response was not observed on the rice line Nipponbare. Preliminary results indicate that a sixth gene, pthXo8, also contributes to the virulence of Xoo (B. Yang, unpublished data). With the exception of avrRxo1, all of the above T3 effectors are related to avrBs3 and pthA, which were first identified in X. campestris pv vesicatoria and X. campestris pv citri, the causal agents of bacterial spot of pepper (Capsicum annuum) and citrus (Citrus species) canker, respectively (Bonas et al., 1989; Swarup et al., 1991). The AvrBs3/PthA-related T3 effectors have properties of transcription factors and possess eukaryotic nuclear localization signal motifs and a potent acidic transcription activation domain in the C-terminal portion (Van den Ackerveken et al., 1996; Zhu et al., 1998). The proteins have DNA-binding activity, and AvrBs3 can bind specific gene promoter sequences in vivo, as shown by chromatin immunoprecipitation and gel shift assays (Yang et al., 2000; Kay et al., 2007). Here, we refer to the family of T3 effectors as the TAL T3 effectors, and each protein has a series of highly conserved repeated sequence of amino acids (Fig. 3 Figure 3. Open in new tabDownload slide Prototypic TAL effector. Each TAL effector contains a central repetitive region of varying numbers of 34 amino acid repeat units (white boxes). The C-terminal region contains three nuclear localization signal motifs (NLS [A, B, and C]) and an acidic activation domain (AD). Figure 3. Open in new tabDownload slide Prototypic TAL effector. Each TAL effector contains a central repetitive region of varying numbers of 34 amino acid repeat units (white boxes). The C-terminal region contains three nuclear localization signal motifs (NLS [A, B, and C]) and an acidic activation domain (AD). ). The consensus repeat, in Xoo and Xoc, is 34 amino acid residues, although some repeats have single codon deletions or internal smaller repetitive portions. With some exceptions, the central repetitive region can be switched among the TAL effector family members while retaining the biological activity of the effector from which the repeat region is derived. Thus, some aspect of TAL effector specificity is dictated by the repetitive structure. The repeats may affect domain spacing as well as, perhaps, the maintenance of individual effector identity. The proteins are believed to function as dimeric proteins within the host (Gurlebeck et al., 2005). The closest structural relative of the repeat region is the so-called tetratricopeptide repeat, which is also approximately 34 amino acid residues in length and widespread in proteins of diverse function (Blatch and Lassle, 1999). The AvrRxo1, on the other hand, is a novel protein of 421 amino acids with a variety of reported motifs, including a eukaryotic thiol protease active site, an ATP/GTP-binding site motif, nine N-myristoylation site motifs, and a putative nuclear localization sequence motif (Zhao et al., 2004). The functionality of the motifs has not yet been experimentally verified. Genes related to avrRxo1 are fairly limited in distribution and present in Acidovorax, Burkholdaria, and X. campestris pv vesicatoria (Zhao et al., 2004; Thieme et al., 2005). However, related genes have not been detected in Xoo. Mutants of avrRxo1 have not been generated; therefore, any virulence contribution to Xoc by avrRxo1 is unknown. IMPLICATION OF INTERACTION BETWEEN T3 EFFECTORS AND THE CORRESPONDING HOST GENES Xa27 and xa13 have provided insight into the diversity of resistance mechanisms in rice. The apparent reason for the broad activity of Xa27 is the presence of avrXa27 in a large number of strains from southeast Asia, including many strains from Korea, China, Japan, and the Philippines (Gu et al., 2004). At the same time, two lines of evidence indicate that strains of Xoo could defeat Xa27. First, avrXa27 is present in a single copy, at least in the two strains that have been examined; therefore, simple loss or mutation of the gene converts incompatible strains to compatible strains on plants with Xa27 (Gu et al., 2005; Ochiai et al., 2005). Second, mutation of avrXa27 in the strain PXO99A converts PXO99A to compatibility on plants containing Xa27 and does not lead to any detectible loss of virulence. Thus, the loss of avrXa27 does not appear to have a fitness cost to the pathogen, and populations of Xoo without avrXa27 could be expected to appear relatively rapidly should Xa27 be deployed. avrXa7, unlike avrXa27, is an important virulence factor for some strains of Xoo, and loss of avrXa7 can result in strains that are only weakly virulent (Bai et al., 2000; Yang et al., 2000). Several factors mitigate the argument that Xa7, by itself, would provide durable resistance due to the fitness cost for loss of avrXa7 in the pathogen. One factor is that a variety of other TAL T3 effector genes are present in Xoo populations that do not have Xa7-mediated elicitor activity but can restore full virulence to strains missing avrXa7 (Yang and White, 2004). We refer to interchangeable TAL effectors with regard to virulence activity as major TAL effectors. In addition to avrXa7, the major TAL effector genes in Xoo are pthXo1, pthXo2, and pthXo3 (Yang and White, 2004). Loss of Xa7-dependent elicitor activity from avrXa7 and concomitant retention of virulence activity has also been observed (Yang et al., 2005). Thus, evasion of Xa7-mediated resistance should, at least theoretically, be possible by deletion of the central repeats, as observed for pthXo4, by recombination among different TAL effector genes, as observed with pthXo5, or by horizontal transfer of alternate virulence factors (Yang et al., 2005). At the same time, field studies in the Philippines indicated that deployment of Xa7 was durable in test plots for more than 10 years (Vera Cruz et al., 2000). Regardless of the possibility of race changes in the pathogen population, pyramiding R genes with cognate T3 effectors that are widespread in the pathogen populations should provide a degree of broad durable resistance. TAL T3 effectors also are involved in recessive resistance. In the case of xa13, induction of the dominant allele Os8N3 is mediated by the TAL effector PthXo1 (Yang et al., 2006). The gene pthXo1 was first identified in Xoo strain PXO99A, which coincidentally is also a strain that is incompatible on plants that are homozygous for xa13 (Yang and White, 2004). PXO99A cannot induce the xa13 allele of Os8N3 and does not grow as well in xa13 plants in comparison with normal plants (Chu et al., 2005; Yang et al., 2006). The introduction of avrXa7, pthXo2, or pthXo3 into PXO99A restores virulence regardless of the presence of pthXo1 (Yang et al., 2006). Thus, xa13-dependent resistance is only effective against strains that rely on pthXo1 and is mimicked phenotypically by mutations in pthXo1 of PXO99A (Yang et al., 2006). The recessive resistance provided by xa13 is phenotypically and qualitatively different from resistance provided by the dominant R gene Xa7, for example (Chu et al., 2004; Yang et al., 2006). Quantitatively, in terms of bacterial growth and lesion length, resistance mediated by xa13 and Xa7 are approximately equal (Bai et al., 2000; Yang et al., 2000, 2006). However, Xa7 resistance is the result of the presence of the appropriate elicitor gene in the pathogen, while xa13 is dependent on the absence of an effective virulence factor (Fig. 4 Figure 4. Open in new tabDownload slide Diverse host responses to Xoo challenge in rice. Leaves were inoculated with water or Xoo by needleless syringe. Leaf 1, Mock inoculation; leaf 2, response to T3SS nonpathogenic mutant of PXO99A; leaf 3, recessive R gene response (note the lack of streaking up and down the leaf blade); leaf 4, fully susceptible response (wild-type strain); leaf 5, PXO99A (avrXa7; response to dominant R gene Xa7). Photographs were taken with back lighting, and yellow zones are indicative of water soaking. Figure 4. Open in new tabDownload slide Diverse host responses to Xoo challenge in rice. Leaves were inoculated with water or Xoo by needleless syringe. Leaf 1, Mock inoculation; leaf 2, response to T3SS nonpathogenic mutant of PXO99A; leaf 3, recessive R gene response (note the lack of streaking up and down the leaf blade); leaf 4, fully susceptible response (wild-type strain); leaf 5, PXO99A (avrXa7; response to dominant R gene Xa7). Photographs were taken with back lighting, and yellow zones are indicative of water soaking. ). The xa13 alleles are natural cases of mutations in loss of function for susceptibility, hence the classification of the dominant allele Os8N3 as a S gene of the host and the target of the virulence factor PthXo1. The defeat of xa13-mediated resistance could theoretically involve induction of the xa13 allele or targeting of alternative host genes that provide the same effect as Os8N3. Some variants of xa13 are weakly inducible in young plants and concomitantly have greater susceptibility to PXO99A as young plants (B. Yang, unpublished data). No strains have been identified that induced the xa13 allele similar to PthXo1-mediated induction of Os8N3, nor have strains with alternate virulence TAL effectors been found that induce Os8N3 in normal plants (Yang et al., 2006). The three aforementioned TAL effectors that restore virulence to PXO99A compensate for PthXo1 by targeting alternative S genes (B. Yang and F.F. White, unpublished data). PthXo1, AvrXa7, PthXo2, and PthXo3 are major virulence T3 effectors in the bacterial blight system, and the targeted host genes are major S genes. Strains lacking at least one of the major T3 effectors are severely debilitated for virulence, as measured by the standard leaf-clipping assays, display similar phenotypes, and, as noted above, compare favorably to results with dominant resistance genes. At present, the reason for resistance or lack of virulence is unclear and undoubtedly related to the benefit to the pathogen that is provided by the expression of the N3 gene in rice. T3 effectors, in general, are hypothesized to interfere with host defense and defense signaling mechanisms. Of course, in the broadest definition, any factor that promotes virulence can be interpreted as an interference with host defense. The question remains whether the major TAL T3 effectors of Xoo are “wrecking balls” and interfere with many normal host functions by expropriating normal developmental pathways or “guided missiles” designed to interfere with a specific host function related to immunity or, alternatively, beneficial functions unrelated specifically to immunity. One model for TAL effectors has proposed that the effectors enhance the spread of the bacteria from local infection sites (Yang et al., 1994; Iyer-Pascuzzi et al., 2008). Os8N3 protein, which is membrane localized, may, for example, result in loss of tissue integrity, allowing release of the bacteria into uninfected tissue (Chu et al., 2006). Alternatively, Os8N3 may promote greater bacterial growth due to leakage of nutrients into extracellular spaces, and consequently, the bacteria may grow to higher titers within the host. Future experimentation regarding bacterial physiology within the host as well as studies involving ectopic expression of S genes will provide insight into these models or provide new hypotheses to test. Xoo strains lacking the major TAL effectors are not nonpathogenic and are still capable of causing water-soaked symptoms if syringe inoculated. The ability to cause water soaking is in contrast to T3SS mutants, which are incapable of secretion of any T3SS effectors. T3SS-deficient mutants are virtually symptomless (Fig. 4). Utilization of the N3 family, based on existing evidence, is unique to the bacterial blight system. Xoc also contains multiple TAL effector genes, but infection does not lead to Os8N3 expression, indicating that, if the effectors play similar roles in heterologous systems, the targets of the effectors are likely to be different (A. Bogdanove, B. Yang, and F.F. White, unpublished data). The genomic sequences are available for three strains of Xoo and one strain of Xoc (Lee et al., 2005; Ochiai et al., 2005; Lu et al., 2008; Salzberg et al., 2008). The full complement of TAL T3 effectors are now known for two strains of Xoo and one strain of Xoc, and both pathogens are unusual due to the high number of TAL effector genes within their individual genomes. Xoo strains PXO99A and MAFF311018 (also named T7174) have 19 and 17 genes, respectively, organized in nine and eight regions of their respective genomes (Ochiai et al., 2005; Salzberg et al., 2008; Fig. 5 Figure 5. Open in new tabDownload slide Genomic positions of TAL effector genes in two strains of Xoo. Genes are labeled and identified by biological function and name, where assigned (adapted from Salzberg et al. [2008] and Ochiai et al. [2005]). The dashed line indicates the location of the PXO99A TAL effector gene 9e in MAFF3113018. Apostrophes after gene letters indicate truncated genes. Colored boxes indicate gene clusters. [See online article for color version of this figure.] Figure 5. Open in new tabDownload slide Genomic positions of TAL effector genes in two strains of Xoo. Genes are labeled and identified by biological function and name, where assigned (adapted from Salzberg et al. [2008] and Ochiai et al. [2005]). The dashed line indicates the location of the PXO99A TAL effector gene 9e in MAFF3113018. Apostrophes after gene letters indicate truncated genes. Colored boxes indicate gene clusters. [See online article for color version of this figure.] ). The individual genes are distinguishable on the basis of the number of repeats in the central repetitive region and by polymorphisms within each repeat sequence, particularly at the 12th and 13th codons. A comparison of the repetitive regions of PXO99A and MAFF311018 indicates a high degree of rearrangements and shuffling of the genes at all of the loci, to the point where only three genes of 17 in MAFF311018 are identical (Salzberg et al., 2008). The large numbers of TAL effector genes in these species may reflect the evolutionary “investment” the strains have in utilizing the TAL effectors for virulence. The major contribution to virulence by pthXo1, for example, corroborates the view that the genes, as a class of T3 effectors, are important, maybe essential, to the ecological niche these bacteria occupy. Yang and Gabriel (1995) proposed that maintenance of a large repertoire of TAL effector genes may increase the frequency of recombination between genes and, as a consequence, increase the diversity of TAL effectors within the pathogen population. Populations with greater diversity of effector genes may then adapt faster to changing host genotypes. The maintenance of high gene numbers may even be exacerbated by rice breeding and R gene deployment by farmers over the millennia. The gene pthXo5, for example, gained virulence function by recombination of the original avrXa7 and a chromosomal gene of PXO99A, now known to be pthXo6 (Yang et al., 2005; Sugio et al., 2007). Some of the genes are apparent pseudogenes, in the sense that a number of the genes are truncated in comparison with known functional genes (Salzberg et al., 2008). Pseudogenes could still serve as recombination substrates for generating new effector genes. Remarkably, Xoc strain BLS256 has 28 TAL effector genes, and the evidence also indicates that none are identical and most are unique with respect to repetitive region sequence. The profiles of changes in host gene expression upon infection by Xoc also appear different (A. Bogdanove, unpublished data). Differences in TAL effector gene effects have been observed between Xoo and Xoc, indicating possible differences in TAL effector utilization for host adaptation (Makino et al., 2006). While many may serve as recombination substrates, the TAL effector genes of PXO99A are not simply substrates for new major TAL effectors. Three TAL effector genes, in addition to pthXo1, are known to have contributions to virulence (Table II), and two are known to be associated with the elevated expression of two host genes distinct from Os8N3. The gene pthXo6 is responsible for the elevated expression of a gene named OsTFX1 (AK108319) and located on chromosome 9 of rice. A mutant of PXO99A in pthXo6 suffered a loss of approximately 35% in the lengths of leaf lesions and a 50% reduction in bacterial population per leaf (Sugio et al., 2007). Ectopic expression of OsTFX1 in rice abrogated the need for pthXo6 in the pathogen but did not make the plants more susceptible to PXO99A (Sugio et al., 2007). OsTFX1 is a member of the large bZip family of transcription factors, and the function of OsTFX1 in a normal plant is unknown. Ectopic expression of the gene, other than compensating for the presence of pthXo6 in the pathogen, did not cause any phenotypic abnormalities in the transgenic plants (Sugio et al., 2007). Elevated expression of OsTFX1 may provide a good diagnostic tool for Xoo, as all strains examined were capable of inducing the gene. Xoc, at least on the basis of a single strain, was unable to induce OsTFX1. pthXo7 is associated with the elevation of OsTFIIAγ1 and has only been found in Xoo strain PXO99A (Sugio et al., 2007). Loss of pthXo7 in PXO99A has no measurable effect on strain virulence in susceptible rice plants. In addition to compatibility on plants with the recessive xa13, PXO99A is also one of the few strains known to be compatible with the other recessive resistance gene xa5. One model for the resistance provided by xa5 is that the alternate TFIIAγ xa5 may not function properly with the TAL effectors and interfere with TAL effector function. In turn, PXO99A may have adapted, in part, to xa5 by boosting the level of TFIIAγ1. A small effect on virulence was found upon transfer to a strain (PXO86) that is incompatible on plants with xa5, suggesting that pthXo7 may be an adaptation to host genotypes containing the xa5 allele of TFIIAγ5 (Sugio et al., 2007). However, pthXo7 does not confer on PXO86 the same level of growth on xa5 plants in comparison with PXO99A; therefore, it does not appear to be the sole or even the principal reason for the ability of PXO99A to cause significant disease in the presence of xa5 (Sugio et al., 2007). It should also be noted that the growth of PXO99A is reduced in plants with the xa5 gene for resistance, which again points to the uniqueness of the xa5 mechanism for resistance. One also cannot, at this point, rule out the possibility of virulence factors and R genes in rice that are adapted specifically to the TFIIAγ xa5 subunit. In the future, xa5 may be considered not only as an R gene, because it also has characteristics of a quantitative trait locus and a resistance modifier/suppressor gene. Research into the molecular basis of Xoo and Xoc interaction with the host promises to provide exciting new insights in the near future into the adaptations between pathogen and host. The full complement of host effects due to the T3 TAL effectors wait to be discovered as more genes, strains, and host genotypes are examined. More recently, the characterization of strains from West Africa has revealed new genotypes of both Xoo and Xoc and indicates that an ability to cause bacterial blight of rice may have arisen in at least three lineages of related bacteria (Gonzalez et al., 2007). The research has also uncovered a variety of novel R genes, and a further understanding of their function in host susceptibility and resistance will be forthcoming. As in other bacterial pathogen-host systems, the features of a host strategy to turn the mechanism of the virulence factors against the pathogen are beginning to emerge. Curiously, in the rice system, the pathogen gains advantage by expropriating the developmental control of host genes and subsequent induction of a membrane protein, in the case of Os8N3. The host, on the other hand, can foil the process by production of a separate membrane protein (XA27) that triggers a rapid defense response. The differential effects of the proteins and the components of the two consequences on host cell physiology await further investigations. XA27, on the basis of sequence relatedness, is unique to rice and, possibly, some relatives (Gu et al., 2004), and it is one of a number of genes that may have been “invented” through the evolutionary process for specific purposes (Xiao et al., 2009). Our understanding of Xoo and Xoc virulence is far from complete. For example, a third TAL effector gene, named pthXo8, has been identified in PXO99A with quantitative effects similar to pthXo6, and the host gene expression associated with pthXo8 is under investigation. Preliminary evidence indicates that the effector is involved