The molecular genetics of virulence of Xanthomonas campestris

Biotechnology Advances 17 (1999) 489–508

Research review paper

The molecular genetics of virulence of Xanthomonas campestris James W.Y.F. Chan, Paul H. Goodwin* Department of Environmental Biology, University of Guelph, Guelph, Ont. N1G 2W1, Canada

Abstract Bacteria belonging to the genus Xanthomonas are important pathogens of many plants, and their virulence appears to be due primarily to secreted and surface compounds that could increase host nutrient loss, or avoid or suppress unfavorable conditions in the host. Type II and III secretory pathways are essential for virulence. Some individual extracellular enzymes (type II-secretion dependent) affect final bacterial population levels, whereas some avirulence gene products (type III-secretion dependent) affect virulence by altering host metabolism. Avr proteins, probably secreted via a pilus, can also be recognized by host resistance gene products. Virulence is also associated with bacterial surface polysaccharides, which may help to avoid host defense responses, and regulatory gene systems, which can control virulence gene expression. Keywords: Plant disease; Bacteria; Secretion; Regulation; Extracellular polysaccharide; Avirulence

1. Introduction The Xanthomonas genus is arguably one of the most ubiquitous group of plant-associated bacteria. Members of this genus have been shown to infect at least 124 monocotyledonous and 268 dicotyledonous plants, while other members of the genus are saprophytic and epiphytic. Within the Xanthomonas genus, Xanthomonas campestris is the most dominant species with at least 141 pathovars [1] identified by classic taxonomical methods [2]. The taxonomy of the Xanthomonas genus has traditionally been determined by a phenetic approach based mainly on biochemical, physiological, morphological and phytopathogenic features. This approach in interpreting phenotypic features was developed to satisfy a practical need for plant pathologists to name pathogens that are specific for particular plant hosts or certain diseases. As a result, a nomenclature scheme for pathovars and species was developed based almost solely on a single phenotypic feature, namely phytopathogenicity [2]. Although * Corresponding author. Tel.: 11-519-824-4120; fax: 11-519-837-0442 E-mail address: [email protected] (P.H. Goodwin) 0734-9750/99/$–see front matter © 1999 Elsevier Science Inc. All rights reserved. PII: S0 7 3 4 - 9 7 5 0 ( 9 9 ) 0 0 0 2 5 -7

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this highly artificial scheme was deemed adequate for plant disease control purposes, modern genetic and molecular genetic techniques and research in the study of molecular plant— microbe interactions have outstripped the usefulness of this traditional classification scheme. Vauterin et al. [3] proposed a comprehensive revision of the classification of the Xanthomonas genus. Under the new nomenclature, the genus comprises 20 DNA homology groups. The most heterogeneous group, Xanthomonas campestris, was divided into 16 DNA homology groups. Group 9, known as X. axonopodis, is the largest and most heterogeneous group and contains most of the X. campestris pathovars. Correlating the genomic groups with their pathogenic specialization is variable. For crucifers, grasses and cereals, pathovars attacking related hosts are clearly within the same genomic groups. Conversely, pathovars attacking different hosts could be found in the same genomic group [3]. It appears that pathogenic specialization does not often correlate well with the genetic relatedness of xanthomonads, and there is a continuous range of biodiversity in this genus, which makes any classification system somewhat artificial [4]. The phytopathogenicity of xanthomonads is a complex phenomenon. Based on mutation data, Daniels [5] estimated that between 20 and 100 genes are involved in phytopathogenicity. Pathogenesis involves many steps, beginning with penetration of the plant through wounds and natural openings. Once inside the plant, Xanthomonas cells multiply in the intercellular spaces until the spaces become filled with bacteria and bacterial extracellular polysaccharide [6]. This is associated with the appearance of water-soaking and increased plant cell permeability, which would increase the loss of nutrients from the plant cell. For some Xanthomonas species and pathovars, the bacteria will invade the vascular tissue, where they multiply and spread through the plant. Later, during foliar or vascular infections, plant cells adjacent to bacterial colonies begin to degrade. The plant organelles degenerate, cell walls swell and fragment, and finally bacterial cells enter and multiply inside the plant cells. In susceptible host plants, disease symptoms appear after several days of bacterial growth. Symptoms caused by Xanthomonas pathogens can include chlorosis, necrosis, wilting, hypertrophy, rotting, die back and cankers [6]. These pathogens have been described as being both biotrophic (i.e., feeding on living host tissue) because they multiply considerably before any damage is visible, and necrogenic (i.e. killing plant cells) because they cause necrosis [7]. However, a more accurate description would be that most xanthomonads are hemibiotrophic; bacteria initially feed on living host cells but then disrupt and kill host cells and use the nutrients in the dead cells.