in manipulation of the small RNA pathways of the host. The curious and novel nature of the TAL T3 effectors sometimes distracts attention from the presence of multiple other T3 effectors in Xoo and Xoc genomes. Recent characterization of the genomic data from MAFF311018 indicates that at least 19 candidate T3 effector genes, in addition to the 17 TAL effector genes, are present in the genome (Furutani et al., 2009; Table III Table III. Candidate non-TAL type III effectors of Xoo Modified from Furutani et al. (2009). Entries in the same row are related by sequence similarity. N, No similar sequence identified; NA, not annotated. Xoo . . . X. axonopodis pv citri 306 . X. campestris pv vesicatoria 85-10 . MAFF311018a . KACC10331 . PXO99a . . . XOO0037 (novel) NA PXO03356 N N XOO0103 (XopF) XOO0074 PXO03413 XAC2785 XCV0414 XCV2942 XOO0148 (AvrBs2) XOO0168 PXO03330 XAC0076 XCV0052 XOO0315 (XopN) XOO0343 PXO02760 XAC2786 XCV2944 XOO3150 (XopN) NA NA XAC2786 XCV2944 XOO1488 (novel) NA NA N N XOO1669 (conserved) XOO1768 PXO01625 XAC3085 XCV3215 XOO2210 (conserved) XOO4824 N NA N XOO2877 (novel) NA PXO00236 N N XOO3803 (conserved) XOO4033 PXO04172 XAC0601 XCV0657 XOO4134 (conserved) XOO4391 PXO03819 XAC0277 XCV0285 XOO2402 (HopAS) XOO2543 PXO01041 XAC2009 XCC2059 XOO3222 (XopP) XOO3425 PXO02107 XAC1208 XCV1236 XOO3426 XOO2875 (XopX, HopAE) XOO3022 PXO00234 XAC0543 XCV3785 XOO4042 XOO4287 PXO03702 XCV0572 XOO4208 (XopQ) XOO4466 PXO03901 XAC433 XCV4438 Xoo . . . X. axonopodis pv citri 306 . X. campestris pv vesicatoria 85-10 . MAFF311018a . KACC10331 . PXO99a . . . XOO0037 (novel) NA PXO03356 N N XOO0103 (XopF) XOO0074 PXO03413 XAC2785 XCV0414 XCV2942 XOO0148 (AvrBs2) XOO0168 PXO03330 XAC0076 XCV0052 XOO0315 (XopN) XOO0343 PXO02760 XAC2786 XCV2944 XOO3150 (XopN) NA NA XAC2786 XCV2944 XOO1488 (novel) NA NA N N XOO1669 (conserved) XOO1768 PXO01625 XAC3085 XCV3215 XOO2210 (conserved) XOO4824 N NA N XOO2877 (novel) NA PXO00236 N N XOO3803 (conserved) XOO4033 PXO04172 XAC0601 XCV0657 XOO4134 (conserved) XOO4391 PXO03819 XAC0277 XCV0285 XOO2402 (HopAS) XOO2543 PXO01041 XAC2009 XCC2059 XOO3222 (XopP) XOO3425 PXO02107 XAC1208 XCV1236 XOO3426 XOO2875 (XopX, HopAE) XOO3022 PXO00234 XAC0543 XCV3785 XOO4042 XOO4287 PXO03702 XCV0572 XOO4208 (XopQ) XOO4466 PXO03901 XAC433 XCV4438 a Previously identified candidate effectors are in parentheses. Conserved, Newly identified candidate effector and conserved in other species; novel, unique to Xoo. Open in new tab Table III. Candidate non-TAL type III effectors of Xoo Modified from Furutani et al. (2009). Entries in the same row are related by sequence similarity. N, No similar sequence identified; NA, not annotated. Xoo . . . X. axonopodis pv citri 306 . X. campestris pv vesicatoria 85-10 . MAFF311018a . KACC10331 . PXO99a . . . XOO0037 (novel) NA PXO03356 N N XOO0103 (XopF) XOO0074 PXO03413 XAC2785 XCV0414 XCV2942 XOO0148 (AvrBs2) XOO0168 PXO03330 XAC0076 XCV0052 XOO0315 (XopN) XOO0343 PXO02760 XAC2786 XCV2944 XOO3150 (XopN) NA NA XAC2786 XCV2944 XOO1488 (novel) NA NA N N XOO1669 (conserved) XOO1768 PXO01625 XAC3085 XCV3215 XOO2210 (conserved) XOO4824 N NA N XOO2877 (novel) NA PXO00236 N N XOO3803 (conserved) XOO4033 PXO04172 XAC0601 XCV0657 XOO4134 (conserved) XOO4391 PXO03819 XAC0277 XCV0285 XOO2402 (HopAS) XOO2543 PXO01041 XAC2009 XCC2059 XOO3222 (XopP) XOO3425 PXO02107 XAC1208 XCV1236 XOO3426 XOO2875 (XopX, HopAE) XOO3022 PXO00234 XAC0543 XCV3785 XOO4042 XOO4287 PXO03702 XCV0572 XOO4208 (XopQ) XOO4466 PXO03901 XAC433 XCV4438 Xoo . . . X. axonopodis pv citri 306 . X. campestris pv vesicatoria 85-10 . MAFF311018a . KACC10331 . PXO99a . . . XOO0037 (novel) NA PXO03356 N N XOO0103 (XopF) XOO0074 PXO03413 XAC2785 XCV0414 XCV2942 XOO0148 (AvrBs2) XOO0168 PXO03330 XAC0076 XCV0052 XOO0315 (XopN) XOO0343 PXO02760 XAC2786 XCV2944 XOO3150 (XopN) NA NA XAC2786 XCV2944 XOO1488 (novel) NA NA N N XOO1669 (conserved) XOO1768 PXO01625 XAC3085 XCV3215 XOO2210 (conserved) XOO4824 N NA N XOO2877 (novel) NA PXO00236 N N XOO3803 (conserved) XOO4033 PXO04172 XAC0601 XCV0657 XOO4134 (conserved) XOO4391 PXO03819 XAC0277 XCV0285 XOO2402 (HopAS) XOO2543 PXO01041 XAC2009 XCC2059 XOO3222 (XopP) XOO3425 PXO02107 XAC1208 XCV1236 XOO3426 XOO2875 (XopX, HopAE) XOO3022 PXO00234 XAC0543 XCV3785 XOO4042 XOO4287 PXO03702 XCV0572 XOO4208 (XopQ) XOO4466 PXO03901 XAC433 XCV4438 a Previously identified candidate effectors are in parentheses. Conserved, Newly identified candidate effector and conserved in other species; novel, unique to Xoo. Open in new tab ). Three of the T3 effector genes are novel to Xoo (Furutani et al., 2009). The value of RLK-type resistance in rice is evident through the investigations of Xa21 and Xa26 (Lee et al., 2006; Cao et al., 2007). At the same time, some Xoo strains are virulent in Xa21 or Xa26 plants (Lee et al., 1999, 2003; Jeung et al., 2006), and evidence from the case of FLS2 and related proteins indicates that PTI is also vulnerable to T3 effector interference (Gohre et al., 2008; Xiang et al., 2008; reviewed in more detail in this issue). Xoo and possibly Xoc appear to have built a lifestyle that is dependent on a single family of T3 effector genes. This strategy, while providing interesting insight to the vulnerabilities and driving forces of rice genome evolution, may also prove the Achilles' heel in future strategies for durable and broad resistance to bacterial blight and streak in rice. ACKNOWLEDGMENTS We thank Dr. Zhongchao Yin for unpublished photographs. 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Nature 428 : 764 – 767 Crossref Search ADS PubMed Author notes * Corresponding author; e-mail [email protected]. The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Frank F. White ([email protected]). [C] Some figures in this article are displayed in color online but in black and white in the print edition. www.plantphysiol.org/cgi/doi/10.1104/pp.109.139360 © 2009 American Society of Plant Biologists This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Dellagi, Alia; Segond, Diego; Rigault, Martine; Fagard, Mathilde; Simon, Clara; Saindrenan, Patrick; Expert, Dominique

doi: 10.1104/pp.109.138636pmid: 19448037

Abstract Siderophores (ferric ion chelators) are secreted by organisms in response to iron deficiency. The pathogenic enterobacterium Erwinia chrysanthemi produces two siderophores, achromobactin and chrysobactin (CB), which are required for systemic dissemination in host plants. Previous studies have shown that CB is produced in planta and can trigger the up-regulation of the plant ferritin gene AtFER1. To further investigate the function of CB during pathogenesis, we analyzed its effect in Arabidopsis (Arabidopsis thaliana) plants following leaf infiltration. CB activates the salicylic acid (SA)-mediated signaling pathway, while the CB ferric complex is ineffective, suggesting that the elicitor activity of this siderophore is due to its iron-binding property. We confirmed this hypothesis by testing the effect of siderophores structurally unrelated to CB, including deferrioxamine. There was no activation of SA-dependent defense in plants grown under iron deficiency before CB treatment. Transcriptional analysis of the genes encoding the root ferrous ion transporter and ferric chelate reductase, and determination of the activity of this enzyme in response to CB or deferrioxamine, showed that these compounds induce a leaf-to-root iron deficiency signal. This root response as well as ferritin gene up-regulation in the leaf were not compromised in a SA-deficient mutant line. Using the Arabidopsis-E. chrysanthemi pathosystem, we have shown that CB promotes bacterial growth in planta and can modulate plant defenses through an antagonistic mechanism between SA and jasmonic acid signaling cascades. Collectively, these data reveal a new link between two processes mediated by SA and iron in response to microbial siderophores. Iron is essential for most forms of life. It is required for the catalytic activity of proteins mediating electron transfer and redox reactions, such as those involved in respiration, photosynthesis, DNA synthesis, and defense against reactive oxygen species. However, it is often unavailable because it is present as insoluble ferric hydroxide complexes in aerobiosis and at neutral pH. In its ferrous form, iron is more soluble and catalyzes the Fenton reaction in the presence of hydrogen peroxide, which leads to the formation of hydroxyl radicals, resulting in protein denaturation, DNA breaks, and lipid peroxidation (Pierre and Fontecave, 1999). Therefore, iron acquisition, utilization, and storage are subject to different levels of homeostatic regulation. In plants, iron is assimilated from the soil through the roots (Briat et al., 2007; Kim and Guerinot 2007). Under iron deficiency, Arabidopsis (Arabidopsis thaliana) activates processes described as strategy I based on the acidification of the soil by H+-ATPases, iron reduction by a ferric chelate reductase (FRO2; Robinson et al., 1999), and Fe2+ transport across the plasma membrane of root epidermal cells via the iron transporter IRT1 (Eide et al., 1996). Iron is then transported to plant organs essentially as citrate and nicotianamine complexes (Briat et al., 2007). Storage and buffering in dedicated compartments including apoplast and organelles (vacuole, plastids) avoid iron toxicity (Briat et al., 2007). In plastids, ferritins represent the major iron-containing proteins. In Arabidopsis, the ferritins AtFER1 to AtFER4 are mainly involved in buffering iron and protect the plant cells against oxidative stress (Ravet et al., 2009). Vacuolar iron stores can be mobilized to the cytosol via the divalent metal transporters AtNRAMP3 and AtNRAMP4 during seedling development (Lanquar et al., 2005). Microorganisms have developed powerful iron acquisition systems based on the production of siderophores, which are selective ferric ion chelators secreted in response to iron deficiency (Andrews et al., 2003; Winkelmann, 2007). Siderophores have low molecular weights and very diverse chemical structures that can contain one or a combination of several types of iron-binding moieties: hydroxamate, catecholate, and hydroxycarboxylate. Once loaded with iron, siderophores are specifically transported through the bacterial envelope via protein transporters; in the cytosol, iron is reduced and distributed to iron-containing molecules. During microbial infection, a competition for iron between the host and the microorganism may take place. Phytopathogenic bacteria and fungi can use siderophores to multiply in the host and to promote infection (Expert, 1999; Haas et al., 2008). Oide et al. (2006) demonstrated that in four ascomycete species, Cochliobolus miyabeanus, Cochliobolus heterostrophus, Fusarium graminareum, and Alternaria brassicicola, siderophores are required for resistance to hydrogen peroxide and for full pathogenicity on their respective hosts maize (Zea mays), rice (Oryza sativa), wheat (Triticum aestivum), and Arabidopsis. Likewise, the fire blight-causing agent Erwinia amylovora takes advantage of its siderophore deferrioxamine (DFO) for the infection of apple (Malus domestica) seedlings and flowers and for the resistance to hydrogen peroxide (Dellagi et al., 1998). The importance of iron homeostasis in plant disease has been largely documented in the case of the bacterial pathogen Erwinia chrysanthemi (Expert, 1999). E. chrysanthemi is an enterobacterium causing soft rot on economically important crops including potato (Solanum tuberosum) and chicory (Cichorium intybus) and ornamentals like Saintpaulia plants (Pérombelon, 2002). The bacterial cells invade the intercellular spaces of parenchymatous tissues and secrete large quantities of plant cell wall-degrading enzymes, leading to tissue disorganization (Murdoch et al., 1999). Under iron deficiency, E. chrysanthemi releases two siderophores: the hydroxycarboxylate achromobactin, which is produced when iron becomes limiting (Münzinger et al., 2000), and the catecholate chrysobactin (CB; Persmark et al., 1989), which prevails under severe iron deficiency. CB and achromobactin production are required for the systemic progression of maceration symptoms on the hosts (Enard et al., 1988; Dellagi et al., 2005; Franza et al., 2005). Neema et al. (1993) showed that the production of CB enables bacterial cells to compete with plant cells for iron, preventing sequestration of this metal by the plant ferritins. Consistently, the low availability of iron in the apoplasm of infected Saintpaulia leaves induces the expression of the fct gene, encoding the bacterial outer membrane ferric-chrysobactin (Fe-CB) transporter, which is up-regulated under iron depletion (Masclaux and Expert, 1995). Unlike in mammals, where the iron-sequestrating proteins of the transferrin family are able to reduce extracellular iron availability upon infection (for review, see Schaible and Kaufmann, 2004; Weinberg, 2009), there is no evidence that these proteins exist in plants. In Arabidopsis, the genes encoding the ferritin AtFER1 and the vacuolar metal transporters AtNRAMP3 and AtNRAMP4 are involved in basal resistance to E. chrysanthemi, indicating that changes in plant iron trafficking occur during infection (Dellagi et al., 2005; Segond et al., 2009). Interestingly, AtFER1 gene expression can be activated by the purified siderophore CB and not by Fe-CB in Arabidopsis leaves (Dellagi et al., 2005). This observation led us to suppose that the siderophore could act as a modulator of plant defense responses, since AtFER1 is part of the defense reactions triggered by E. chrysanthemi. Thus, we investigated the role of CB in the activation of Arabidopsis defense responses triggered by E. chrysanthemi upon infection, namely the salicylic acid (SA), the jasmonic acid (JA), and the ethylene (ET) pathways, which are three major signaling pathways involved in the plant's immune network (Glazebrook, 2005; Fagard et al., 2007). In this work, we show that CB activates the SA-dependent defense pathway and that this process is dependent on iron availability in the plant. Not only CB but other types of siderophores could be elicitors, revealing a new function for these iron ligands in plant-microbe interactions. We also show that, when infiltrated into leaves, siderophores provoke iron deficiency in the roots. This work describes a new link between iron and immunity, which appears to be more complex than a simple nutritional competition. RESULTS CB Triggers the Signaling Cascade Mediated by SA In Arabidopsis, E. chrysanthemi triggers defenses mediated by the major signaling molecules SA, JA, and ET, as revealed by marker gene expression analysis 24 h after bacterial inoculation (Fagard et al., 2007). We thus investigated whether the siderophore CB is able to activate similar responses. We monitored the expression of SA-, ET-, and JA-dependent defense genes by reverse transcription (RT)-PCR after water or CB infiltration in Arabidopsis leaves, using 0.25, 0.5, and 1 mm CB. All concentrations gave similar results (Supplemental Fig. S1), but the reproducibility of the data was best with 1 mm CB. Therefore, we used 1 mm CB in all the following experiments. We found that 24 h post infiltration (hpi), CB strongly activates the expression of the SA marker gene PR1 (Fig. 1A). Figure 1. Open in new tabDownload slide PR1 gene expression and SA production in Arabidopsis leaves following CB treatment. A, Expression patterns of PR1 (SA pathway) and PDF1.2 (ET/JA pathway) were monitored by RT-PCR using RNAs extracted from Col-0 leaves at the indicated times after infiltration of distilled water or CB. The constitutive EF1α gene was used as a control. B, GUS staining of leaves from transgenic PR1::GUS plants 24 h after infiltration of water, CB, or SA. C, Total SA content was measured by HPLC in Col-0 leaves 24 h after the indicated treatments (μg SA g−1 fresh weight [FW]). n = 6, error bars indicate sd, and the asterisk indicates a significant difference from the control by the Mann-Whitney test (P < 0.01). Figure 1. Open in new tabDownload slide PR1 gene expression and SA production in Arabidopsis leaves following CB treatment. A, Expression patterns of PR1 (SA pathway) and PDF1.2 (ET/JA pathway) were monitored by RT-PCR using RNAs extracted from Col-0 leaves at the indicated times after infiltration of distilled water or CB. The constitutive EF1α gene was used as a control. B, GUS staining of leaves from transgenic PR1::GUS plants 24 h after infiltration of water, CB, or SA. C, Total SA content was measured by HPLC in Col-0 leaves 24 h after the indicated treatments (μg SA g−1 fresh weight [FW]). n = 6, error bars indicate sd, and the asterisk indicates a significant difference from the control by the Mann-Whitney test (P < 0.01). We did not find any significant modification in the expression of PDF1.2, which is a good marker for the ET/JA pathway (Penninckx et al., 1998). Thus, we focused on the SA pathway. We then used Arabidopsis lines expressing GUS fusions to the PR1 promoter. We observed a strong GUS staining 24 h following infiltration of CB, which was not detected after water infiltration (Fig. 1B). The intensity of GUS staining in leaves treated with CB was similar to that observed in SA-treated leaves, used as positive controls. To determine whether the activation of PR1 expression correlated with an accumulation of SA, we measured the SA content by HPLC in Arabidopsis leaves 24 h after CB treatment. Figure 1C shows that siderophore treatment results in a 2- to 3-fold increase in SA content 24 hpi compared with control leaves. Altogether, these data show that CB triggers the SA defense pathway when infiltrated into Arabidopsis leaves. SA can be synthesized in Arabidopsis through two distinct pathways, involving either Phe ammonia-lyase or isochorismate synthase (ICS1/SID2). Because it was previously found that the ICS1/SID2 pathway is involved in the up-regulation of PR1 after E. chrysanthemi infection (Fagard et al., 2007), we proceeded on the hypothesis that this pathway could be also required for the CB-induced response. Therefore, we monitored the accumulation of SA in ecotype Columbia (Col-0) and in a sid2 mutant (Nawrath and Métraux, 1999). While CB infiltration resulted in a 2- to 3-fold accumulation of total SA in Col-0 leaves compared with control leaves 24 hpi, no significant accumulation of this hormone was observed in sid2 leaves (Fig. 2A). Figure 2. Open in new tabDownload slide Roles of genes of the SA pathway in the signaling cascade triggered by CB. A, Total SA content (measured as in Fig. 1) fold induction (i.e. ratio of SA content in CB-treated leaves to SA content in water control). n = 6, error bars indicate sd, and the asterisk indicates a significant difference from Col-0 using the Mann-Whitney test (P < 0.01). B, RNAs from leaves were harvested after infiltration of water or CB (genotypes and times indicated). RT-PCR results with the indicated defense gene-specific primers are shown. The constitutive EF1α gene was used as a control. C, Expression profiles of AtFER1 and PR1 monitored by RT-PCR. RNAs were extracted from leaves of the indicated mutant genotypes harvested at the given times after the treatment indicated. Figure 2. Open in new tabDownload slide Roles of genes of the SA pathway in the signaling cascade triggered by CB. A, Total SA content (measured as in Fig. 1) fold induction (i.e. ratio of SA content in CB-treated leaves to SA content in water control). n = 6, error bars indicate sd, and the asterisk indicates a significant difference from Col-0 using the Mann-Whitney test (P < 0.01). B, RNAs from leaves were harvested after infiltration of water or CB (genotypes and times indicated). RT-PCR results with the indicated defense gene-specific primers are shown. The constitutive EF1α gene was used as a control. C, Expression profiles of AtFER1 and PR1 monitored by RT-PCR. RNAs were extracted from leaves of the indicated mutant genotypes harvested at the given times after the treatment indicated. We can conclude that the SID2 gene is necessary for the biosynthesis of SA in response to CB. In order to check whether up-regulation of the PR1 gene in response to CB is dependent on SA biosynthesis, we monitored by RT-PCR the accumulation of its transcript in Col-0 and sid2 leaves treated with water or CB. PR1 expression was strongly activated by the siderophore 24 hpi in Col-0 leaves. By contrast, the presence of PR1 transcripts was hardly detected in sid2 leaves infiltrated with the siderophore (Fig. 2B). The up-regulation of PR1 by CB is thus dependent on the accumulation of SA in the leaves via the SID2 gene activity. We also monitored the expression of PAD4 and EDS5 genes known to act upstream of PR1 in the SA-mediated response. PAD4 encodes a protein similar to lipases and is required for resistance and accumulation of SA following infection with Pseudomonas syringae pv maculicola and Hyaloperonospora parasitica (Glazebrook et al., 1996, 1997; Zhou et al., 1998). EDS5 encodes a MATE-type multidrug efflux pump presumably involved in SA efflux from the chloroplast and is required for resistance to P. syringae and H. parasitica and the accumulation of SA in response to P. syringae (Nawrath and Métraux, 1999; Nawrath et al., 2002). We found that both PAD4 and EDS5 are up-regulated between 7 and 24 h after CB treatment (Fig. 2B). This response is independent of SA accumulation, since it was similar in Col-0 and in the sid2 deficient lines (Fig. 2B). To determine whether the SA-mediated response activated by CB requires functional PAD4 and EDS5 genes, as well as the NPR1 gene encoding the SA sensor protein (Mou et al., 2003), CB or water was infiltrated onto the leaves of Col-0, eds5, pad4, and npr1 plants (Fig. 2C). No up-regulation of PR1 was observed in eds5 and npr1 mutants, indicating that the corresponding genes must be functional to mediate the response to CB. In the pad4 mutant, the expression of PR1 was still up-regulated. Collectively, these results indicate that CB activates a signaling pathway leading to PR1 up-regulation that is independent of PAD4 but dependent on SA production via SID2 and EDS5 and on the perception of this hormone via NPR1. As CB infiltration in Arabidopsis leaves activates the expression of the ferritin-encoding gene AtFER1 (Dellagi et al., 2005), we asked whether this response requires the integrity of the SA pathway. We monitored the expression of this gene in Col-0, sid2, eds5, and npr1 leaves treated with CB. AtFER1 up-regulation was observed in all lines in response to the siderophore (Fig. 2, B and C). Similar results were obtained with the pad4 mutant line. These results indicate that AtFER1 up-regulation by CB is independent of the SA-mediated signaling pathway. The Iron-Chelating Property of Siderophores Is Required for the Activation of the SA-Mediated Signaling Pathway AtFER1 gene transcription is not activated by Fe-CB (Dellagi et al., 2005). Siderophore iron-binding activity is measured by calculating the pFe {defined as −log [Fe3+], where [Fe3+] = free [Fe3+] in solution calculated at determined concentrations of ligand and Fe(III) and pH; for CB, pFe = 14.5 [Tomisić et al., 2008]}. Thus, we analyzed the expression of the SA pathway in response to leaf infiltration with CB or Fe-CB (Fig. 3A). Figure 3. Open in new tabDownload slide Importance of the chemical state of different siderophores in the activation of the SA pathway. Col-0 leaves were treated with water, CB, Fe-CB, DFO, or Fe-DFO as indicated. RNAs were extracted from leaves harvested 24 h after treatment, and RT-PCR with the indicated defense markers was performed. The constitutive EF1α gene was used as a control. Figure 3. Open in new tabDownload slide Importance of the chemical state of different siderophores in the activation of the SA pathway. Col-0 leaves were treated with water, CB, Fe-CB, DFO, or Fe-DFO as indicated. RNAs were extracted from leaves harvested 24 h after treatment, and RT-PCR with the indicated defense markers was performed. The constitutive EF1α gene was used as a control. We found that the PR1, EDS5, and PAD4 genes were not up-regulated by Fe-CB. These results indicate that the elicitor activity of the siderophore is related to its chemical state. To determine whether the activation of the SA pathway is specific to CB, we tested the activity of a structurally unrelated siderophore, DFO. DFO is produced by the bacterial plant pathogen E. amylovora (Kachadourian et al., 1997) and is able to activate the transcription of AtFER1 in Arabidopsis leaves (Dellagi et al., 2005). Compounds of the DFO family harbor three hydroxamate groups that can bind Fe3+ very efficiently (pFe = 24.2; Tomisić et al., 2008). We found that, like CB, DFO infiltrated onto Arabidopsis leaves results in transcript accumulation of genes from the SA pathway (Fig. 3B). Ferrioxamine (Fe-DFO) did not induce this response. The same results were obtained with ferrichrome, another hydroxamate-type siderophore (data not shown). Collectively, these results suggest that the presence of siderophores in intercellular spaces of Arabidopsis leaves, when they are iron free, induces an SA-mediated response similar to that activated by pathogens. This process is not specific to the siderophore structure, as it can be activated by either catecholates or hydroxamates. Activation of the SA-Mediated Signaling Pathway by CB Depends on Iron Availability to the Plant When present in the plant tissue, a siderophore should rapidly take up iron from iron-containing molecules, suggesting that this metal plays a critical role in activation of the SA-dependent process, depending on whether it is bound or not to the ligand. To check whether the nutritional iron status of the plant influences the SA-mediated response, we compared the effect of CB on plants grown under iron-sufficient and iron-deficient conditions. We used hydroponically grown plants for which nutritional iron was adjusted as described in “Materials and Methods” and analyzed the expression of PR1. The results (Fig. 4) Figure 4. Open in new tabDownload slide Influence of plant iron nutrition on PR1 and AtFER1 gene expression levels following CB treatment. Hydroponically grown plants under 50 μ m Fe-EDTA for approximately 6 weeks (+Fe) or under 50 μ m Fe-EDTA for 5 weeks and then without iron for 5 d (−Fe) were infiltrated with water or CB (1 mm) as indicated. RT-PCR with the indicated gene-specific primers was performed with RNAs extracted from leaves harvested at the indicated times after treatment. The constitutive EF1α gene was used as a control. Figure 4. Open in new tabDownload slide Influence of plant iron nutrition on PR1 and AtFER1 gene expression levels following CB treatment. Hydroponically grown plants under 50 μ m Fe-EDTA for approximately 6 weeks (+Fe) or under 50 μ m Fe-EDTA for 5 weeks and then without iron for 5 d (−Fe) were infiltrated with water or CB (1 mm) as indicated. RT-PCR with the indicated gene-specific primers was performed with RNAs extracted from leaves harvested at the indicated times after treatment. The constitutive EF1α gene was used as a control. indicate that PR1 is up-regulated in plants grown under iron sufficiency, while this was not the case in iron-deficient plants. We measured the amounts of SA in leaves treated with CB from plants grown under both conditions. In agreement with PR1 expression profiles, we found that iron-deficient plants do not accumulate significant amounts of SA (data not shown). These results show that iron present in the plant growth medium is necessary for up-regulation of the SA-mediated pathway in response to CB. Up-regulation of AtFER1 is observed in the presence of iron (Gaymard et al., 1996). As expected, the up-regulation of AtFER1 occurring in response to CB treatment (Dellagi et al., 2005) was not detected with plants grown under iron-deficient conditions (Fig. 4). Siderophores Trigger an Iron Deficiency Response in the Roots As the presence of siderophores in the plant leaves can lead to iron withholding, we investigated whether these ligands are able to trigger an iron deficiency reaction in the plant. We analyzed the expression of IRT1 and FRO2 genes, encoding the iron transporter IRT1 (Eide et al., 1996) and the ferric chelate reductase FRO2 (Robinson et al., 1999), known to respond to iron deficiency in the root. Both genes appeared to be up-regulated in roots 7 h after CB leaf treatment compared with control plants (Fig. 5A). Figure 5. Open in new tabDownload slide Iron deficiency root response caused by CB or DFO leaf infiltration. Col-0 plants were grown under hydroponic conditions with 50 μ m Fe-EDTA. A and B, Leaves were infiltrated with water, CB, or DFO, and then RNAs were extracted from roots harvested at the indicated times after treatment. Expression patterns of IRT1 and FRO2 genes were analyzed by northern blots. Ethidium bromide staining is shown as a loading control. C, Ferric chelate reductase activity measured in roots at the indicated times after CB treatment. n = 6, error bars indicate sd, and the asterisk indicates a significant difference from the control using the Mann-Whitney test (P < 0.01). FW, Fresh weight. Figure 5. Open in new tabDownload slide Iron deficiency root response caused by CB or DFO leaf infiltration. Col-0 plants were grown under hydroponic conditions with 50 μ m Fe-EDTA. A and B, Leaves were infiltrated with water, CB, or DFO, and then RNAs were extracted from roots harvested at the indicated times after treatment. Expression patterns of IRT1 and FRO2 genes were analyzed by northern blots. Ethidium bromide staining is shown as a loading control. C, Ferric chelate reductase activity measured in roots at the indicated times after CB treatment. n = 6, error bars indicate sd, and the asterisk indicates a significant difference from the control using the Mann-Whitney test (P < 0.01). FW, Fresh weight. Fe-CB infiltration in leaves did not activate the expression of these genes, as expected (Fig. 5A). We also observed a similar response to that observed with CB after infiltration of DFO (Fig. 5B). We then determined the enzymatic activity of FRO2 in roots from plants treated with CB. Figure 5C shows that 24 h after siderophore infiltration in leaves of hydroponically grown plants, the FRO2 activity in roots was three times higher than in control plants. These data indicate that the presence of a siderophore in Arabidopsis leaves causes an iron deficiency in the roots, suggesting the propagation of a leaf-to-root signal. The SA- and ET-Mediated Signaling Pathways Are Dispensable for IRT1 and FRO2 Up-Regulation by CB As the SA pathway is induced by CB and ET is involved in the up-regulation of FRO2 and IRT1 genes in response to iron deficiency (Lucena et al., 2006), we investigated the role of SA and ET in expression of the root response induced by CB. We used the sid2 and ein2 mutants, the latter being affected in ET perception (Alonso et al., 1999). CB infiltration in the leaves of these mutants led to the activation of IRT1 and FRO2, and notably, expression levels of these genes were higher in the sid2 mutant (Fig. 6). Figure 6. Open in new tabDownload slide IRT1 and FRO2 gene expression in Arabidopsis SA and ET mutants following CB leaf treatment. Plants were grown under hydroponic conditions with 50 μ m Fe-EDTA. Expression patterns of IRT1 and FRO2 genes were analyzed by northern blots using RNA extracted from roots harvested from ET (ein2) and SA (sid2) mutants at the indicated times after treatment. Ethidium bromide staining is shown as a loading control. Figure 6. Open in new tabDownload slide IRT1 and FRO2 gene expression in Arabidopsis SA and ET mutants following CB leaf treatment. Plants were grown under hydroponic conditions with 50 μ m Fe-EDTA. Expression patterns of IRT1 and FRO2 genes were analyzed by northern blots using RNA extracted from roots harvested from ET (ein2) and SA (sid2) mutants at the indicated times after treatment. Ethidium bromide staining is shown as a loading control. Thus, the SA and ET pathways are not required to mediate the iron deficiency root response induced by CB. CB Manipulates Plant Defenses and Promotes in Planta Bacterial Growth The data presented above indicate that siderophores act as elicitors of plant defense controlled by the SA hormone. As E. chrysanthemi triggers a set of defenses in Arabidopsis during infection, including the SA-mediated signaling pathway (Fagard et al., 2007), we wondered if the activation of this pathway was reduced after inoculation of a siderophore-deficient mutant compared with the wild-type strain. Thus, we used the E. chrysanthemi CB-deficient mutant affected in the cbsE gene (Franza et al., 2005). Expression of the PR1 gene was monitored 3, 7, and 24 h after infiltration of Arabidopsis leaves with wild-type cells or the siderophore-deficient mutant. As observed previously, the PR1 gene was strongly up-regulated by the wild-type bacteria compared with the control plants (Fig. 7A). Figure 7. Open in new tabDownload slide Effects of CB on the expression of PR1 and PDF1.2 genes during E. chrysanthemi infection. A, Col-0 leaves were infiltrated with 10 mm MgSO4 or 107 colony-forming units mL−1 bacterial suspension of E. chrysanthemi wild type (E.ch) or CB negative mutant (E.ch cbs). Leaves were harvested at the indicated times after treatment. Expression patterns of PR1 (SA pathway) and PDF1.2 (ET/JA pathway) were monitored by RT-PCR. The constitutive EF1α gene was used as a control. B, RT-PCR using RNAs extracted from leaves 24 h after treatment with 0.05% (w/v) methanol (cont), JA, or JA + CB. C, Plants were infiltrated with water or CB 48 h before inoculation with a bacterial suspension of wild-type E. chrysanthemi cells. Leaves were harvested at the times indicated after bacterial infiltration, and then bacterial counts were performed as indicated in “Materials and Methods.” n = 6, error bars indicate sd, and the asterisk indicates a significant difference from the control using the Mann-Whitney test (P < 0.05). Figure 7. Open in new tabDownload slide Effects of CB on the expression of PR1 and PDF1.2 genes during E. chrysanthemi infection. A, Col-0 leaves were infiltrated with 10 mm MgSO4 or 107 colony-forming units mL−1 bacterial suspension of E. chrysanthemi wild type (E.ch) or CB negative mutant (E.ch cbs). Leaves were harvested at the indicated times after treatment. Expression patterns of PR1 (SA pathway) and PDF1.2 (ET/JA pathway) were monitored by RT-PCR. The constitutive EF1α gene was used as a control. B, RT-PCR using RNAs extracted from leaves 24 h after treatment with 0.05% (w/v) methanol (cont), JA, or JA + CB. C, Plants were infiltrated with water or CB 48 h before inoculation with a bacterial suspension of wild-type E. chrysanthemi cells. Leaves were harvested at the times indicated after bacterial infiltration, and then bacterial counts were performed as indicated in “Materials and Methods.” n = 6, error bars indicate sd, and the asterisk indicates a significant difference from the control using the Mann-Whitney test (P < 0.05). Infection by the siderophore-deficient mutant resulted in reduced expression of this gene. This result indicates that CB, during bacterial infection, contributes to the activation of the SA pathway, although it is not the only elicitor of this process. Interestingly, expression of PDF1.2, the gene marker of the ET/JA pathway that is not activated by wild-type bacteria 24 h after infiltration, was strongly up-regulated in response to the siderophore-deficient mutant (Fig. 7A). This result suggests that CB represses the expression of PDF1.2. As PDF1.2 expression is known to be activated by JA, we analyzed the expression of PDF1.2 in leaves treated with JA or with both JA and CB (Fig. 7B). We observed an accumulation of PDF1.2 transcripts in response to JA, which was not detected after coinfiltration of CB and the hormone. This result confirms that CB can repress the expression of PDF1.2. Previous studies using bacterial mutants unable to produce CB or achromobactin have shown that these siderophores promote the infection process in host plants (Enard et al., 1988; Dellagi et al., 2005; Franza et al., 2005). We thus asked whether infiltration of CB onto Arabidopsis leaves prior to E. chrysanthemi inoculation could affect the bacterial growth. We infiltrated water or 1 mm CB onto Arabidopsis leaves 48 h before bacterial challenge. Bacterial populations were determined over 2 d post inoculation (Fig. 7C). In the control leaves preinfiltrated with water, E. chrysanthemi grew by less than 1 order of magnitude after 2 d of infection. In the leaves pretreated with CB, we observed a much faster growth and an increase in bacterial counts by 1 order of magnitude. These data indicate that preinfiltration of CB stimulates bacterial growth. DISCUSSION The plant pathogenic bacterium E. chrysanthemi requires the production of siderophores for systemic progression in host tissues (Enard et al., 1988; Dellagi et al., 2005; Franza et al., 2005). Production of siderophores and pectinases is controlled by iron availability, indicating that high-affinity iron uptake by this bacterium is a critical factor during pathogenesis (Franza et al., 2002). In order to know the role of siderophores in the infection process more precisely, we need to understand how these compounds are perceived in the host. In this work, we have investigated the plant's response to the siderophore CB. We found that two physiological functions are modulated by this molecule: plant defense and iron assimilation. Role of CB in the Activation of the SA-Mediated Signaling Pathway We found that CB in Arabidopsis activates the SA-mediated signaling pathway leading to PR1 gene expression. Our results using the sid2 mutant show that CB activates SA biosynthesis in Arabidopsis. The structural similarity between SA and CB allowed the hypothesis that CB or its potential degradation products could act as precursor(s) in SA biosynthesis. However, since other siderophores with no structural relationship to SA are also able to trigger the SA pathway, we excluded this hypothesis. CB also requires NPR1 to activate the expression of PR1. The NPR1 protein is an important player in SA signaling and in systemic acquired resistance (Dong, 2004). In the cytosol, it is present as disulfide-bound oligomers that monomerize following reduction consecutive to SA-controlled redox changes (Mou et al., 2003). The monomers are translocated to the nucleus, where they interact with TGA transcription factors that recognize cis elements in PR gene promoters (Johnson et al., 2003). This means that the response induced by the siderophore could also result in a cellular redox change involving SA and leading to the activation of the PR1 gene via NPR1. We also show that siderophore-mediated PR1 up-regulation does not require PAD4. PAD4 encodes a triacyl-glycerol lipase acting upstream of SA (Jirage et al., 1999) and is necessary for SA accumulation and amplification of SA-dependent defense responses (Zhou et al., 1998). It is not required for PR1 up-regulation during the hypersensitive response observed in an incompatible interaction involving resistant plants. However, PAD4 is required for full PR1 up-regulation in compatible interactions involving susceptible plants (Zhou et al., 1998). In light of these data, it is possible that the SA response triggered by a siderophore is strong enough and is comparable to an incompatible interaction, except that there is no reaction of cell death. Indeed, we never observed necrosis after siderophore treatment at the macroscopic level or at the microscopic level after trypan blue staining (data not shown). Our results indicate that CB activates the AtFER1 gene independently of the SA-mediated signaling pathway (Fig. 8). Figure 8. Open in new tabDownload slide Diagram showing the responses of Arabidopsis to microbial siderophores. Leaf infiltration of iron-free siderophores (CB or DFO) activates the SA-mediated signaling pathway leading to PR1 up-regulation, the basal defense marker PAD4, ferritin accumulation via AtFER1, and root iron uptake via IRT1 and FRO2. Up-regulation of IRT1 and FRO2 appeared to be partially repressed by SA. Activation of the SA pathway and AtFER1 up-regulation depends on iron availability to the plant (indicated with dashed arrows). Further details are discussed in the text. Figure 8. Open in new tabDownload slide Diagram showing the responses of Arabidopsis to microbial siderophores. Leaf infiltration of iron-free siderophores (CB or DFO) activates the SA-mediated signaling pathway leading to PR1 up-regulation, the basal defense marker PAD4, ferritin accumulation via AtFER1, and root iron uptake via IRT1 and FRO2. Up-regulation of IRT1 and FRO2 appeared to be partially repressed by SA. Activation of the SA pathway and AtFER1 up-regulation depends on iron availability to the plant (indicated with dashed arrows). Further details are discussed in the text. Thus, it may be possible that regulation of the ferritin gene by CB takes place upstream of the SA response. It would be helpful to determine whether AtFER1 contributes to the responses induced by CB in an atfer1 mutant. As an iron-buffering molecule, ferritin could contribute to changes in the cellular iron status and activate downstream signals. Role of CB in the Activation of the Iron Deficiency Root Response We recently reported that infection of Arabidopsis