2. Extracellular enzymes, phytotoxins and polysaccharides—production and virulence Extracellular enzymes may play a role by facilitating plant tissue maceration, and have long been considered important virulence determinants. Strong evidence for the role of extracellular enzymes in the virulence of Xanthomonas campestris pv. campestris, the causal agent of black rot of crucifers, has come from mutations in the secretion pathway genes, which causes both a retention of certain extracellular enzymes in the cells and a loss of virulence [8,9]. These results prompted investigations of a number of genes that encode individual extracellular enzymes to examine the separate roles of these genes in virulence.

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Two major endoglucanase genes were obtained from X. campestris pv. campestris by Gough et al. [10,11]. Marker exchange of one of the endoglucanases with a Tn5 insertion generated an endoglucanase-minus X. campestris pv. campestris mutant. This major endoglucanase structural gene encodes a 53-kD extracellular protein. Strong sequence homology has been identified with an enzyme from Cellulomonas fimi, and a domain that is conserved in cellulase family A was clearly identified. The other major endoglucanase structural gene encodes a cytoplasmic enzyme. Little periplasmic endoglucanase activity was found in X. campestris pv. campestris. While the endoglucanase-minus mutant generated no detectable endoglucanase activity, it nonetheless retained phytopathogenicity to seedlings, which indicated that endoglucanase alone is not important in the disease. Pectate lyase (polygalacturonate lyase) genes have been cloned by X. campestris pv. campestris, X. campestris pv. vesicatoria and X. campestris pv. malvacearum [12–14]. There are three isoforms of pectate lyase in X. campestris pv. campestris. Although the synergistic effects of these enzymes are unknown, pectate lyase isozyme I is the least active of the three isozymes. Marker-exchange Tn5 mutants lacking pectate lyase isozyme I retained the same pathogenicity as the wild type in inoculated seedlings [13]. X. campestris pv. campestris also encodes a pectate transeliminase that exhibits the same molecular weight as pectate lyase isozyme I from this bacterium [8]. From X. campestris pv. vesicatoria, a 1.4 kb PstI restriction fragment has been demonstrated to encode for a pectate lyase gene which has no homology to DNA or Erwinia chrysanthemi, a soft-rot bacterium [12]. There is only a single isoform of pectate lyase in X. campestris pv. vesicatoria. In addition, it has been demonstrated that only a subpopulation of X. campestris pv. vesicatoria exhibited pectolytic activity on sodium polypectate medium. Nevertheless, the 1.4-kb pectate lyase gene is able to hybridize to DNA from all xanthomonads regardless of their ability to break down pectic compounds. Both pectolytic and nonpectolytic X. campestris pv. vesicatoria isolates are able to cause bacterial spot disease in tomatoes or peppers. The causal agent of cotton blight, X. campestris pv. malvacearum also contained one isoform of pectate lyase, which is reported to crosshybridize to DNA from a number of pathovars [14]. It was demonstrated that X. campestris pv. malvacearum is capable of producing high levels of pectate lyase in vitro, but highlyproducing strains did not show significantly increased disease severity in inoculated plants. Thus, it appears that the pectate lyase serves very little role, if any, in virulence, but may still play a role related to bacterial nutrition. A gene encoding an extracellular protease has also been cloned from X. campestris pv. campestris and two major extracellular proteases (denoted PRT1 and PRT2) have been identified [15–17]. The two major extracellular proteases belonged to different protease classes and showed different patterns of peptide bond cleavage. The PRT1 protease is similar to members of the subtilisin family of serine proteases, which require a divalent metal ion for stability and activity. The PRT2 protease is similar to the neutral protease from Aeromonas proteolytica and the two extracellular metalloproteases from Erwinia chrysanthemi, which require Zn12 for activity and Ca12 for stability. Whether these are expressed in planta remains uncertain. A mutant lacking PRT1 and PRT2 proteases produced slightly attenuated symptoms when introduced into mature plant leaves. Similar results have been shown with protease-deficient mutants of X. campestris pv. oryzae, X. campestris pv. malvacearum and X. campestris pv. glycines [15,18]. Thus, proteases in Xanthomonas campestris may serve