by E. chrysanthemi triggers an iron deficiency response in the roots (Segond et al., 2009). This work shows that this response is also induced by infiltration of a siderophore on the leaf, suggesting that this ligand is responsible for the root reaction when it is released by bacterial cells during infection. The elicitor activity of siderophores is likely due to their strong iron-chelating capacity rather than to recognition by a plant receptor. Indeed, we found that the siderophores induce a reaction similar to iron deficiency consisting of IRT1 and FRO2 expression and production of the FRO2 enzymatic activity. It is tempting to speculate that the iron taken up by the roots is rapidly translocated to the leaves, a process that may cause an oxidative stress (Fig. 8). This oxidative stress could activate the SA pathway and AtFER1 gene expression, as these two responses are known to be inducible by reactive oxygen species (Leon et al. 1995; Gaymard et al., 1996; Petit et al., 2001). Two observations are in agreement with this hypothesis. First, under iron deficiency, CB treatment does not induce the up-regulation of PR1 or that of AtFER1, indicating that iron is required for activation of the SA response and confirming that this metal is essential to AtFER1 up-regulation. Second, the expression of IRT1 is rapid (7 hpi) but decreases between 7 and 24 h, indicating that the iron deficiency signal disappears during this period, likely because of a negative feedback due to iron uptake via IRT1. The timing of activation of the various SA marker genes (7–24 hpi) is compatible with this interpretation. In addition, the protein IRT1 can transport other cations than Fe2+, including Zn2+, Mn2+, and Cd2+ (Korshunova et al., 1999), and it is conceivable that some of these metals are taken up by the plant after IRT1 induction and contribute to the responses observed. We investigated whether SA or ET is involved in the activation of the iron deficiency response by CB. Our data show that the ein2 mutation does not affect IRT1 and FRO2 up-regulation following CB treatment, suggesting that ET is not involved in this process. On the other hand, we found that IRT1 and FRO2 transcripts accumulate to higher levels in the sid2 mutant compared with the wild-type ecotype. This observation suggests that SA could exert a negative control of the iron deficiency response. This is consistent with the iron-binding capacity of SA (pFe = 12.1; Nurchi et al., 2009), a property that might confer onto this molecule a cellular iron-sensing function, as suggested in bacteria (Adilakshmi et al., 2000). Role of CB in the Control of E. chrysanthemi Pathogenesis CB pretreatment enhances the multiplication of E. chrysanthemi cells in the leaf (Fig. 7), which is in agreement with the fact that siderophore-deficient mutants are affected in their aggressiveness (Enard et al., 1988; Dellagi et al., 2005; Franza et al., 2005). The weaker activation of PR1 after inoculation of the CB-deficient mutant compared with the wild-type strain indicates that CB produced during infection is likely responsible for activation of the SA pathway. However, under this condition, the PR1 gene is still expressed, indicating the existence of other elicitors of the SA pathway. Achromobactin, the second E. chrysanthemi siderophore, could contribute to this response, and oligogalacturonides generated by pectinases are likely to be involved (Fagard et al., 2007). We also found that, unlike wild-type bacteria, the CB-deficient mutant activates the expression of a marker of the JA/ET pathway, PDF1.2, 24 hpi. This suggests that CB represses this pathway that is involved in Arabidopsis resistance to E. chrysanthemi (Fagard et al., 2007). By activating the biosynthesis of SA via CB, the bacteria modulate the plant defense responses and take advantage of the antagonism between the SA and JA pathways. Furthermore, as siderophores activate iron uptake in the roots, the plant iron content must increase, thus explaining the beneficial effect of CB on E. chrysanthemi growth in the leaves. Some siderophores secreted by soil-borne Pseudomonas species (pyoverdin and pyocyanin) can promote systemic plant protection against soil and foliar pathogens, a phenomenon known as induced systemic resistance (Audenaert et al., 2002; Haas and Défago, 2005). Induced systemic resistance is known to be dependent on the ET and JA pathways and independent of the SA pathway (Pieterse et al., 1998). In this work, we show that the elicitor activity of the siderophore CB that we observed is SA dependent, indicating that this process is different from induced systemic resistance. In conclusion, this work shows that microbial pathogens can modulate the activity of the plant iron acquisition system via the modulation of siderophore production during infection and that this process can lead to changes in the expression of plant immune responses. These changes may be to the advantage of the pathogen or may help the plant to resist the pathogen. This could explain why in a number of plant-pathogen interactions, no role for siderophores was found in virulence, while in others, siderophores are important pathogenicity factors. The future challenges now are to better understand the molecular mechanisms by which siderophores activate the SA pathway and the root iron deficiency response. MATERIALS AND METHODS Plant Material and Growth Conditions Arabidopsis (Arabidopsis thaliana) seeds from the Col-0 ecotype were obtained from the Institut National de la Recherche Agronomique Versailles collection. The sid2-1 mutant was kindly donated by J.-P. Métraux. Seeds of ein2-1 (Guzman and Ecker, 1990), npr1-1 (Cao et al., 1994), pad4-1 (Glazebrook et al., 1997), and eds5-1 (Glazebrook et al., 1996) were provided by the Nottingham Arabidopsis Stock Center (Scholl et al., 2000). The PR1::GUS lines were kindly provided by F. Ausubel. Plants were grown as described by Fagard et al. (2007). Hydroponic cultures were performed as described by Segond et al. (2009) and were used for all experiments where roots were used for RNA extraction or FRO2 activity monitoring. Experiments with iron-starved plants were performed as follows. Plants were first grown under the above-described conditions for 5 weeks and then transferred to iron-deficient medium after washing the roots for 5 min with medium containing the reductant sodium dithionite (5.7 mm) and the chelator bathophenanthrolinedisulfonic acid (0.3 mm), both from Sigma. The plants were kept under high-moisture conditions during the experiments. Chemical Preparations Leaf treatment with the following chemicals was performed by infiltration of the solutions at the indicated concentrations in the intercellular leaf space using a syringe without a needle. CB was a gift from J. Buyer (Lu et al., 1996), and Fe-CB was prepared as described by Rauscher et al. (2002). DFO and Fe-DFO (a gift from R. Kachadourian) were prepared as described by Kachadourian et al. (1997). JA and SA were purchased from Sigma-Aldrich. Siderophores and SA were used at a concentration of 1 mm diluted in distilled water. JA was used at 0.1 mm in 0.05% (w/v) methanol. Bacterial Strains and Culture Conditions The wild-type strain, Erwinia chrysanthemi 3937 (our collection), was isolated from Saintpaulia ionantha (African violet). The CB-deficient mutant PPV11 is derived from strain 3937 that contains an insertional element inactivating the biosynthetic CB gene cbsE (cbs-E1::Ω; Franza et al., 2005). Growth conditions were as described by Dellagi et al. (2005). Plant Inoculations and Determination of Bacterial Growth To monitor bacterial growth after siderophore treatment, we first infiltrated water or CB on the entire leaf. Forty-eight hours later, a small hole was made with a needle within the leaf, and then 5 μL of a bacterial suspension at a density of 5 × 107 colony-forming units mL−1 made up in 50 mm potassium phosphate buffer (pH 7) was spotted on the hole. Leaves were harvested in 0.9% NaCl and ground using a pestle and sterile sand. The resulting suspensions were used for serial dilutions followed by plating on an appropriate medium. For RNA extractions and GUS fusions, we used a syringe without a needle to infiltrate the entire leaf or a portion of the leaf with SA, siderophore solution, or bacterial suspensions at 5 × 107 colony-forming units mL−1 in 10 mm MgSO4 (half a leaf was infiltrated for GUS staining). RNA Extraction, Northern Blotting, and RT-PCR Northern-blot hybridization was carried out as described by Dellagi et al. (2005). IRT1 and FRO2 probes were prepared as described by Segond et al. (2009). For RT-PCR analysis, reverse transcription was performed as described by Fagard et al. (2007). PCR runs were of 94°C for 4 min, 26 to 30 cycles, and each cycle consisting of 94°C for 30 s, 54°C to 58°C for 30 s, and 72°C for 1 min, with a final step of 72°C for 10 min to complete polymerization. Primers for EF1α, PR1, PAD4, CHIB, and EDS5 were described by Fagard et al. (2007). The other gene-specific primers were as follows: PDF1.2-F (AT5G44420; 5′-TCATGGCTAAGTTTGCTTCCATCATCACCC-3′) and PDF1.2-R (5′-GTAGATTTAACATGGGAC-3′). Equal cDNA amounts were checked by performing different PCR cycles with EF1α primers (Supplemental Fig. S2). Experiments were repeated at least three times. Representative data are shown. In Planta GUS Expression Detection In planta GUS expression detection was performed as described by Dellagi et al. (2005). Experiments were repeated three times with similar results. Root FRO Assays Root FRO activity was performed as described by Yi and Guerinot (1996). Briefly, roots from control plants or plants treated with the siderophore were incubated in a solution containing 0.1 mm Fe(III)-EDTA and 0.3 mm ferrozine in distilled water in the dark. After 20 min, the absorbance of the solution was measured at 562 nm, using the same solution without roots as a control. SA Treated leaves were harvested and then weighed before freezing in liquid nitrogen. They were ground in a frozen state in Eppendorf tubes using TissueLyser II (Qiagen). [7-14C]SA (1 nCi, 54 mCi mmol−1; New England Nuclear) was used for recovery determination. Total SA was extracted and analyzed as described by Baillieul et al. (1995) with a Nova-Pak 4-μm C-18 column (150 × 3.9 mm; Waters) as part of the Waters system (1525 Binary HPLC Pump, 2475 Multi λ Fluorescence Detector, 2996 Photodiode Array Detector, 717 Autosampler; Waters). Data were analyzed using Empower Pro Software (Waters). Supplemental Data The following materials are available in the online version of this article. Supplemental Figure S1. Defense gene expression in Arabidopsis leaves infiltrated with CB. Supplemental Figure S2. Validation of the RT-PCR approach using EF1α as a constitutively expressed gene. ACKNOWLEDGMENTS We thank Floriant Belvert for help with SA quantification. We thank J.-P. 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[W] The online version of this article contains Web-only data. www.plantphysiol.org/cgi/doi/10.1104/pp.109.138636 © 2009 American Society of Plant Biologists This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Römer, Patrick; Strauss, Tina; Hahn, Simone; Scholze, Heidi; Morbitzer, Robert; Grau, Jan; Bonas, Ulla; Lahaye, Thomas

doi: 10.1104/pp.109.139931pmid: 19448036

Abstract The pepper (Capsicum annuum) bacterial spot (Bs) resistance gene Bs3 and its allelic variant Bs3-E mediate recognition of the Xanthomonas campestris pv vesicatoria type III effector protein AvrBs3 and its deletion derivative AvrBs3Δrep16. Recognition specificity resides in the Bs3 and Bs3-E promoters and is determined by a defined promoter region, the UPA (for up-regulated by AvrBs3) box. Using site-directed mutagenesis, we defined the exact boundaries of the UPA AvrBs3 box of the Bs3 promoter and the UPA AvrBs3Δrep16 box of the Bs3-E promoter and show that both boxes overlap by at least 11 nucleotides. Despite partial sequence identity, the UPA AvrBs3 box and the UPA AvrBs3Δrep16 box were bound specifically by the corresponding AvrBs3 and AvrBs3Δrep16 proteins, respectively, suggesting that selective promoter binding of AvrBs3-like proteins is the basis for promoter activation specificity. We also demonstrate that the UPA AvrBs3 box retains its functionality at different positions within the pepper Bs3 promoter and confers AvrBs3 inducibility in a novel promoter context. Notably, the transfer of the UPA AvrBs3 box to different promoter locations is always correlated with a new transcriptional start site. The analysis of naturally occurring Bs3 alleles revealed many pepper accessions that encode a nonfunctional Bs3 variant. These accessions showed no apparent abnormalities, supporting the supposition that Bs3 functions only in disease resistance and not in other developmental or physiological processes. Plant pathogenic microbes deliver a cocktail of effector proteins into the host cytoplasm that collectively promote microbial growth (Kamoun, 2006; Block et al., 2008; Göhre and Robatzek, 2008; Cunnac et al., 2009; Hogenhout et al., 2009). Many effector proteins were first termed avirulence (Avr) proteins because their presence evoked a hypersensitive response (HR) in plants expressing a matching resistance (R) gene (Grant et al., 2006; Bent and Mackey, 2007). Although the appearance of an HR often correlates with disease resistance, its causal role in plant immunity has not been fully elucidated (Greenberg and Yao, 2004). Avr proteins were identified initially as activators of the plant immune reaction, but many were later found to contribute to pathogen virulence on host plants that lack a corresponding R gene (Jones and Dangl, 2006). Meanwhile, the in planta function of a number of effectors has been studied at the molecular level, and some seem to function as enzymes (Mudgett, 2005; Abramovitch et al., 2006; Chisholm et al., 2006). Yet, some effectors of the bacterial genus Xanthomonas harbor nuclear localization signals and a transcriptional activation domain (Van den Ackerveken et al., 1996; Yang et al., 2000; Szurek et al., 2001) and have been termed transcription activator-like effector proteins (TALes; Kay and Bonas, 2009). The prototype TALe, AvrBs3, was identified from Xanthomonas campestris pv vesicatoria (Xcv) based on its avirulence activity in pepper (Capsicum annuum; Bonas et al., 1989) and was later shown to contribute to bacterial virulence in susceptible pepper genotypes (Wichmann and Bergelson, 2004). The most characteristic structural feature of TALes is a variable number of tandemly arranged, nearly perfect copies of a 34-amino acid motif that mediates binding of the TALe AvrBs3 to host target promoters (Kay et al., 2007). Although TALes are generally highly homologous to each other, their activity in plants is subject to exquisite specificity (Schornack et al., 2006). For example, the pepper Bs3 and the tomato (Solanum lycopersicum) Bs4 R genes mediate recognition of the 96.6% identical Xcv AvrBs3 and AvrBs4 proteins, respectively (Ballvora et al., 2001; Schornack et al., 2005; Römer et al., 2007). Tomato Bs4 is expressed constitutively at low levels and encodes a nucleotide-binding Leu-rich repeat-type R protein (Schornack et al., 2004, 2005). By contrast, the pepper Bs3 gene is transcriptionally induced by AvrBs3 and encodes a YUCCA-like flavin monooxygenase (Römer et al., 2007). Thus, Bs3- and Bs4-mediated recognition are mechanistically distinct despite the fact that they mediate recognition of almost identical effector proteins. Previously, we showed that the pepper Bs3 gene mediates recognition of AvrBs3 but not of its deletion derivative AvrBs3Δrep16, which lacks repeat units 11 to 14 (Herbers et al., 1992). Reciprocally, the Bs3-E allele mediates recognition of AvrBs3Δrep16 but not of AvrBs3. Recent studies demonstrated that AvrBs3 and AvrBs3Δrep16 specifically activate the matching Bs3 and Bs3-E promoters, respectively (Römer et al., 2007). The Bs3-E gene is an allele of Bs3 that carries a 13-bp insertion in its promoter compared with Bs3. Comparison of the Bs3 and other AvrBs3-inducible promoters from pepper revealed a conserved DNA element, the so-called UPA (for up-regulated by AvrBs3) box (Kay et al., 2007; Römer et al., 2007). Notably, the Bs3-E-specific 13-bp insertion is located within the UPA box of the Bs3 promoter. Electrophoretic mobility shift assays (EMSA) showed that AvrBs3 has a higher affinity to the Bs3 promoter as compared with the Bs3-E promoter. Yet, in EMSA, AvrBs3Δrep16 also had a higher affinity for the Bs3 promoter than for the Bs3-E promoter. Thus, it seemed that promoter binding of AvrBs3 or AvrBs3Δrep16 is not the basis for promoter activation specificity (Römer et al., 2007). In order to gain further insight into the molecular basis of TALe specificity and Bs3-mediated resistance, we have now carried out site-directed mutagenesis to define the exact boundaries of the UPA boxes of the Bs3 promoter (herein designated UPA AvrBs3 box) and the Bs3-E promoter (herein designated UPA AvrBs3Δrep16 box). We present new EMSA results that demonstrate that promoter binding is indeed the basis for promoter activation specificity by TALes. Finally, the analysis of a collection of naturally occurring Bs3 alleles revealed that pepper accessions encoding nonfunctional Bs3 variants are phenotypically normal. These data support the role of Bs3 exclusively in disease resistance. RESULTS Mapping the Promoter Sequences Used for Activation by AvrBs3 and AvrBs3Δrep16 Previously, we showed that transcript abundance of the pepper Bs3 resistance gene is increased 24 h post infection with Xcv strains that deliver AvrBs3 (Römer et al., 2007). We have now carried out semiquantitative reverse transcription (RT)-PCR to monitor Bs3 transcript accumulation on Xcv-infected leaves in a time-course experiment. As shown in Figure 1A Figure 1. Open in new tabDownload slide Analysis of Bs3 and Bs3-E transcript abundance. Transcript abundance was determined by semiquantitative RT-PCR. The constitutively expressed gene elongation factor 1α (EF1α) served as a normalization control. A, An increase of Bs3 transcript abundance was detectable 6 h post infection (hpi) with Xcv strain 85-10 expressing avrBs3. Leaves of 5- to 6-week-old plants of pepper cv ECW-30R (Bs3 genotype) were inoculated with Xcv (OD600 = 0.4) via blunt syringe. Inoculated leaf tissue was harvested at 0, 3, 6, 9, 12, 18, and 24 h post infection, and RNA was extracted and reverse transcribed into cDNA. To determine the earliest time point at which Bs3 transcript was detectable, 35 and 40 PCR cycles were carried out. B, Bs3 and Bs3-E transcripts were detectable only after infection with Xcv strains expressing the matching Avr protein. Tissue-specific analysis of Bs3 and Bs3-E transcripts was performed on cDNA from uninfected leaf, flower, fruit, and root tissue of pepper ECW-30R (Bs3) and ECW (Bs3-E). As a positive control for RT-PCR, we used cDNA derived from ECW-30R and ECW leaves that were inoculated with avrBs3- and avrBs3Δrep16-expressing Xcv strains (OD600 = 0.4), respectively. Figure 1. Open in new tabDownload slide Analysis of Bs3 and Bs3-E transcript abundance. Transcript abundance was determined by semiquantitative RT-PCR. The constitutively expressed gene elongation factor 1α (EF1α) served as a normalization control. A, An increase of Bs3 transcript abundance was detectable 6 h post infection (hpi) with Xcv strain 85-10 expressing avrBs3. Leaves of 5- to 6-week-old plants of pepper cv ECW-30R (Bs3 genotype) were inoculated with Xcv (OD600 = 0.4) via blunt syringe. Inoculated leaf tissue was harvested at 0, 3, 6, 9, 12, 18, and 24 h post infection, and RNA was extracted and reverse transcribed into cDNA. To determine the earliest time point at which Bs3 transcript was detectable, 35 and 40 PCR cycles were carried out. B, Bs3 and Bs3-E transcripts were detectable only after infection with Xcv strains expressing the matching Avr protein. Tissue-specific analysis of Bs3 and Bs3-E transcripts was performed on cDNA from uninfected leaf, flower, fruit, and root tissue of pepper ECW-30R (Bs3) and ECW (Bs3-E). As a positive control for RT-PCR, we used cDNA derived from ECW-30R and ECW leaves that were inoculated with avrBs3- and avrBs3Δrep16-expressing Xcv strains (OD600 = 0.4), respectively. , Bs3 transcript was detectable as early as 6 h post infection and peaks at 18 h post infection. Expression of Bs3 and Bs3-E was also studied in uninfected leaf, flower, fruit, and root tissue. However, we were unable to detect Bs3 or Bs3-E transcript in uninfected plant tissue (Fig. 1B). To define the minimal Bs3 and Bs3-E promoter regions, we generated progressive 5′ deletions and fused these to the Bs3 and Bs3-E coding sequences (cds). Functionality and specificity of corresponding T-DNA constructs were tested in leaves of Nicotiana benthamiana by Agrobacterium tumefaciens-mediated codelivery of a cauliflower mosaic virus 35S (35S) promoter-driven avrBs3 (35S:avrBs3) or avrBs3Δrep16 (35S:avrBs3Δrep16) gene, respectively (Fig. 2). Figure 2. Open in new tabDownload slide The UPA boxes of the Bs3 and Bs3-E promoters are crucial to their inducibility by the matching AvrBs3 and AvrBs3Δrep16 proteins. To define the minimal Bs3 and Bs3-E promoters, progressive 5′ deletions of the Bs3 promoter (343 Bs3, 166 Bs3, and 90 Bs3; A and B) and the Bs3-E promoter (356 Bs3-E, 179 Bs3-E, and 90 Bs3-E; C and D) were fused to the Bs3 and Bs3-E cds, respectively. The Bs3 and Bs3-E promoter constructs were delivered as denoted into N. benthamiana leaves via A. tumefaciens (OD600 = 0.8) together with constructs containing the 35S promoter-driven avrBs3Δrep16 (left side of the leaf; + avrBs3Δrep16) or avrBs3 (right side of the leaf; + avrBs3) genes (B and D). In addition, avrBs3 and avrBs3Δrep16 were expressed individually in the absence of a Bs3 or a Bs3-E promoter construct (avrBs3* or avrBs3Δrep16*). Dashed lines mark inoculated areas. Four days post infiltration (dpi), leaves were harvested and cleared with ethanol to visualize the HR (dark areas in B and D). Schemes in A and C show the length of the Bs3 promoter (A) and the Bs3-E promoter (C) deletions examined. Promoter deletion constructs are designated according to the length of the respective promoter fragment with respect to the ATG start codon and are displayed to scale. The total length of promoter regions and the 5′ UTR of Bs3 and Bs3-E are 1,023 and 1,036 bp, respectively. All similarly sized Bs3 and Bs3-E promoters are identical in their 5′ and 3′ ends but differ in size due to the presence of a 13-bp insertion/deletion polymorphism that is present in the Bs3-E promoter and absent from the Bs3 promoter. Hatched boxes represent the promoter and the 5′ UTR of the genes. Small white and black boxes represent the UPA boxes from the Bs3 (UPA AvrBs3) and Bs3-E (UPA AvrBs3Δrep16) promoters, respectively. Note that the Bs3 and Bs3-E promoters differ only within these UPA boxes but are otherwise identical and therefore are displayed in identical color. Gray boxes represent the cds of the Bs3 and Bs3-E genes. + and – indicate the presence and absence of an HR in N. benthamiana (A and C). Figure 2. Open in new tabDownload slide The UPA boxes of the Bs3 and Bs3-E promoters are crucial to their inducibility by the matching AvrBs3 and AvrBs3Δrep16 proteins. To define the minimal Bs3 and Bs3-E promoters, progressive 5′ deletions of the Bs3 promoter (343 Bs3, 166 Bs3, and 90 Bs3; A and B) and the Bs3-E promoter (356 Bs3-E, 179 Bs3-E, and 90 Bs3-E; C and D) were fused to the Bs3 and Bs3-E cds, respectively. The Bs3 and Bs3-E promoter constructs were delivered as denoted into N. benthamiana leaves via A. tumefaciens (OD600 = 0.8) together with constructs containing the 35S promoter-driven avrBs3Δrep16 (left side of the leaf; + avrBs3Δrep16) or avrBs3 (right side of the leaf; + avrBs3) genes (B and D). In addition, avrBs3 and avrBs3Δrep16 were expressed individually in the absence of a Bs3 or a Bs3-E promoter construct (avrBs3* or avrBs3Δrep16*). Dashed lines mark inoculated areas. Four days post infiltration (dpi), leaves were harvested and cleared with ethanol to visualize the HR (dark areas in B and D). Schemes in A and C show the length of the Bs3 promoter (A) and the Bs3-E promoter (C) deletions examined. Promoter deletion constructs are designated according to the length of the respective promoter fragment with respect to the ATG start codon and are displayed to scale. The total length of promoter regions and the 5′ UTR of Bs3 and Bs3-E are 1,023 and 1,036 bp, respectively. All similarly sized Bs3 and Bs3-E promoters are identical in their 5′ and 3′ ends but differ in size due to the presence of a 13-bp insertion/deletion polymorphism that is present in the Bs3-E promoter and absent from the Bs3 promoter. Hatched boxes represent the promoter and the 5′ UTR of the genes. Small white and black boxes represent the UPA boxes from the Bs3 (UPA AvrBs3) and Bs3-E (UPA AvrBs3Δrep16) promoters, respectively. Note that the Bs3 and Bs3-E promoters differ only within these UPA boxes but are otherwise identical and therefore are displayed in identical color. Gray boxes represent the cds of the Bs3 and Bs3-E genes. + and – indicate the presence and absence of an HR in N. benthamiana (A and C). Functional Bs3 and Bs3-E promoter deletion derivatives were expected to be transcriptionally induced by AvrBs3 and AvrBs3Δrep16, respectively, resulting in the expression of the Bs3 and Bs3-E proteins and triggering an HR. Data observed with overexpression of bacterial effectors in a heterologous system generally need to be treated with caution. Yet, we have previously demonstrated that Bs3-mediated recognition specificity is unaffected even if highly related TALes are expressed to high levels in a heterologous system (Schornack et al., 2005; Römer et al., 2007). Furthermore, our assay reflects the natural situation where activation of the Bs3 promoter results in the HR (Römer et al., 2007). Using A. tumefaciens-mediated delivery, it was found that Bs3 promoter fragments containing 1,023, 343, or 166 bp upstream of the ATG start codon triggered the HR in combination with 35S:avrBs3 but not with 35S:avrBs3Δrep16 (Fig. 2B). By contrast, a 90-bp Bs3 promoter fragment, which lacks the previously predicted UPA AvrBs3 box (TATATAAACCN2-3CC; Kay et al., 2007), was not able to trigger the HR in N. benthamiana when codelivered with the 35S:avrBs3 construct. In addition, we tested a set of equivalent Bs3-E promoter deletion derivatives (Fig. 2C; note that similarly sized Bs3 and Bs3-E promoter deletions are identical at their 5′ and 3′ termini but differ in size due to a 13-bp insertion/deletion polymorphism). As observed with the Bs3 promoter deletion derivatives, a Bs3-E promoter deletion that contains only 90 bp upstream of the ATG start codon (Fig. 2D; construct 90 Bs3-E) did not trigger HR when codelivered with a 35S:avrBs3Δrep16 construct. The exact location of the UPA AvrBs3Δrep16 box of the Bs3-E promoter has not been defined; however, it likely will include the 13-bp insertion/deletion polymorphism present in the Bs3-E promoter and absent from the Bs3 promoter, because this is the only difference between the two promoters. Consistent with this region playing a role in activation is the result that the uninducible promoter derivatives 90 Bs3 and 90 Bs3-E do not contain the regions that are polymorphic between Bs3 and Bs3-E. To determine if other sequences, apart from the predicted UPA AvrBs3 box, are crucial to AvrBs3-mediated transcriptional activation, we generated a set of Bs3 promoter mutants carrying different deletions between the ATG start codon and the UPA AvrBs3 box (Fig. 3A). Figure 3. Open in new tabDownload slide Extended Bs3 promoter deletions 3′ of the UPA AvrBs3 box suppress the AvrBs3-triggered and Bs3-mediated HR. To analyze if sequences apart from the UPA AvrBs3 box are needed for AvrBs3-mediated activation of the Bs3 promoter, regions between the UPA AvrBs3 box and the ATG start codon were deleted. A, Graphical display of Bs3 promoter deletion constructs. The designation of the deletions refers to the first and last deleted bp in the given promoter starting from the first bp 5′ of the ATG start codon. Deletions are displayed as white boxes, and their size is indicated in bp (triangles). The Bs3 promoter region, the UPA AvrBs3 box, and the Bs3 cds are displayed as hatched, black, and gray boxes, respectively. The indicated 5′ UTR refers to the Bs3 wild-type gene. Arrows above the boxes mark the TSS of the given promoter. The scale at the bottom indicates the distances with respect to the ATG start codon. With the exception of the Bs3 cds, all elements are drawn to scale. All deletions were fused individually in front of the Bs3 cds. + and – indicate the presence and absence of an HR in N. benthamiana for each construct when being codelivered with a 35S:avrBs3 T-DNA. The sequences, deletions, and TSS of all promoters are also provided as Supplemental Data Set S1. B, Functional analysis of Bs3 promoter deletion constructs. Individual deletion constructs were delivered together with 35S:avrBs3 via A. tumefaciens (OD600 = 0.8) into N. benthamiana leaves. Dashed lines mark the inoculated areas. The leaves were harvested at 4 dpi and cleared with ethanol to visualize the HR (dark areas). Figure 3. Open in new tabDownload slide Extended Bs3 promoter deletions 3′ of the UPA AvrBs3 box suppress the AvrBs3-triggered and Bs3-mediated HR. To analyze if sequences apart from the UPA AvrBs3 box are needed for AvrBs3-mediated activation of the Bs3 promoter, regions between the UPA AvrBs3 box and the ATG start codon were deleted. A, Graphical display of Bs3 promoter deletion constructs. The designation of the deletions refers to the first and last deleted bp in the given promoter starting from the first bp 5′ of the ATG start codon. Deletions are displayed as white boxes, and their size is indicated in bp (triangles). The Bs3 promoter region, the UPA AvrBs3 box, and the Bs3 cds are displayed as hatched, black, and gray boxes, respectively. The indicated 5′ UTR refers to the Bs3 wild-type gene. Arrows above the boxes mark the TSS of the given promoter. The scale at the bottom indicates the distances with respect to the ATG start codon. With the exception of the Bs3 cds, all elements are drawn to scale. All deletions were fused individually in front of the Bs3 cds. + and – indicate the presence and absence of an HR in N. benthamiana for each construct when being codelivered with a 35S:avrBs3 T-DNA. The sequences, deletions, and TSS of all promoters are also provided as Supplemental Data Set S1. B, Functional analysis of Bs3 promoter deletion constructs. Individual deletion constructs were delivered together with 35S:avrBs3 via A. tumefaciens (OD600 = 0.8) into N. benthamiana leaves. Dashed lines mark the inoculated areas. The leaves were harvested at 4 dpi and cleared with ethanol to visualize the HR (dark areas). The Bs3 promoter deletions were fused in front of the Bs3 cds and were delivered into N. benthamiana leaves via A. tumefaciens T-DNA transfer together with a 35S:avrBs3 gene (Fig. 3B). A Bs3 promoter deletion derivative that lacks a 44-bp region 3′ of the UPA AvrBs3 box (Bs3Δ56-99; Fig. 3A) was still capable of triggering an AvrBs3-dependent HR. By contrast, Bs3 promoter mutants with deletions larger than 59 bp were not functional. These data suggest that a minimum distance between the UPA AvrBs3 box and the ATG start codon is needed in order to generate a transcript that encodes a functional Bs3 protein. We carried out RACE to determine the transcriptional start site (TSS) of all constructs. As anticipated, most constructs that did not mediate an AvrBs3-dependent HR produced transcripts that lack the ATG start codon of the Bs3 cds (constructs Bs3Δ1-99, Bs3Δ1-91, and Bs3Δ1-80; Fig. 3A; Supplemental Data Set S1). Construct Bs3Δ1-59, which also does not mediate an AvrBs3-dependent HR, produces a transcript that starts with the ATG start codon of the Bs3 cds but that contains no 5′ untranslated region (UTR). It seems likely that the absence of an HR with construct Bs3Δ1-59 is due to the lack of a 5′ UTR. Therefore, an essential aspect in the analysis of UPA box-containing promoters is to consider a spacing between the UPA box and the start of transcription. While a deletion analysis is capable of coarse characterization of regulatory regions in a promoter, linker-scanning mutagenesis permits a higher resolution identification of short, defined sequence motifs and their effect on promoter activity (McKnight and Kingsbury, 1982). We analyzed a set of 12 linker-scanning Bs3 promoter mutants. Each mutant contained a 15-bp insertion located between positions +31 and −191 relative to the TSS (Fig. 4, A and B Figure 4. Open in new tabDownload slide Linker-scanning mutagenesis of the Bs3 promoter region identifies the UPA AvrBs3 box as a functionally crucial element. A, Distribution of transposon footprints in the Bs3 promoter. To identify sequences that are crucial for AvrBs3-mediated transcriptional activation of the Bs3 promoter, linker-scanning mutagenesis was carried out. Twelve transposon footprint mutants (T; black triangles above or below the hatched box) were obtained, containing a 15-bp insertion. Numbering of the mutants refers to the nucleotide that is adjacent to the 3′ end of the respective transposon footprint with respect to the TSS. The Bs3 promoter region and the 5′ UTR are displayed as hatched areas. The transcriptional start site of the wild-type Bs3 gene is indicated with a black horizontal line. The UPA AvrBs3 box and the Bs3 cds are displayed as black and gray boxes, respectively. + and – indicate the presence and absence of an HR with each promoter construct in N. benthamiana leaves when codelivered with a 35S:avrBs3 T-DNA (+ avrBs3). avrBs3* denotes a tissue patch in which no Bs3 promoter construct but only a 35S:avrBs3 T-DNA was delivered. The 5′ UTR in the Bs3 wild-type promoter is 59 bp in size, and thus insertions T + 31 and T + 41 are located in the 5′ UTR. B, Insertions adjacent to or within the predicted UPA AvrBs3 box. The numbers above the nucleotide sequence indicate the distance with respect to the TSS. Boldface letters represent the predicted UPA AvrBs3 box. Sequences of transposon mutants are available as Supplemental Data Set S2. C, Functional analysis of linker-scanning-derived Bs3 promoter mutants. Each mutant was fused in front of the Bs3 cds and was delivered together with 35S:avrBs3 into N. benthamiana leaves via A. tumefaciens (OD600 = 0.8). Dashed lines mark the inoculated areas. Leaves were harvested at 4 dpi and cleared with ethanol to visualize the HR (dark areas). Figure 4. Open in new tabDownload slide Linker-scanning mutagenesis of the Bs3 promoter region identifies the UPA AvrBs3 box as a functionally crucial element. A, Distribution of transposon footprints in the Bs3 promoter. To identify sequences that are crucial for AvrBs3-mediated transcriptional activation of the Bs3 promoter, linker-scanning mutagenesis was carried out. Twelve transposon footprint mutants (T; black triangles above or below the hatched box) were obtained, containing a 15-bp insertion. Numbering of the mutants refers to the nucleotide that is adjacent to the 3′ end of the respective transposon footprint with respect to the TSS. The Bs3 promoter region and the 5′ UTR are displayed as hatched areas. The transcriptional start site of the wild-type Bs3 gene is indicated with a black horizontal line. The UPA AvrBs3 box and the Bs3 cds are displayed as black and gray boxes, respectively. + and – indicate the presence and absence of an HR with each promoter construct in N. benthamiana leaves when codelivered with a 35S:avrBs3 T-DNA (+ avrBs3). avrBs3* denotes a tissue patch in which no Bs3 promoter construct but only a 35S:avrBs3 T-DNA was delivered. The 5′ UTR in the Bs3 wild-type promoter is 59 bp in size, and thus insertions T + 31 and T + 41 are located in the 5′ UTR. B, Insertions adjacent to or within the predicted UPA AvrBs3 box. The numbers above the nucleotide sequence indicate the distance with respect to the TSS. Boldface letters represent the predicted UPA AvrBs3 box. Sequences of transposon mutants are available as Supplemental Data Set S2. C, Functional analysis of linker-scanning-derived Bs3 promoter mutants. Each mutant was fused in front of the Bs3 cds and was delivered together with 35S:avrBs3 into N. benthamiana leaves via A. tumefaciens (OD600 = 0.8). Dashed lines mark the inoculated areas. Leaves were harvested at 4 dpi and cleared with ethanol to visualize the HR (dark areas). ; Supplemental Data Set S2). The Bs3 promoter mutants were fused to the Bs3 cds and were delivered via A. tumefaciens T-DNA transfer together with the 35S:avrBs3 construct into N. benthamiana leaves (Fig. 4C) to test for HR induction. Two of the 12 mutants (T−46 and T−55) no longer triggered an HR. The insertions that affected Bs3 promoter function are within (T−55) or adjacent to (T−46) the previously defined UPA AvrBs3 box (Fig. 4B) that spans a region from −47 to −61 bp. Given that the T−46 insertion is located adjacent to but not within the previously predicted UPA AvrBs3 box, the functionally relevant nucleotides of the box likely extend farther into the 3′ region of the promoter. Given that all insertions that affected Bs3 promoter function were located at or in the UPA AvrBs3 box, it seems likely that this is the only sequence motif that is crucial to AvrBs3-mediated promoter activation. Recognition Specificity of the Bs3-E Promoter The Bs3 and Bs3-E promoters differ only by a 13-bp insertion (CTCTATTCCACTA) in Bs3-E compared with Bs3 (Römer et al., 2007). Although it is conceivable that this polymorphic area defines recognition specificity, the particular sequences that contribute to specificity in the Bs3-E promoter remained unclear. We hypothesized that the 13-bp insertion in the Bs3-E promoter contains the complete UPA AvrBs3Δrep16 box of the Bs3-E promoter. Here, we tested this hypothesis by placing the 13-bp insertion of the Bs3-E promoter into two different locations of the Bs3 promoter and fused the Bs3 promoter derivatives (Bs3 −20i and Bs3 +31i) in front of the Bs3 cds (Fig. 5A). Figure 5. Open in new tabDownload slide The Bs3-E promoter-specific 13-bp insertion exerts its function on recognition specificity in a position-dependent manner. A, The 13-bp insertion sequence (CTCTATTCCACTA) that is specific to the Bs3-E promoter (insertion at position −50 of the Bs3 promoter) was placed into the promoter and 5′ UTR of the Bs3 gene. Numbering of the corresponding mutant derivatives (Bs3 −20i and Bs3 +31i) refers to the nucleotide in the Bs3 promoter that is adjacent to the 3′ end of the 13-bp insertion. The nucleotide sequence highlighted in boldface letters represents the UPA AvrBs3Δrep16 box (see Fig. 7). The Bs3 promoter region and the 5′ UTR are displayed as a hatched box. The UPA AvrBs3 box and the Bs3 cds are displayed as black and gray boxes, respectively. B, Functional analysis of the Bs3 promoter derivatives Bs3 −20i and Bs3 +31i. Both Bs3 promoter derivatives were fused to the Bs3 cds and were codelivered with 35S promoter-driven avrBs3 (+ avrBs3) or avrBs3Δrep16 (+ avrBs3Δrep16) genes into N. benthamiana leaves via A. tumefaciens (OD600 = 0.8). Leaf areas in which only a 35S:avrBs3 or a 35S:avrBs3Δrep16 T-DNA was delivered are marked as avrBs3* or avrBs3Δrep16*, respectively. Dashed lines mark the inoculated areas. Leaves were harvested at 4 dpi and cleared with ethanol to visualize the HR (dark areas). Figure 5. Open in new tabDownload slide The Bs3-E promoter-specific 13-bp insertion exerts its function on recognition specificity in a position-dependent manner. A, The 13-bp insertion sequence (CTCTATTCCACTA) that is specific to the Bs3-E promoter (insertion at position −50 of the Bs3 promoter) was placed into the promoter and 5′ UTR of the Bs3 gene. Numbering of the corresponding mutant derivatives (Bs3 −20i and Bs3 +31i) refers to the nucleotide in the Bs3 promoter that is adjacent to the 3′ end of the 13-bp insertion. The nucleotide sequence highlighted in boldface letters represents the UPA AvrBs3Δrep16 box (see Fig. 7). The Bs3 promoter region and the 5′ UTR are displayed as a hatched box. The UPA AvrBs3 box and the Bs3 cds are displayed as black and gray boxes, respectively. B, Functional analysis of the Bs3 promoter derivatives Bs3 −20i and Bs3 +31i. Both Bs3 promoter derivatives were fused to the Bs3 cds and were codelivered with 35S promoter-driven avrBs3 (+ avrBs3) or avrBs3Δrep16 (+ avrBs3Δrep16) genes into N. benthamiana leaves via A. tumefaciens (OD600 = 0.8). Leaf areas in which only a 35S:avrBs3 or a 35S:avrBs3Δrep16 T-DNA was delivered are marked as avrBs3* or avrBs3Δrep16*, respectively. Dashed lines mark the inoculated areas. Leaves were harvested at 4 dpi and cleared with ethanol to visualize the HR (dark areas). We anticipated that Bs3 −20i and Bs3 +31i would be activated by both AvrBs3 and AvrBs3Δrep16. However, our data demonstrated that only 35S:avrBs3 triggered the HR in combination with Bs3 −20i and Bs3 +31i, whereas 35S:avrBs3Δrep16 failed to trigger the HR with these promoter derivatives (Fig. 5B). Based on these data, it appears that the 13-bp insertion represents a part of, but not the complete, UPA AvrBs3Δrep16 box. To further define the recognition specificity of the Bs3-E