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nutritional roles rather than serve as virulence determinants. Conversely, one can argue that the proteolysis of plant cell walls during the later stages of disease development is critical for overcoming plant host defense mechanisms. Therefore, the role of proteases in Xanthomonas virulence remains unclear. An a-amylase gene has been isolated from a X. campestris pv. campestris strain and expressed in E. coli [19]. The amino acid sequence of the 45-kDa protein resembled the a-amylase of Aeromonas hydrophila N-terminal amino acid sequence analysis confirmed that the enzyme is exported. However, since marker-exchange and in planta experiments were not conducted, the role of the extracellular a-amylase in the virulence of X. campestris has yet to be established. Although it appears that extracellular enzymes play a small role in virulence in plant seedling assays, they do affect bacterial growth and symptoms in mature plants. Mutants of X. campestris pv. campestris in protease, an endoglucanase enzyme, export genes and a regulatory gene for enzyme synthesis and all showed an approximate 10-fold decrease in population along with some reduction in symptoms. The only exception to this was a mutant defective in one of the two endoglucanase genes [20]. A number of phytotoxins have been proposed to be involved in the development of symptoms due to infection by xanthomonads [6]. However, there is as yet a lack of molecular genetic evidence to support a role for phytotoxins. An exception is the albicidin phytotoxins produced by X. albilineans, a pathogen of sugarcane. Transformation of X. albilineans with a plant esterase that detoxifies albicidins resulted in strains that did not release albicidins or cause disease [21]. A characteristic of the Xanthomonas genus is the appearance of mucoid colonies when cultured on media supplemented with glucose. The slimy appearance is the result of copious amounts of extracellular polysaccharide (EPS), also known as xanthan gum, produced by the cells. The unique and unusual rheological properties of xanthan gum in solution has given rise to many industrial applications [22–24], and has also made it one of the most extensively studied microbial polysaccharides [25]. Xanthan gum is a pentasaccharide repeating unit composed of a cellulosic backbone to which trisaccharide side chains are attached at the C-3 position on alternating D-glucosyl residues [26]. Depending on the growth conditions, the trisaccharide side chains contain varying molar rations of D-mannose and D-glucuronic acid may be acetylated or pyruvylated, respectively. Two genetic loci responsible for EPS production and regulation have been independently isolated from X. campestris pv. campestris [27,28]. These genes are clustered, and some mutations in these loci have pleiotropic effects because EPS and lipopolysaccharides (LPS) are partially composed of similar sugar subunits [27,29–31]. One gene cluster for EPS biosynthesis, known as the gum region, is composed of 12 genes and is mainly expressed as a single operon [32]. The gum gene products are responsible for assembly and polymerization of the sugar subunits and then secretion of the polymer [33]. There is evidence that Xanthomonas EPS can be an important virulence determinant. Although it was initially reported that mutations affecting EPS production in X. campestris pv. campestris did not significantly reduce its virulence to plant seedlings [29], a subsequent study using 4- to 5-week-old plants showed that the same EPS mutant had very limited bacterial growth and produced almost no symptoms [20]. It was suggested that the EPS can mask the bacteria to prevent recognition and plant defense responses. Mutation of X. campestris pv. campestris in the gumD gene, which