promoter, we placed insertions of one, two, or three nucleotides into the Bs3 promoter instead of the 13-bp sequence (Fig. 6A). Figure 6. Open in new tabDownload slide An insertion of 2 bp into the Bs3 promoter causes a change in recognition specificity. A, Schematic representation of insertions that were placed into the Bs3 promoter region. All insertions are indicated below the Bs3 promoter with yellow background. The 13-bp natural insertion of the Bs3-E promoter is displayed above the Bs3 promoter with yellow background. Insertions were placed into the Bs3 promoter at the position where the Bs3-E promoter contains the 13-bp insertion with respect to the Bs3 promoter. Red letters highlight insertions corresponding to the 5′ end of the 13-bp insertion in the Bs3-E promoter. Brown and blue letters display the predicted UPA AvrBs3 box of the Bs3 promoter and the experimentally defined UPA AvrBs3Δrep16 box of the Bs3-E promoter, respectively (see Fig. 7). Green and gray boxes represent the Bs3 promoter and the Bs3 cds, respectively. + and – indicate the presence and absence of an HR in N. benthamiana leaves upon codelivery of each construct with a 35S promoter-driven avrBs3 (AvrBs3) or avrBs3Δrep16 (AvrBs3Δrep16) gene, respectively. B, Functional analysis of Bs3 promoter insertion mutants. Representative mutants were expressed transiently in N. benthamiana leaves via A. tumefaciens (OD600 = 0.8) together with T-DNA constructs containing 35S promoter-driven avrBs3 (+ avrBs3) or avrBs3Δrep16 (+ avrBs3Δrep16) genes. Leaf areas in which only a 35S:avrBs3 or a 35S:avrBs3Δrep16 T-DNA was delivered are marked as avrBs3* or avrBs3Δrep16*, respectively. Dashed lines mark the inoculated areas. Leaves were harvested at 4 dpi and cleared with ethanol to visualize the HR (dark areas). Figure 6. Open in new tabDownload slide An insertion of 2 bp into the Bs3 promoter causes a change in recognition specificity. A, Schematic representation of insertions that were placed into the Bs3 promoter region. All insertions are indicated below the Bs3 promoter with yellow background. The 13-bp natural insertion of the Bs3-E promoter is displayed above the Bs3 promoter with yellow background. Insertions were placed into the Bs3 promoter at the position where the Bs3-E promoter contains the 13-bp insertion with respect to the Bs3 promoter. Red letters highlight insertions corresponding to the 5′ end of the 13-bp insertion in the Bs3-E promoter. Brown and blue letters display the predicted UPA AvrBs3 box of the Bs3 promoter and the experimentally defined UPA AvrBs3Δrep16 box of the Bs3-E promoter, respectively (see Fig. 7). Green and gray boxes represent the Bs3 promoter and the Bs3 cds, respectively. + and – indicate the presence and absence of an HR in N. benthamiana leaves upon codelivery of each construct with a 35S promoter-driven avrBs3 (AvrBs3) or avrBs3Δrep16 (AvrBs3Δrep16) gene, respectively. B, Functional analysis of Bs3 promoter insertion mutants. Representative mutants were expressed transiently in N. benthamiana leaves via A. tumefaciens (OD600 = 0.8) together with T-DNA constructs containing 35S promoter-driven avrBs3 (+ avrBs3) or avrBs3Δrep16 (+ avrBs3Δrep16) genes. Leaf areas in which only a 35S:avrBs3 or a 35S:avrBs3Δrep16 T-DNA was delivered are marked as avrBs3* or avrBs3Δrep16*, respectively. Dashed lines mark the inoculated areas. Leaves were harvested at 4 dpi and cleared with ethanol to visualize the HR (dark areas). The Bs3 promoter insertion mutants were fused to the Bs3 cds and delivered via A. tumefaciens together with 35S:avrBs3Δrep16 or 35S:avrBs3 into N. benthamiana leaves. We found that only the insertions Bs3+CT, Bs3+CTA, and Bs3+CTC triggered the HR in combination with AvrBs3Δrep16 (Fig. 6). By contrast, all other insertions of one, two, and three nucleotides did not result in an AvrBs3Δrep16-responsive promoter (Fig. 6B). We concluded that the CTC motif at the 5′ end of the 13-bp insertion in the Bs3-E promoter is part of the UPA AvrBs3Δrep16 box. Given that not only a CTC but also a CTA insertion triggered the AvrBs3Δrep16-dependent HR, it seems likely that the C nucleotide at the 3′ terminal end can be functionally replaced by an A nucleotide. Since the Bs3+CT insertion is followed by an A nucleotide, this promoter mutant also contains a CTA motif at the 5′ end of the insertion site. This probably explains why Bs3+CT is functionally identical to the Bs3+CTA construct. Systematic Substitution Mutagenesis of the Bs3 Promoter Previously, the sequence of the UPA AvrBs3 box was determined to be TATATAAACCN2-3CC by comparing three different AvrBs3-inducible promoters (Kay et al., 2007; Römer et al., 2007). However, in this study, we found that the Bs3 promoter mutant T−46, which contains an insertion adjacent to the 3′ end of the predicted UPA AvrBs3 box, was no longer capable of triggering an AvrBs3-dependent HR (Fig. 4). This suggested that the UPA AvrBs3 box extends farther into the 3′ direction than previously assumed. To determine the functionally relevant nucleotides of the UPA AvrBs3 box more precisely, we performed a systematic substitution mutagenesis of the Bs3 promoter to permutate each base from position −41 to −63 (Fig. 7A). Figure 7. Open in new tabDownload slide Substitution mutagenesis of the Bs3 and Bs3-E promoters permits exact containment of the corresponding UPA boxes. A, Generation of Bs3 and Bs3-E promoter substitution mutants. Individual nucleotides in the Bs3 or Bs3-E promoter were replaced by all three alternative nucleotides via site-directed mutagenesis. The top sequence is part of the Bs3 wild-type promoter. Nucleotide positions are relative to the TSS of the Bs3 promoter. The five sequences below refer to representative substitution mutants. Mutagenized nucleotides are displayed in red letters. In total, 69 Bs3 and 30 Bs3-E substitution mutants were generated. B, Functional analysis of Bs3 promoter mutants. The 69 different Bs3 promoter mutants and the wild-type Bs3 promoter were fused to the Bs3 cds and were codelivered with a 35S promoter-driven avrBs3 construct into N. benthamiana leaves via A. tumefaciens (OD600 = 0.8). The phenotypes were scored at 4 dpi. The colored boxes summarize the results of the phenotypic scoring. Nucleotide positions displayed in the top row are numbered relative to the TSS of Bs3. Green boxes display the wild-type Bs3 promoter that triggers the HR. Each light green box represents a substitution mutant that triggers the HR. Substitution mutants that did not trigger the HR are displayed in red. Boxes representing substitution mutants with an intermediate phenotype (weak HR) are displayed as light green and red triangles. Please note that the collection of green boxes represents one data point (the wild-type Bs3 promoter), while the other boxes represent the results that were observed with distinct substitution mutants. The UPA AvrBs3 box deduced from this analysis is shown at bottom in blue letters. The underlined sequence represents the previously predicted UPA AvrBs3 box (Kay et al., 2007). C, Functional analysis of Bs3-E promoter mutants. Thirty distinct Bs3-E promoter mutants and the wild-type Bs3-E promoter were fused to the Bs3-E cds and delivered with a 35S:avrBs3Δrep16 construct into N. benthamiana leaves via A. tumefaciens (OD600 = 0.8). The phenotypes were scored at 4 dpi. Color coding is as in B, with the difference that green boxes represent the Bs3-E promoter. Nucleotide positions that are displayed in the top row of boxes are relative to the TSS of Bs3-E. The deduced minimal UPA AvrBs3Δrep16 box of Bs3-E is displayed in brown letters. Yellow background marks the Bs3-E promoter-specific 13-bp insertion. The gray background marks corresponding positions in the Bs3 and Bs3-E promoters, which yielded almost identical results in the substitution mutant analysis. Figure 7. Open in new tabDownload slide Substitution mutagenesis of the Bs3 and Bs3-E promoters permits exact containment of the corresponding UPA boxes. A, Generation of Bs3 and Bs3-E promoter substitution mutants. Individual nucleotides in the Bs3 or Bs3-E promoter were replaced by all three alternative nucleotides via site-directed mutagenesis. The top sequence is part of the Bs3 wild-type promoter. Nucleotide positions are relative to the TSS of the Bs3 promoter. The five sequences below refer to representative substitution mutants. Mutagenized nucleotides are displayed in red letters. In total, 69 Bs3 and 30 Bs3-E substitution mutants were generated. B, Functional analysis of Bs3 promoter mutants. The 69 different Bs3 promoter mutants and the wild-type Bs3 promoter were fused to the Bs3 cds and were codelivered with a 35S promoter-driven avrBs3 construct into N. benthamiana leaves via A. tumefaciens (OD600 = 0.8). The phenotypes were scored at 4 dpi. The colored boxes summarize the results of the phenotypic scoring. Nucleotide positions displayed in the top row are numbered relative to the TSS of Bs3. Green boxes display the wild-type Bs3 promoter that triggers the HR. Each light green box represents a substitution mutant that triggers the HR. Substitution mutants that did not trigger the HR are displayed in red. Boxes representing substitution mutants with an intermediate phenotype (weak HR) are displayed as light green and red triangles. Please note that the collection of green boxes represents one data point (the wild-type Bs3 promoter), while the other boxes represent the results that were observed with distinct substitution mutants. The UPA AvrBs3 box deduced from this analysis is shown at bottom in blue letters. The underlined sequence represents the previously predicted UPA AvrBs3 box (Kay et al., 2007). C, Functional analysis of Bs3-E promoter mutants. Thirty distinct Bs3-E promoter mutants and the wild-type Bs3-E promoter were fused to the Bs3-E cds and delivered with a 35S:avrBs3Δrep16 construct into N. benthamiana leaves via A. tumefaciens (OD600 = 0.8). The phenotypes were scored at 4 dpi. Color coding is as in B, with the difference that green boxes represent the Bs3-E promoter. Nucleotide positions that are displayed in the top row of boxes are relative to the TSS of Bs3-E. The deduced minimal UPA AvrBs3Δrep16 box of Bs3-E is displayed in brown letters. Yellow background marks the Bs3-E promoter-specific 13-bp insertion. The gray background marks corresponding positions in the Bs3 and Bs3-E promoters, which yielded almost identical results in the substitution mutant analysis. The resulting 69 Bs3 promoter substitution mutants were fused in front of the Bs3 cds and were delivered via A. tumefaciens together with 35S:avrBs3Δrep16 or 35S:avrBs3 into N. benthamiana leaves. We identified three functionally distinct classes of substitution mutants. Forty-two substitution mutants were functionally identical to the Bs3 wild-type promoter and triggered an HR in combination with 35S:avrBs3 but not in combination with 35S:avrBs3Δrep16 (Fig. 7B, light green boxes). Twenty substitution mutants produced no HR in combination with either 35S:avrBs3 or 35S:avrBs3Δrep16 (Fig. 7B, red boxes). The remaining seven substitution mutants showed a reduced HR phenotype (Fig. 7B, boxes with red and light green triangles). This reduced HR phenotype was observed in multiple repetitions and thus is highly reproducible. Inspection of the Bs3 promoter substitution mutants revealed that functionally relevant nucleotides of the UPA AvrBs3 box span the region from −61 to −44. Thus, the experimentally defined UPA AvrBs3 box (TATATAAACCTAACCATC) extends three nucleotides farther in the 3′ direction as compared with the UPA AvrBs3 box described previously (TATATAAACCN2-3CC; Fig. 7; Kay et al., 2007). Based on the experimentally defined UPA AvrBs3 box, the Bs3 promoter mutant T−46 (Fig. 4B) contains an insertion within and not adjacent to the UPA AvrBs3 box and explains why this insertion mutant is incapable of triggering the AvrBs3-dependent HR (Fig. 4C). Another notable observation of our systematic mutagenesis is that the functional consequences of substitutions in the UPA AvrBs3 box differ depending on which position is mutated. For example, any mutation at position −61, −57, or −47 abolished the AvrBs3-mediated HR (Fig. 7B). By contrast, none of the substitutions at positions −59, −50, −49, −46, and −45 had a detectable effect on the AvrBs3-mediated HR. Substitution Mutagenesis of the Bs3-E Promoter To analyze the UPA AvrBs3Δrep16 box, we carried out substitution mutagenesis of selected positions within the Bs3-E promoter (Fig. 7C). The observation that Bs3 promoter mutants containing a 3-bp CTC insertion triggered an HR in combination with AvrBs3Δrep16 (Fig. 6) suggests that this sequence motif defines the 3′ end of the UPA AvrBs3Δrep16 box. In agreement with this hypothesis, we identified Bs3-E promoter substitution mutants in the CTC motif (positions −63 to −61) that do not mediate the HR in combination with AvrBs3Δrep16 (Fig. 7C). All Bs3-E promoter mutants that contained substitutions located in the 3′ direction of position −61 were functionally identical to the Bs3-E wild-type promoter. By contrast, many Bs3-E promoter mutants that contained substitutions located in the 5′ direction of the CTC motif lost the capability to mediate an AvrBs3Δrep16-triggered HR (Fig. 7C). Taken together, our data define the UPA AvrBs3Δrep16 box region as extending from position −74 to position −61 (TATATAAACCTCTC; Fig. 7C). Although it remains to be clarified if the UPA AvrBs3Δrep16 box extends at its 5′ end beyond position −74, it seems evident that the UPA AvrBs3 and the UPA AvrBs3Δrep16 boxes overlap in at least 11 nucleotides (Fig. 7C). In this context, it is notable that substitution mutations in corresponding positions in the UPA AvrBs3 and UPA AvrBs3Δrep16 boxes (−61 [Bs3] versus −74 [Bs3-E], −57 [Bs3] versus −70 [Bs3-E], and −53 [Bs3] versus −66 [Bs3-E]) have almost identical functional consequences. For example, all substitutions in position −57 of the Bs3 promoter and in position −70 of the Bs3-E promoter resulted in nonfunctional promoters (Fig. 7, B and C). Similarly, substitutions to an A nucleotide at position −53 of the Bs3 and position −66 of the Bs3-E promoter had no obvious functional consequences, while mutations to G or T resulted in nonfunctional Bs3 or Bs3-E promoter mutants. DNA-Binding Specificity of AvrBs3 and AvrBs3Δrep16 Previously, EMSA showed that AvrBs3 binds with high affinity to the Bs3 promoter and weakly to the Bs3-E promoter (Römer et al., 2007). Unexpectedly, EMSA showed also that AvrBs3Δrep16 had a higher affinity to the Bs3 promoter as compared with the Bs3-E promoter fragment (Römer et al., 2007). EMSA with AvrBs3Δrep16 and AvrBs3 was carried out with DNA probes of identical size that had identical sequence at their 3′ end but not at their 5′ end (Fig. 8A). Figure 8. Open in new tabDownload slide AvrBs3 and AvrBs3Δrep16 bind with high affinity to the Bs3 and Bs3-E promoters, respectively. A, Probes derived from the Bs3 and Bs3-E promoter sequences used in EMSA. The experimentally defined UPA AvrBs3 and UPA AvrBs3Δrep16 boxes of the Bs3 and Bs3-E promoters (see Fig. 7) are displayed as boldface black and gray letters, respectively. Numbering is relative to the TSS of Bs3 and Bs3-E, respectively. Hatched background indicates the 13-bp insertion of the Bs3-E promoter. Sequences of biotin-labeled probes are indicated by lines above and below the promoter sequences. Black and gray lines mark the Bs3 and Bs3-E promoter probe sequences used in these experiments; the dashed line marks the Bs3-E promoter probe sequence used previously (Römer et al., 2007). Note that the probe sequence used previously (dashed line) does not cover the entire UPA AvrBs3Δrep16 box (gray letters). B, EMSA of 100 fmol of biotin-labeled Bs3-derived (36 bp) and Bs3-E-derived (49 bp) probes incubated with 100, 250, and 500 fmol of GST:AvrBs3 (AvrBs3) and GST:AvrBs3Δrep16 (AvrBs3Δrep16) fusion proteins, respectively. Positions of bound and free probes are indicated: arrow, bound probe; asterisk, free probe. The top signals correspond to the slots. C, Coomassie Brilliant Blue-stained 8% SDS-PAGE. GST translational fusions to AvrBs3 and AvrBs3Δrep16 used for EMSA studies were expressed in Escherichia coli, purified, and quantified by Bradford analysis (Bradford, 1976). Subsequently, 1.5 and 3 μg of GST:AvrBs3, GST:AvrBs3Δrep16, and bovine serum albumin (BSA) standard were separated by SDS-PAGE and stained with Coomassie Brilliant Blue. Fragments of the expected size (GST:AvrBs3, 150.8 kD; AvrBs3Δrep16, 136.8 kD) are indicated by asterisks. Marker proteins (M) are indicated with their molecular masses in kD (PageRuler prestained protein ladder; Fermentas). D, EMSA competition assay of 100 fmol of biotin-labeled Bs3 (left) or Bs3-E (right) probe incubated with 500 fmol of GST:AvrBs3 (AvrBs3) or GST:AvrBs3Δrep16 (AvrBs3Δrep16), respectively. A molar excess of nonlabeled Bs3 and Bs3-E fragments of 25×, 50×, and 100× was used for competition. All experiments were repeated twice with similar results. Figure 8. Open in new tabDownload slide AvrBs3 and AvrBs3Δrep16 bind with high affinity to the Bs3 and Bs3-E promoters, respectively. A, Probes derived from the Bs3 and Bs3-E promoter sequences used in EMSA. The experimentally defined UPA AvrBs3 and UPA AvrBs3Δrep16 boxes of the Bs3 and Bs3-E promoters (see Fig. 7) are displayed as boldface black and gray letters, respectively. Numbering is relative to the TSS of Bs3 and Bs3-E, respectively. Hatched background indicates the 13-bp insertion of the Bs3-E promoter. Sequences of biotin-labeled probes are indicated by lines above and below the promoter sequences. Black and gray lines mark the Bs3 and Bs3-E promoter probe sequences used in these experiments; the dashed line marks the Bs3-E promoter probe sequence used previously (Römer et al., 2007). Note that the probe sequence used previously (dashed line) does not cover the entire UPA AvrBs3Δrep16 box (gray letters). B, EMSA of 100 fmol of biotin-labeled Bs3-derived (36 bp) and Bs3-E-derived (49 bp) probes incubated with 100, 250, and 500 fmol of GST:AvrBs3 (AvrBs3) and GST:AvrBs3Δrep16 (AvrBs3Δrep16) fusion proteins, respectively. Positions of bound and free probes are indicated: arrow, bound probe; asterisk, free probe. The top signals correspond to the slots. C, Coomassie Brilliant Blue-stained 8% SDS-PAGE. GST translational fusions to AvrBs3 and AvrBs3Δrep16 used for EMSA studies were expressed in Escherichia coli, purified, and quantified by Bradford analysis (Bradford, 1976). Subsequently, 1.5 and 3 μg of GST:AvrBs3, GST:AvrBs3Δrep16, and bovine serum albumin (BSA) standard were separated by SDS-PAGE and stained with Coomassie Brilliant Blue. Fragments of the expected size (GST:AvrBs3, 150.8 kD; AvrBs3Δrep16, 136.8 kD) are indicated by asterisks. Marker proteins (M) are indicated with their molecular masses in kD (PageRuler prestained protein ladder; Fermentas). D, EMSA competition assay of 100 fmol of biotin-labeled Bs3 (left) or Bs3-E (right) probe incubated with 500 fmol of GST:AvrBs3 (AvrBs3) or GST:AvrBs3Δrep16 (AvrBs3Δrep16), respectively. A molar excess of nonlabeled Bs3 and Bs3-E fragments of 25×, 50×, and 100× was used for competition. All experiments were repeated twice with similar results. Based on our data described above, it is now clear that the Bs3-E promoter probe did not span the complete UPA AvrBs3Δrep16 box, which likely accounts for the unexpected EMSA results (Römer et al., 2007). Therefore, we repeated the EMSA with Bs3 and Bs3-E promoter-derived probes that are sequence identical at their 3′ and 5′ ends but that, due to the 13-bp insertion in the Bs3-E promoter, are not identical in size. Hence, the new probe for the Bs3-E promoter is likely to contain the complete UPA AvrBs3Δrep16 box. As shown in Figure 8, GST:AvrBs3Δrep16 bound with high affinity to the Bs3-E-derived and with low affinity to Bs3-derived promoter fragments (Fig. 8, B and C). Similarly, GST:AvrBs3 binds with high and low affinity to the Bs3- and Bs3-E-derived promoter fragments, respectively. Competition assays with labeled Bs3-derived promoter fragments and nonlabeled Bs3- and Bs3-E-derived promoter fragments, and vice versa, further confirm that AvrBs3 has high affinity to the Bs3 promoter fragment and only low affinity to the Bs3-E promoter fragment (Fig. 8D). Competition assays for AvrBs3Δrep16 showed that it binds with high affinity to the Bs3-E promoter fragment and with low affinity to the Bs3 promoter fragment. Together, these data strongly suggest that, in contrast to our previous statements (Römer et al., 2007), specific binding of AvrBs3 or AvrBs3Δrep16 to their matching promoters is the basis for promoter activation specificity. A Bs3 Promoter with an Inverted UPA AvrBs3 Box Does Not Trigger the Bs3-Mediated HR To clarify if the UPA AvrBs3 box acts in a directional manner, we generated a Bs3 promoter derivative in which the UPA AvrBs3 box is replaced by the reverse-complement sequence (Fig. 9A). Figure 9. Open in new tabDownload slide A Bs3 promoter with an inversely orientated UPA AvrBs3 box does not trigger a Bs3-dependent HR. A, Part of the Bs3 promoter wild-type sequence (Bs3) and a derivative with an inverted UPA AvrBs3 box (Bs3 UPArev) are shown, highlighted with gray boxes. The experimentally determined UPA AvrBs3 box is shown in white letters (see Fig. 7). Numbering is relative to the TSS of Bs3. B, Functional analysis of the inverted box in the Bs3 promoter. The promoters depicted in A were fused to the Bs3 cds and delivered via A. tumefaciens (OD600 = 0.8) alone (asterisks) or together with a 35S promoter-driven avrBs3 gene (+ avrBs3) into N. benthamiana leaves. Dashed lines mark the inoculated areas. Leaves were harvested at 4 dpi and cleared with ethanol to visualize the HR (dark areas). Figure 9. Open in new tabDownload slide A Bs3 promoter with an inversely orientated UPA AvrBs3 box does not trigger a Bs3-dependent HR. A, Part of the Bs3 promoter wild-type sequence (Bs3) and a derivative with an inverted UPA AvrBs3 box (Bs3 UPArev) are shown, highlighted with gray boxes. The experimentally determined UPA AvrBs3 box is shown in white letters (see Fig. 7). Numbering is relative to the TSS of Bs3. B, Functional analysis of the inverted box in the Bs3 promoter. The promoters depicted in A were fused to the Bs3 cds and delivered via A. tumefaciens (OD600 = 0.8) alone (asterisks) or together with a 35S promoter-driven avrBs3 gene (+ avrBs3) into N. benthamiana leaves. Dashed lines mark the inoculated areas. Leaves were harvested at 4 dpi and cleared with ethanol to visualize the HR (dark areas). The corresponding Bs3 promoter mutant (Bs3 UPArev) was fused to the Bs3 cds and was delivered via A. tumefaciens together with 35S:avrBs3 into N. benthamiana leaves. The Bs3 UPArev construct did not trigger the HR in combination with AvrBs3, suggesting that the UPA AvrBs3 box acts in a directional manner (Fig. 9B). Functionality of the UPA AvrBs3 Box Is Independent of the Promoter Context The 3′ end of the UPA AvrBs3 box in the Bs3 promoter is located 44 bp upstream of the TSS and 102 bp upstream of the ATG start codon. We wondered if the UPA AvrBs3 box retains its functionality when moved to other positions within the Bs3 promoter. As starting material for this experiment, we used a Bs3 promoter substitution mutant (designated Bs3 UPAmut; the wild-type T nucleotide at position −61 is replaced by an A nucleotide) that, when fused in front of the Bs3 cds, did not trigger the HR in N. benthamiana leaves upon A. tumefaciens-mediated codelivery with 35S:avrBs3 (Fig. 10). Figure 10. Open in new tabDownload slide Functionality of the UPA AvrBs3 box is independent of its position and the promoter context. A, Graphical representation of promoter constructs tested. Mutations and insertions of UPA AvrBs3 boxes were made via site-directed mutagenesis. Green boxes represent the Bs3 and Bs3-E promoters. Yellow boxes represent the Bs4 promoter. Blue and brown boxes represent the UPA AvrBs3 and UPA AvrBs3Δrep16 boxes, respectively. Note that the Bs3 and Bs3-E promoters differ only within and adjacent to their UPA boxes but are otherwise identical and therefore displayed in identical color. A white vertical line within the UPA AvrBs3 box represents a substitution mutation (T→A at position −61; see Fig. 7B) within the UPA AvrBs3 box. All promoter elements are displayed to scale. The numbers and horizontal lines above the promoters provide a scale and denote the distances with respect to the ATG start codon. Arrows above the promoter boxes mark the TSS of the given promoter. Numbers below the arrows denote the distance between the 3′ end of the UPA AvrBs3 box and the respective TSS. Gray boxes represent the Bs3 cds. + and – indicate the presence and absence of an HR in N. benthamiana upon codelivery of each promoter construct with a 35S:avrBs3 construct. B, Functional analysis of Bs3 and Bs4 promoter derivatives. The promoter constructs depicted in A were codelivered together with a 35S:avrBs3 construct (+ avrBs3) into N. benthamiana leaves via A. tumefaciens (OD600 = 0.8). Dashed lines mark the inoculated areas. Four days after inoculation, the leaves were harvested and cleared with ethanol to better visualize the HR (dark areas). Figure 10. Open in new tabDownload slide Functionality of the UPA AvrBs3 box is independent of its position and the promoter context. A, Graphical representation of promoter constructs tested. Mutations and insertions of UPA AvrBs3 boxes were made via site-directed mutagenesis. Green boxes represent the Bs3 and Bs3-E promoters. Yellow boxes represent the Bs4 promoter. Blue and brown boxes represent the UPA AvrBs3 and UPA AvrBs3Δrep16 boxes, respectively. Note that the Bs3 and Bs3-E promoters differ only within and adjacent to their UPA boxes but are otherwise identical and therefore displayed in identical color. A white vertical line within the UPA AvrBs3 box represents a substitution mutation (T→A at position −61; see Fig. 7B) within the UPA AvrBs3 box. All promoter elements are displayed to scale. The numbers and horizontal lines above the promoters provide a scale and denote the distances with respect to the ATG start codon. Arrows above the promoter boxes mark the TSS of the given promoter. Numbers below the arrows denote the distance between the 3′ end of the UPA AvrBs3 box and the respective TSS. Gray boxes represent the Bs3 cds. + and – indicate the presence and absence of an HR in N. benthamiana upon codelivery of each promoter construct with a 35S:avrBs3 construct. B, Functional analysis of Bs3 and Bs4 promoter derivatives. The promoter constructs depicted in A were codelivered together with a 35S:avrBs3 construct (+ avrBs3) into N. benthamiana leaves via A. tumefaciens (OD600 = 0.8). Dashed lines mark the inoculated areas. Four days after inoculation, the leaves were harvested and cleared with ethanol to better visualize the HR (dark areas). Next, we inserted a wild-type UPA AvrBs3 box into Bs3 UPAmut at a distance of 293 bp (Bs3 UPA293) or 424 bp (Bs3 UPA424) upstream of the ATG start codon. Bs3 UPA293 and Bs3 UPA424 promoter constructs were fused in front of the Bs3 cds and were tested functionally via A. tumefaciens-mediated delivery. Both promoter constructs triggered an HR in N. benthamiana upon codelivery with 35S:avrBs3 (Fig. 10B). Thus, the UPA AvrBs3 box retains its functionality when moved within the Bs3 promoter. Next, we tested if the UPA AvrBs3 box would retain its function when moved to a promoter context different from the pepper Bs3 promoter. We used the promoter of the tomato R gene Bs4, which is transcribed constitutively at a low level (Schornack et al., 2005) and which does not trigger the HR when fused in front of the Bs3 cds (Fig. 10B). We placed the UPA AvrBs3 box and a mutated box variant into the Bs4 promoter (termed Bs4 UPA and Bs4 UPAmut, respectively) and fused these Bs4 promoter derivatives in front of the Bs3 cds. A. tumefaciens-mediated delivery of 35S:avrBs3 triggered an HR in N. benthamiana leaves upon codelivery of the Bs4 UPA but not the Bs4 UPAmut promoter construct. Thus, the UPA AvrBs3 box retains its functionality not only in different locations within the pepper Bs3 but also in the context of the tomato Bs4 promoter. The TSS of the Bs3 promoter is 44 bp downstream of the 3′ end of the UPA AvrBs3 box (Römer et al., 2007). We carried out RACE to determine the TSS generated in constructs that trigger the HR in combination with 35S:avrBs3 (Bs3 UPA293, Bs3 UPA424, and Bs4 UPA). As shown in Figure 10A, the TSS of Bs3 UPA293, Bs3 UPA424, and Bs4 UPA were 46, 41, and 42 bp downstream of the 3′ end of the respective UPA AvrBs3 boxes. Hence, the TSS in AvrBs3-inducible promoters appears to be dictated by the location of the UPA AvrBs3 box. Naturally Occurring Nonfunctional Bs3 Alleles To study natural diversity of Bs3 alleles, we examined accessions of the Capsicum species C. annuum, C. baccatum, and C. chinense. We determined the promoter sequences (244 bp 5′ of the ATG start codon) and cds of 49 Capsicum accessions, identifying 23 different haplotypes (Table I Table I. Functional analysis of naturally occurring Bs3 alleles Haplotype No.a . Promoter Type . Promoter, Comments . Coding Sequence, Commentsb . Functional Analysisc . . . . . . . . avrBs3 d . avrBs3Δrep16 e . 35S:cds f . Prom:Bs3cds g . 1* Bs3-E −/− +/+ + + 2 Bs3-E DNA polymorphisms also present in Bs3 cds −/n +/n 3‡ Bs3 +/+ −/− + − 4 Bs3-E Cds identical to Bs3 cds −/− +/+ 5 Bs3-E Loss of function due to mutation in second exon (1,093, CCT[P]→CTT[L]) −/− −/− − 6 Bs3-E −/− +/+ 7 Bs3-E −/− +/+ 8 Bs3-E 11-bp deletion in first exon (Δ595–605) results in a frameshift and thus an early stop codon (haplotypes 8–13) −/− −/− 9 Bs3-E −/− −/− − 10 Bs3-E −/− −/− 11 Bs3-E −/− −/− − 12 Bs3-E −/− −/− − 13 Bs3-E −/− −/− 14 Bs3-E −/− +/+ 15 Bs3-E −/− +/+ 16 Unique C→G (123) substitution in predicted UPA AvrBs3Δrep16 box Loss of function due to mutation in first exon (366, GGT[G]→GTC[S]) −/− −/− − − 17 Bs3-E n/− n/+ + 18 Identical to no. 19 C→G (161) substitution n/n n/n 19 Identical to no. 18 C→G (161) substitution −/− +/+ 20 Identical to no. 22 C→A (18) substitution Substitution in first exon (472, TAC[Y]→TAG[stop]) results in a stop codon −/− −/− 21 Bs3-E −/− −/+ + 22 Identical to no. 20 C→A (18) substitution n/− n/+ + + 23 Unique Deletion (Δ199) and C→T (121) substitution in predicted UPA AvrBs3Δrep16 box −/− −/− + − Haplotype No.a . Promoter Type . Promoter, Comments . Coding Sequence, Commentsb . Functional Analysisc . . . . . . . . avrBs3 d . avrBs3Δrep16 e . 35S:cds f . Prom:Bs3cds g . 1* Bs3-E −/− +/+ + + 2 Bs3-E DNA polymorphisms also present in Bs3 cds −/n +/n 3‡ Bs3 +/+ −/− + − 4 Bs3-E Cds identical to Bs3 cds −/− +/+ 5 Bs3-E Loss of function due to mutation in second exon (1,093, CCT[P]→CTT[L]) −/− −/− − 6 Bs3-E −/− +/+ 7 Bs3-E −/− +/+ 8 Bs3-E 11-bp deletion in first exon (Δ595–605) results in a frameshift and thus an early stop codon (haplotypes 8–13) −/− −/− 9 Bs3-E −/− −/− − 10 Bs3-E −/− −/− 11 Bs3-E −/− −/− − 12 Bs3-E −/− −/− − 13 Bs3-E −/− −/− 14 Bs3-E −/− +/+ 15 Bs3-E −/− +/+ 16 Unique C→G (123) substitution in predicted UPA AvrBs3Δrep16 box Loss of function due to mutation in first exon (366, GGT[G]→GTC[S]) −/− −/− − − 17 Bs3-E n/− n/+ + 18 Identical to no. 19 C→G (161) substitution n/n n/n 19 Identical to no. 18 C→G (161) substitution −/− +/+ 20 Identical to no. 22 C→A (18) substitution Substitution in first exon (472, TAC[Y]→TAG[stop]) results in a stop codon −/− −/− 21 Bs3-E −/− −/+ + 22 Identical to no. 20 C→A (18) substitution n/− n/+ + + 23 Unique Deletion (Δ199) and C→T (121) substitution in predicted UPA AvrBs3Δrep16 box −/− −/− + − a Haplotypes 1 and 3 represent the Bs3-E (*) and Bs3 (‡) alleles, respectively. Capsicum accessions corresponding to the haplotypes are provided in Supplemental Figure S1. b Polymorphic nucleotides are underlined, and the encoded amino acids are given in square brackets. Numbering of polymorphisms refers to Supplemental Figures S1 and S2. c Functionality of the Bs3 alleles was analyzed via Xcv inoculation of the pepper accessions and via A. tumefaciens-mediated transient expression in N. benthamiana. Results are displayed in columns d and e , always with the data observed in pepper first and the data observed in N. benthamiana second. The presence or absence of an HR is indicated by + or −, respectively. n, Not tested. Capsicum accessions were tested with Xcv strains expressing either avrBs3 (d) or avrBs3Δrep16 (e). For A. tumefaciens-mediated transient expression in N. benthamiana, the cloned Bs3 alleles (promoter and cds) were delivered in combination with a cauliflower mosaic virus 35S promoter-driven avrBs3 (d) or avrBs3Δrep16 (e) construct. f The cds of the given Bs3 allele was expressed under the control of the 35S promoter (35S:cds). g The promoter of a given Bs3 allele was fused in front of the Bs3 wild-type cds and was delivered into N. benthamiana in combination with a 35S-driven avrBs3Δrep16 T-DNA (Prom:Bs3 cds). Open in new tab Table I. Functional analysis of naturally occurring Bs3 alleles Haplotype No.a . Promoter Type . Promoter, Comments . Coding Sequence, Commentsb . Functional Analysisc . . . . . . . . avrBs3 d . avrBs3Δrep16 e . 35S:cds f . Prom:Bs3cds g . 1* Bs3-E −/− +/+ + + 2 Bs3-E DNA polymorphisms also present in Bs3 cds −/n +/n 3‡ Bs3 +/+ −/− + − 4 Bs3-E Cds identical to Bs3 cds −/− +/+ 5 Bs3-E Loss of function due to mutation in second exon (1,093, CCT[P]→CTT[L]) −/− −/− − 6 Bs3-E −/− +/+ 7 Bs3-E −/− +/+ 8 Bs3-E 11-bp deletion in first exon (Δ595–605) results in a frameshift and thus an early stop codon (haplotypes 8–13) −/− −/− 9 Bs3-E −/− −/− − 10 Bs3-E −/− −/− 11 Bs3-E −/− −/− − 12 Bs3-E −/− −/− − 13 Bs3-E −/− −/− 14 Bs3-E −/− +/+ 15 Bs3-E −/− +/+ 16 Unique C→G (123) substitution in predicted UPA AvrBs3Δrep16 box Loss of function due to mutation in first exon (366, GGT[G]→GTC[S]) −/− −/− − − 17 Bs3-E n/− n/+ + 18 Identical to no. 19 C→G (161) substitution n/n n/n 19 Identical to no. 18 C→G (161) substitution −/− +/+ 20 Identical to no. 22 C→A (18) substitution Substitution in first exon (472, TAC[Y]→TAG[stop]) results in a stop codon −/− −/− 21 Bs3-E −/− −/+ + 22 Identical to no. 20 C→A (18) substitution n/− n/+ + + 23 Unique Deletion (Δ199) and C→T (121) substitution in predicted UPA AvrBs3Δrep16 box −/− −/− + − Haplotype No.a . Promoter Type . Promoter, Comments . Coding Sequence, Commentsb . Functional Analysisc . . . . . . . . avrBs3 d . avrBs3Δrep16 e . 35S:cds f . Prom:Bs3cds g . 1* Bs3-E −/− +/+ + + 2 Bs3-E DNA polymorphisms also present in Bs3 cds −/n +/n 3‡ Bs3 +/+ −/− + − 4 Bs3-E Cds identical to Bs3 cds −/− +/+ 5 Bs3-E Loss of function due to mutation in second exon (1,093, CCT[P]→CTT[L]) −/− −/− − 6 Bs3-E −/− +/+ 7 Bs3-E −/− +/+ 8 Bs3-E 11-bp deletion in first exon (Δ595–605) results in a frameshift and thus an early stop codon (haplotypes 8–13) −/− −/− 9 Bs3-E −/− −/− − 10 Bs3-E −/− −/− 11 Bs3-E −/− −/− − 12 Bs3-E −/− −/− − 13 Bs3-E −/− −/− 14 Bs3-E −/− +/+ 15 Bs3-E −/− +/+ 16 Unique C→G (123) substitution in predicted UPA AvrBs3Δrep16 box Loss of function due to mutation in first exon (366, GGT[G]→GTC[S]) −/− −/− − − 17 Bs3-E n/− n/+ + 18 Identical to no. 19 C→G (161) substitution n/n n/n 19 Identical to no. 18 C→G (161) substitution −/− +/+ 20 Identical to no. 22 C→A (18) substitution Substitution in first exon (472, TAC[Y]→TAG[stop]) results in a stop codon −/− −/− 21 Bs3-E −/− −/+ + 22 Identical to no. 20 C→A (18) substitution n/− n/+ + + 23 Unique Deletion (Δ199) and C→T (121) substitution in predicted UPA AvrBs3Δrep16 box −/− −/− + − a Haplotypes 1 and 3 represent the Bs3-E (*) and Bs3 (‡) alleles, respectively. Capsicum accessions corresponding to the haplotypes are provided in Supplemental Figure S1. b Polymorphic nucleotides are underlined, and the encoded amino acids are given in square brackets. Numbering of polymorphisms refers to Supplemental Figures S1 and S2. c Functionality of the Bs3 alleles was analyzed via Xcv inoculation of the pepper accessions and via A. tumefaciens-mediated transient expression in N. benthamiana. Results are displayed in columns d and e , always with the data observed in pepper first and the data observed in N. benthamiana second. The presence or absence of an HR is indicated by + or −, respectively. n, Not tested. Capsicum accessions were tested with Xcv strains expressing either avrBs3 (d) or avrBs3Δrep16 (e). For A. tumefaciens-mediated transient expression in N. benthamiana, the cloned Bs3 alleles (promoter and cds) were delivered in combination with a cauliflower mosaic virus 35S promoter-driven avrBs3 (d) or avrBs3Δrep16 (e) construct. f The cds of the given Bs3 allele was expressed under the control of the 35S promoter (35S:cds). g The promoter of a given Bs3 allele was fused in front of the Bs3 wild-type cds and was delivered into N. benthamiana in combination with a 35S-driven avrBs3Δrep16 T-DNA (Prom:Bs3 cds). Open in new tab ; Supplemental Figs. S1 and S2; Supplemental Data Sets S3 and S4). The Bs3-E haplotype (defined as haplotype 1) was the most prevalent, found in nine of 49 sequenced accessions. Within the promoter region, 39 accessions constituting 14 haplotypes were identical to the Bs3-E haplotype. By contrast, we identified only two accessions that were sequence identical to the Bs3 gene in promoter and cds (Table I). We identified two haplotypes (nos. 16 and 23) that carried mutations in the predicted UPA AvrBs3Δrep16 box (Supplemental Fig. S5). Infection experiments showed that neither avrBs3-expressing nor avrBs3Δrep16-expressing Xcv strains triggered the HR in pepper accessions corresponding to haplotypes 16 and 23 (Table I). We PCR amplified and cloned the two corresponding Bs3 alleles (promoter and cds) and tested their functionality via A. tumefaciens-mediated transient expression in N. benthamiana leaves. Both alleles failed to trigger the HR when codelivered with 35S:avrBs3Δrep16. We also cloned the promoters of haplotypes 16 and 23 in front of the Bs3-E cds to test if the lack of functionality of these alleles is due to promoter polymorphisms. Indeed, transcriptional fusions of the Bs3-E cds to the promoters of haplotypes 16 and 23 did not mediate recognition of AvrBs3Δrep16. In a reciprocal experiment, we fused the strong constitutive 35S promoter in front of the cds of haplotypes 16 and 23. Functional analysis demonstrated that the cds of haplotype 23 but not of haplotype 16 triggered an HR. Inspection of the cds of haplotype 16 revealed a G-to-T substitution in the first exon that causes a Gly (GGC)-to-Val (GTC) exchange in a conserved Gly residue of the predicted FAD-binding domain of the Bs3 protein (Table I; Supplemental Figs. S1A and S2). It is likely, therefore, that this difference renders haplotype 16 inactive. In addition to haplotype 16, we found 16 accessions (nine haplotypes [nos. 5, 8–13, 16, and 20]) that, due to polymorphisms in the cds, are unlikely to encode a functional Bs3 protein. One remarkable finding in this context is an 11-bp deletion in exon 1 that is found in 12 accessions (six haplotypes [nos. 8–13]). This deletion results in a frameshift and an early stop codon in the cds (Supplemental Figs. S1A and S2). Similarly, we identified in haplotype 20 a C-to-G substitution in exon 1 that changes a Tyr (TAC) to a stop codon (TAG). We expressed the cds of haplotype 20 and some representative haplotypes that are likely to encode nonfunctional Bs3 proteins under the control of the 35S promoter and confirmed via A. tumefaciens-mediated transient expression that these alleles are indeed incapable of triggering an HR. Altogether, our data show that many Capsicum accessions contain a nonfunctional Bs3 cds. None of these lines with nonfunctional Bs3 alleles showed any observable morphological phenotypes. DISCUSSION UPA Boxes Matching to Different TALes Show No Obvious Sequence Homology We wished to investigate how bacterial TALes specifically interact with and activate corresponding host plant promoters. To do so, we analyzed the Bs3 and Bs3-E promoters to gain insights into how they are specifically activated by the highly related transcription activators AvrBs3 and AvrBs3Δrep16, respectively. Using mutational analysis, we defined the UPA AvrBs3 box of the Bs3 promoter and the UPA AvrBs3Δrep16 box of the Bs3-E promoter (Fig. 7). As both boxes overlap by at least 11 bp, it is unclear if the conserved sequence motif that is present in the UPA AvrBs3 and UPA AvrBs3Δrep16 boxes may also be part of UPA boxes that are targeted by TALes distinct from AvrBs3 and AvrBs3Δrep16. Therefore, we inspected the promoters of the rice (Oryza sativa) genes Xa27, Os8N3, OsTFX1, and OsTFIIAγ1 that have been shown previously to be induced by the matching Xanthomonas oryzae pv oryzae TALes AvrXa27, PthXo1, PthXo6, and PthXo7, respectively (Gu et al., 2005; Chu et al., 2006; Yang et al., 2006; Sugio et al., 2007). None of these promoters contains a sequence that matches the 11-bp sequence present in the UPA AvrBs3 and UPA AvrBs3Δrep16 boxes. Thus, there is no evidence that UPA boxes matching different TALes share a consensus sequence. Our previous studies suggested that the hypervariable residues of TALes determine the promoter target sequence. This hypothesis is based on the analysis of AvrHah1, a TALe from Xanthomonas gardneri, which is recognized in pepper Bs3 plants and thus likely activates the Bs3 promoter. AvrHah1 shares