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encodes a glucosyltransferase involved in synthesis of the first xanthan lipid intermediate, also resulted in reduced virulence, but the mutation appeared to have pleiotropic effects as both EPS and xanthomonadin pigment production were altered [34]. However, Katzen et al. [33] found that mutation of gumD caused a 50% reduction in virulence and a complete loss of EPS synthesis without affecting pigment formation. Mutation of gumF, gumG and gumI which encode two acetyltransferases and a glycosytransferase, respectively, also reduced virulence indicating that shortening the sugar side chains and altering the acetylation of the sugars affects virulence [33]. Acetylation has been shown to be directly related to both EPS viscosity and virulence in X. campestris pv. campestris [35]. In contrast, mutation of gumL, encoding a pyruvate transferase, did not affect virulence or alter the amount of EPS produced. Mutation of several of the other gum genes appeared to be lethal, perhaps due to the accumulation of toxic lipid-linked xanthan intermediates [33]. In X. oryzae pv. oryzae, evidence suggests that EPS production is controlled by the rpfC gene, which is part of a two-component signal transducing system [36]. When the rpfC gene was mutated, both EPS production and virulence to rice were greatly reduced, even though bacterial growth rates and maximum population levels were only slightly reduced [36]. Interestingly, when the rpfC homolog was mutated in X. campestris pv. campestris virulence was also attenuated [37]. The biological role of EPS can probably be better defined by its three important properties: it is highly hydrated so that it provides protection against desiccation and hydrophobic molecules; it is highly anionic, which allows it to concentrate nutrients and immobilize toxic elements; and it has adhesion quality that allows an organism to adsorb to biological surfaces [38]. Therefore, Xanthomonas EPS is likely a major determinant in the ability of the genus to colonize a diverse number of habitats in addition to plant host tissue. There is some evidence for lipopolysaccharide (LPS) involvement in the suppression of hypersensitive response (HR) in plant hosts [39]. Although the LPS-related response is unknown, it is well established that pre-inoculation of plant leaves with X. campestris pv. campestris LPS core-oligosaccharide at least 10 h before inoculation of an avirulent strain could prevent HR symptoms. The opsX gene required for the assembly of LPS has been cloned from X. campestris pv. citrumelo [40]. Mutation of opsX resulted in a loss of virulence on the citrus host marked by rapid cell death within the plant host, although HR, host range and virulence to beans remained unaffected. While it is tempting to conclude that LPS is responsible for the protection from plant inhibitors, mutations in opsx are also pleiotropically affected in growth rate, capsular slime and EPS. Nevertheless, defects in LPS could be partially responsible for the protection from plant inhibitors [41]. In X. campestris pv. campestris, mutation in rfaX, another gene involved in LPS assembly, increased the pathogen’s sensitivity to antibiotics as well as reducing bacterial growth in both host and nonhost plants. Recent evidence strongly suggests that LPS is likely to be involved in the association of bacteria with plant cell walls during the infection process [42]. 3. Extracellular enzymes and polysaccharides regulation Virulence in Xanthomonas is coordinatedly regulated like that of other pathogens responsible for diseases in animals and plants. The regulation of enzymes and polysaccharide ex-