blocks of high homology with AvrBs3 within the hypervariable repeat residues (Supplemental Fig. S3A; Schornack et al., 2008). Thus, comparative analysis of AvrHah1 and AvrBs3 suggested that TALes that share blocks of high homology within their hypervariable repeat residues are likely to target similar UPA boxes. Consistent with this idea is the fact that AvrBs3 and AvrBs3Δrep16 are identical in their first 10 repeat units (Supplemental Fig. S3A) and that the corresponding UPA AvrBs3 and UPA AvrBs3Δrep16 boxes are partially identical. Given that the hypervariable residues of AvrXa27, PthXo1, PthXo6, and PthXo7 share no homology to AvrBs3 within their hypervariable residues (Supplemental Fig. S3, B and C), it is also expected that matching UPA boxes would not share homology with the UPA AvrBs3 box. Taken together, these data indicate that no consensus UPA box exists. Given that not only the Bs3 but also the pepper UPA10 and UPA20 genes are induced by AvrBs3 (Kay et al., 2007), these promoters should contain a UPA AvrBs3 box. Indeed the UPA10, UPA20, and Bs3 promoters share sequence conservation, which is pronounced in the 5′ end of the corresponding UPA AvrBs3 boxes (positions −61 to −52; Supplemental Fig. S4). Nucleotides at position −46 to −44 of the Bs3 promoter showed less sequence similarity to the corresponding promoter regions of UPA10 and UPA20. In agreement with this observation, our substitution mutagenesis showed that sequence variation at positions −46 and −45 had no detectable effect and substitutions at position −44 had only a minor effect on the Bs3-mediated HR (Fig. 7; Supplemental Fig. S4). In this context, it is notable however, that the Bs3 promoter transposon mutant T−46, which basically lacks the last three nucleotides of the UPA AvrBs3 box (corresponding to positions −46 to −44; Fig. 4), did not trigger an HR. Mutant T−46 is equivalent to a “triple” substitution mutant in which positions −46, −45, and −44 are mutated. Notably, the insertion mutant T−46 showed no Bs3-mediated HR (Fig. 4), while all single nucleotide substitution mutants at positions −46 to −44 showed no or only partial reduction of the Bs3 HR (Fig. 7). We conclude that the suppression of the Bs3 HR in T−46 is due to the additive effect of three substitutions. This implies that single nucleotide substitutions at positions −46 and −45 must affect the inducibility of the Bs3 promoter, although these substitutions had no detectable effect on the Bs3 HR. Our mutational studies relied on a transient A. tumefaciens-mediated delivery system using constitutively expressed TALe genes in combination with Bs3 and Bs3-E promoter derivatives into N. benthamiana. This assay is useful for rapid analysis of in vitro-generated promoter derivatives. The assay also appears to be highly representative of the native interaction during infection, because naturally occurring Bs3 alleles by Xcv infection assay yielded identical results (see ECW, ECW-30R, PI593576, PI593491, PI631131, PI357635, PI593574, PI406948, PI631152, PI224415, PI599426, PI566809, PI181907, PI497971, CGN17230, CGN17227, PI593557, PI631137, CGN17042, and CGN17025 in Supplemental Fig. S1). Therefore, we are confident that our reporter system is an efficient and accurate assay to study interactions between TALes and corresponding host promoters. Sequence analysis of naturally occurring Bs3 alleles uncovered that haplotypes 16 and 23 carry mutations in the predicted UPA AvrBs3Δrep16 box (Supplemental Fig. S5). As anticipated, no HR was observed in either haplotype upon infection with avrBs3Δrep16-expressing Xcv strains as well as upon inoculation with an A. tumefaciens strain delivering a 35S promoter-driven avrBs3Δrep16 gene (Table I). Furthermore, when promoters of both haplotypes were fused to the Bs3 wild-type cds, they were unable to trigger an AvrBs3Δrep16-induced HR. Notably, haplotype 16 carries a C-to-G substitution at position −63 relative to the TSS (Supplemental Fig. S5) and thus resembles the C-to-G substitution mutant generated in this study (Fig. 7). By contrast, other haplotypes that contained promoter polymorphisms outside of the UPA AvrBs3Δrep16 box (e.g. haplotypes 19 and 22) were still capable of triggering an AvrBs3Δrep16-dependent HR (Table I). Thus, the definition of the UPA AvrBs3Δrep16 box by the aid of substitution mutants is supported by the functional analysis of naturally occurring Bs3 alleles. What Defines the TSS in a Gene with a UPA Box? We discovered that the UPA AvrBs3 box retained its function when moved to different locations within the pepper Bs3 promoter and also in the heterologous tomato Bs4 promoter (Fig. 10). We also determined that the placement of the UPA AvrBs3 box in different promoter locations changed the TSS. The distance between the TSS and the UPA AvrBs3 box was conserved in all constructs and was found to be between 41 and 46 bp with respect to the 3′ end of the UPA AvrBs3 box. Thus, the TSS of AvrBs3-inducible genes seems to be dictated by the location of the UPA AvrBs3 box. The initiation of mRNA synthesis in eukaryotic cells requires the assembly of general transcription factors and RNA polymerase II into a preinitiation complex at the core promoter. The only known sequence-specific DNA-binding protein among the general transcription factors is the TATA-binding protein, a subunit of the general transcription factor TFIID. Since AvrBs3 was shown to bind to the UPA AvrBs3 box-containing promoters (Kay et al., 2007; Römer et al., 2007) and since the TSS is dictated by the position of the UPA AvrBs3 box, it is possible that AvrBs3 replaces TATA-binding protein in its function as a sequence-specific DNA-binding protein in the preinitiation complex. Given that the UPA AvrBs3 box contains a TATA-like sequence motif, it is tempting to speculate that AvrBs3 binds to this promoter motif. However, mutational analysis of the UPA AvrBs3 box (Fig. 7B) did not provide any evidence that the TATA motif is functionally more important than other areas within the UPA AvrBs3 box. The observation that the UPA AvrBs3 box works at different promoter locations and within different promoter contexts suggests that diverse UPA boxes corresponding to different TALes may be arranged in tandem to make a complex promoter. If such a complex promoter is fused to the Bs3 cds, it should mediate recognition of multiple TALes. Such a promoter may confer durable resistance against a range of pathogens containing a range of TALes, as is the case of several Xanthomonas species, including X. oryzae pv oryzae, X. oryzae pv oryzicola, and X. axonopodis pv citri. How Did Bs3 Evolve and Does it Have a Function aside from Resistance? Previously, we showed that Bs3 and Bs3-E are transcriptionally induced by AvrBs3 and AvrBs3Δrep16, respectively, but not by the TALe AvrBs4 (Römer et al., 2007). Here, we show that Bs3 and Bs3-E transcripts are not detectable via RT-PCR in uninfected leaf, flower, fruit, or root tissue (Fig. 1B). This raises the question of whether the Bs3 protein has a biological function other than in recognition of TALes. Given that expression of Bs3 triggers cell death, a potentially detrimental function that requires strict control of gene expression, one wonders how this gene evolved. The sequence analysis of Bs3 alleles from different Capsicum accessions provides some clues to these questions. About one-third of the analyzed Capsicum accessions (16 of 49) contained a Bs3 cds with a predicted early stop codon (Table I; Supplemental Figs. S1A and S2). It is known that cell death plays an important role not only in disease resistance but also in developmental processes (Lam, 2004). Thus, we wondered if the accessions that carry a nonfunctional Bs3 cds show any phenotypes that would be indicative of a contribution of Bs3 to developmental or physiological processes. Yet, we observed no altered phenotypes in these accessions other than the change in their response to bacterial effectors, indicating that Bs3 is important only in the context of disease resistance. More than three-quarters of the analyzed Capsicum accessions (38 of 49) were identical to the Bs3-E haplotype within their promoter sequences. About three-quarters of these Bs3-E promoter-containing accessions (29 of 38) differed from the Bs3-E haplotype in their cds. By contrast, the three accessions that contain the Bs3 promoter were identical in their promoter and cds. The Bs3-E haplotype is more prevalent and possibly more ancient than the Bs3 haplotype. Thus, the Bs3 promoter is possibly the consequence of a 13-bp deletion in the Bs3-E promoter. Consistent with this hypothesis, haplotype 4 contains the Bs3-E promoter but is identical to the Bs3 haplotype in its cds (Supplemental Fig. S1A) and thus might represent the progenitor of the Bs3 haplotype. The predicted Bs3 protein is homologous to YUCCA-like proteins from Arabidopsis (Arabidopsis thaliana), some of which are involved in auxin biosynthesis (Schlaich, 2007; Chandler, 2009). Overexpression of YUCCA-like genes leads to phenotypes characteristic of auxin-overproducing mutants but not to an HR, as in the case of the Bs3 cds (Zhao et al., 2001; Kim et al., 2007; Römer et al., 2007). Hence, YUCCA-like proteins and Bs3 are functionally distinct. Yet, based on sequence homology, Bs3 can be considered as a YUCCA deletion derivative because it lacks a stretch of approximately 70 amino acids present in all predicted YUCCA proteins (Römer et al., 2007). We speculated that the Bs3 gene evolved from a YUCCA-like gene in pepper and anticipated that some Capsicum accessions would contain an ancestral Bs3 gene that encodes a “full-length” YUCCA-like protein instead of the YUCCA deletion derivative. However, none of the Capsicum haplotypes analyzed here contained a Bs3 cds that encodes a full-length YUCCA protein. Thus, the postulated deletion in the Bs3 cds most likely preceded the speciation of Capsicum. Given that Bs3 as a potential YUCCA deletion derivative triggers cell death, its expression must be strictly regulated. Therefore, it appears likely that a promoter mutation that made this promoter transcriptionally inactive preceded the deletion in the cds. In agreement with this idea, knockout mutations in Arabidopsis YUCCA-like genes produced no obvious phenotypes due to genetic redundancy (Schlaich, 2007). In summary, the analyses of Capsicum accessions suggest that the Bs3 gene has no function aside from disease resistance and provide some insights into how and when this potentially detrimental gene evolved. MATERIALS AND METHODS Generation of Promoter Deletion Constructs of Bs3 and Bs3-E Progressive 5′ promoter deletions of the Bs3 gene were PCR amplified from genomic DNA of pepper (Capsicum annuum ‘ECW-30R’; Minsavage et al., 1990). The PCR was carried out with Phusion high-fidelity DNA polymerase (New England Biolabs) and the primers Prom-90 bp-fwd-PR (5′-CACCAGTTATCATCCCCTTTCTCTTTTCTC-3′), Prom-179 bp-fwd-PR (5′-CACCGCACACCCTGGTTAAACAATGAACACG-3′), and Prom-356 bp-fwd-PR (5′-CACCTCATAGTCAAGCTAACGAAACTTATGC-3′) in combination with the primer Final-entry-02-rev (5′-CATTTGTTCTTTCCAAATTTTGGCAATATCTTGTGCAAC-3′). PCR fragments were cloned into pENTR-D (Invitrogen), sequenced, and transferred into the T-DNA vector pGWB1 (Nakagawa et al., 2007) via Gateway recombination (Invitrogen). pGWB1 derivatives were transformed into Agrobacterium tumefaciens GV3101 (Holsters et al., 1980) for transient expression assays. The Bs3-E alleles were cloned in the same way using genomic DNA from pepper cv ECW as template. Internal promoter deletions were generated by the Phusion site-directed mutagenesis kit (New England Biolabs). We used a Bs3 gene (promoter and cds) cloned in pENTR-D (Invitrogen) as template (Römer et al., 2007) to create deletions. Primers that were used are available upon request. All constructs were sequenced and transferred by Gateway LR recombination into pGWB1 (Nakagawa et al., 2007). pGWB1 derivatives were transformed into A. tumefaciens GV3101 for transient expression assays. Creation of Promoter Substitution and Insertion Mutants Substitution mutants in the Bs3 promoter were generated via site-directed mutagenesis using the Phusion site-directed mutagenesis kit (New England Biolabs). We used a Bs3 gene (promoter and cds) cloned in pENTR-D (Invitrogen) as template DNA (Römer et al., 2007). We employed primers that contain at a given position all nucleotides except the nucleotide present in the wild-type sequence. The different permutations were selected by sequence analysis of cloned fragments. The promoter constructs were transferred via Gateway LR recombination into pGWB4 (encodes a C-terminal GFP epitope tag; Nakagawa et al., 2007). pGWB4 derivatives were transformed into A. tumefaciens GV3101 for transient expression in planta. The same approach was used for creation of the Bs3-E promoter mutants, with the difference that we used a cloned Bs3-E gene as template DNA (Römer et al., 2007). For the insertion of nucleotides in the Bs3 promoter, we used the Phusion site-directed mutagenesis kit (New England Biolabs) and the primers site-dir-02-N-fwd-PR (5′-CTGACCAATTTTATTATATAAACCTNAACCATCCTCAC-3′), site-dir-02-NN-fwd-PR (5′-CTGACCAATTTTATTATATAAACCTNNAACCATCCTCAC-3′), site-dir-02-AA+N-fwd-PR (5′-CTGACCAATTTTATTATATAAACCTAANAACCATCCTCAC-3′), or site-dir-02-CT+N-fwd-PR (5′-CTGACCAATTTTATTATATAAACCTCTNAACCATCCTCAC-3′) in combination with the primer site-dir-02-rev-PR (5′-GCAAACGTGTTCATTGTTTAACCAGGGTG-3′). All primers used are phosphorylated at their 5′ termini. We used a Bs3 gene (promoter and cds) cloned in the vector pENTR-D (Invitrogen) as template DNA (Römer et al., 2007). After sequence analysis, cloned fragments were transferred into pGWB1 by Gateway LR recombination (Nakagawa et al., 2007). For insertion of a 13-bp sequence at different locations within the Bs3 promoter, we used the Phusion site-directed mutagenesis kit (New England Biolabs) in combination with primers Prom-Bs3+13-20nU-fwd-PR (5′-CTCTATTCCACTACCTTTCTCTTTTCTCCTCTTG-3′) + Prom-Bs3+13-20nU-rev-PR (5′-GGATGATAACTTGAAGTTGTGGGATG-3′) or primers Prom-Bs3+13+31UTR-fwd-PR (5′-CTCTATTCCACTACAAGTAGTCCTAGTTGCACAT-3′) + Prom-Bs3+13+31UTR-rev-PR (5′-TGTTTTGATAGATTTAGCGGGTGACAAG-3′). A Bs3 gene (promoter and cds) cloned in pENTR-D (Invitrogen) was used as template (Römer et al., 2007). For the insertion of a UPA AvrBs3 box, we also used the Phusion site-directed mutagenesis kit. As template, we used pENTR-D, which contains a Bs3 gene with a mutation (−61T is replaced by A) in the original UPA AvrBs3 box. For the insertion of the Bs3 UPA293 box, we used primers box-02-293-fwd-PR (5′-CAATTTTATTATATAAACCTAACCATCCTCACAACCAAGTAAACTCAAAGAACTAATCATTGAAC-3′) and box-02-293-rev-PR (5′-CATACTAATTTCATATTTCCCTTGCATAAG-3′). To insert the Bs3 UPA424 box, we used primers box-03-424-fwd-PR (5′-CAATTTTATTATATAAACCTAACCATCCTCACAACCACATTAGATTGTACTTGCTTTTTACCACAGATAC-3′) and box-03-424-rev-PR (5′-TCATGTATCATTCGCATTTCAAAGTAAAACTAAGG-3′). For the generation of Bs4 UPA and Bs4 UPAmut constructs, we used a Bs4 promoter fragment of 302 bp (Schornack et al., 2005) in pENTR-D (Invitrogen). In the Bs4 UPAmut, all C nucleotides in the UPA AvrBs3 box are replaced by G nucleotides. The boxes were inserted via Phusion site-directed mutagenesis using primers Bs3inBs4-promfwd-PR (5′-CAATTTTATTATATAAACCTAACCATCCTCACAACGTTTCAAGTGGTACTTGT-3′) and Bs3ubm1inBs4-promfwd-PR (5′-CAATTTTATTATATAAAGGTAAGGATCCTCACAACGTTTCAAGTGGTACTTGT-3′) in combination with the primer Bs3in4-promrev-PR (5′-GTGAAAGCTTGTATTAACATTCGCTTTG-3′). After sequence analysis, the promoter constructs were transferred by Gateway LR recombination into the T-DNA vector pK7-GW-Bs3. pK7-GW-Bs3 was generated on the basis of pK7FWG2 (Karimi et al., 2002). We removed the HindIII and BamHI fragments by restriction digest from pK7FWG2 (contains the cauliflower mosaic virus 35S promoter, the Gateway attR cassette, and the GFP cds) and replaced it by a synthesized DNA fragment that contains SacI and EcoRV restriction sites followed by the cauliflower mosaic virus 35S terminator. Next, a Gateway attR cassette was placed into the EcoRV site, resulting in pK7-GW. The Bs3 cds was amplified from genomic DNA of pepper cv ECW-30R using the primers final-entry-SacI-01-fwd-PR (5′-GGGGGGAGCTCATGATGAATCAGAATTGCTTTAATTCTTGTTC-3′) and final-entry-SacI-02-rev-PR (5′-GGGGGGAGCTCCATTTGTTCTTTCCAAATTTTGGCAATATC-3′). These primers add a SacI restriction site on both ends of the Bs3 cds. The Bs3 cds was cloned into the SacI restriction site of pK7-GW, resulting in pK7-GW-Bs3. Linker-Scanning Mutagenesis of the Bs3 Promoter For insertion mutagenesis, we used the GPS-LS Linker Scanning System (New England Biolabs). As template, we used pENTR-D containing 343 bp 5′ of the ATG start codon of the Bs3 gene fused to the Bs3 cDNA. This plasmid was created by splicing using overlap extension PCR. The promoter was amplified from genomic DNA of ECW-30R pepper plants using primers Prom-356 bp-fwd-PR (5′-CACCTCATAGTCAAGCTAACGAAACTTATGC-3′) and B5-rev-PR (5′-CATACGGAACACTGTATTGCTTAAGG-3′). For cDNA synthesis, pepper ECW-30R plants were inoculated with a blunt syringe using Xanthomonas campestris pv vesicatoria strain 85-10 expressing avrBs3 (pDS300F; optical density at 600 nm [OD600] = 0.4). RNA extraction and cDNA synthesis were done as described previously (Römer et al., 2007). The Bs3 cDNA was amplified with Phusion high-fidelity DNA polymerase and the primers Final-entry-01-fwd-PR (5′-ATGATGAATCAGAATTGCTTTAATTCTTGTTC-3′) and Final-entry-01-rev-PR (5′-CTACATTTGTTCTTTCCAAATTTTGGCAATATCTTGTGC-3′). PCR products of the cDNA and the promoter region were mixed in a 1:1 ratio and PCR amplified using Prom-356 bp-fwd-PR and Final-entry-01-rev-PR primers. The PCR product was cloned into pENTR-D (Invitrogen), sequenced, and used for the transposon mutagenesis. Transposon insertions in the promoter region were identified via PCR. The identified transposon mutants were treated according to the manual, so that only the 15-bp insertion of the transposon was left. The resulting transposon mutants were sequenced and then recombined into pGWB1. Plant Material and Infection Assays Pepper and Nicotiana benthamiana plants were grown as described previously (Römer et al., 2007). Pepper germplasm was supplied by the U.S. Department of Agriculture (accessions preceded by “PI” or “Grif”) and the Plant Genetic Resources cluster of the Centre for Genetic Resources, The Netherlands (accessions preceded by CGN). Information on corresponding pepper accessions is available at http://www.ars-grin.gov/npgs/acc/acc_queries.html and http://www.cgn.wur.nl/UK/CGN+Plant+Genetic+Resources/. Agrobacterium-mediated transient transformation of N. benthamiana leaves, Xcv infection assays of Capsicum species, RT-PCR, RACE, and EMSA were carried out as described previously (Römer et al., 2007). Generally, Xanthomonas and Agrobacterium infection assays were routinely carried out at three independent time points. At each time point, each bacterial strain or combination of strains was inoculated into four different leaves. Agrobacterium strains delivering T-DNAs that encode bacterial effector genes were generally used in multiple infiltrations, always including an appropriate positive and negative control on each inoculated leaf. Supplemental Data The following materials are available in the online version of this article. Supplemental Figure S1. Nucleotide polymorphisms of Bs3 alleles. Supplemental Figure S2. Proteins that are encoded by naturally occurring Bs3 alleles. Supplemental Figure S3. Sequence comparison of TALe hypervariable residues. Supplemental Figure S4. Comparison of UPA AvrBs3 boxes of different AvrBs3-inducible promoters. Supplemental Figure S5. Alignment of UPA AvrBs3Δrep16 box variants in naturally occurring Bs3 alleles. Supplemental Data Set S1. Bs3 promoter deletions. Supplemental Data Set S2. Nucleotide sequences of linker-scanning Bs3 promoter mutants. Supplemental Data Set S3. Nucleotide sequences of naturally occurring Bs3 alleles in Fasta format. Supplemental Data Set S4. Sequence alignment of naturally occurring Bs3 alleles. ACKNOWLEDGMENTS We are grateful to Diana Horvath, Annett Strauss, and Christoph Peterhänsel for helpful comments on earlier versions of the manuscript. We acknowledge the technical support of Sabine Recht, Jens Hausner, Carola Kretschmer, Bianca Rosinsky, and Marina Schulze. 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The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Thomas Lahaye ([email protected]). [W] The online version of this article contains Web-only data. www.plantphysiol.org/cgi/doi/10.1104/pp.109.139931 © 2009 American Society of Plant Biologists This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)