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pression in Xanthomonas can be divided into four components: sensory transduction, positive and negative regulation, export, and global regulation. Bacteria have evolved sensory transduction systems to detect environmental changes and to relay the information in the form of intracellular signals. The two-component signal transducing systems in bacteria are usually composed of a kinase sensor and a response regulator, although hybrid kinases serving both functions, also exist [43,44]. At least three two-component signal transduction systems have been discovered in X. campestris with two serving a possible role in the regulation of extracellular polysaccharide and enzyme production [37, 45,46]. Thus, it is conceivable that the cascades of events leading to extracellular polysaccharide and enzyme production could be initiated by a number of two-component signal transducing systems. The two-component signal transducing system encoded by rpfC and rpfG (regulation of pathogenicity factors) has been proposed to be an element in the positive regulation of extracellular enzymes and polysaccharide production [37,47]. Sequence data for rpfC and rpfG revealed that these genes both encoded hybrid kinases. The purpose of having redundant roles within a gene cluster remains unclear. These genes are responsible for the regulation of the rpf gene cluster, which mediates the secretion of enzymes and polysaccharide. The full biochemical roles of the gene products from the rpf gene cluster, containing at least eight genes, have not been elucidated. However, the cluster does not encode structural genes responsible for the synthesis of extracellular enzymes and polysaccharide. Mutations in any of the rpf genes caused a reduction in extracellular enzymes and polysaccharide to less than 10% of the wild-type levels and also a reduction in virulence [37,48]. The rpf gene cluster is not unique to X. campestris pv. campestris since the genes could cross-hybridize to DNA of other pathovars and similar gene clusters from X. campestris pv. translucens have been cloned [49]. The rpfB and rpfF genes may code for a diffusible factor that belongs to a class of ‘autoinducers’ in prokaryotes [50]. The predicted enzyme activities of rpfB and rpfF suggests that they are involved in the production of a fatty acid derivative that acts as the diffusible factor. A possible function of the diffusible factor may be as a cell density ‘sensor’ so that under a high bacterial cell density inside a plant host, bacterial cells coming in contact with the autoinducer will increase enzyme production by approximately tenfold. This evolutionary strategy provides the pathogen with a method to ‘fine tune’ its enzyme production when it is most needed, such as when a high bacterial cell density has been reached and plant nutrients are beginning to be depleted [50]. In this scenario, when the bacteria become starved, extracellular enzymes would be produced to degrade plant polymers and create more substrates for further bacterial growth. Mutation of rpfb and rpfF results in a loss of virulence [50]. Another similar but separate diffusible factor is encoded by the pigB gene of X. campestris pv. campestris, which regulates both EPS and xanthomonadin pigment production [51,52]. The pigB gene is not important for virulence if the bacteria are infiltrated into host tissue, but it is important for epiphytic growth and fewer lesions developed from infections that resulted from epiphytic populations [53]. The rpfN gene is responsible for the negative regulation of enzyme and polysaccharide production [54]. When introduced into a wild-type X. campestris pv. campestris, the overexpression of rpfN gene caused repression of enzyme and polysaccharide production. Con-

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versely, a mutation in the rpfN gene resulted in the overproduction of enzymes and polysaccharide. The biochemical role of RpfN remains unknown, although it is thought to interact with a protein that binds to the sequences upstream of the promoter of protease and endoglucanase genes. Evidence for a highly regulated virulence system is also apparent from an experiment using a promoter-probe system [55]. Host plant-inducible genes in X. campestris were identified using a shotgun cloning method, by ligating X. campestris pv. campestris chromosomal DNA to a broad host range plasmid containing a promoterless chloramphenicol acetyltransferase gene (CAT). The constructs were subsequently conjugated into X. campestris pv. campestris and virulence was assayed in chloramphenicol-treated turnip seedlings. Four chloramphenicol-resistant transconjugants were found to be virulent in the seedlings. It remains to be seen how these plant host-inducible genes interact with proteins from the sensory transduction, positive and negative regulation pathways. A gene designated as clp (catabolite activation protein-like protein) has been found to play a significant role in the regulation of virulence [56]. There is a 45% DNA homology between the clp gene and the gene for CAP from E. coli, which is responsible for the global regulation (i.e. catabolite repression) of operons scattered throughout the chromosome in the presence of cAMP. However, the Clp enzyme activity of X. campestris pv. campestris was not influenced by varying concentrations of cAMP. CAP mutations in E. coli are impaired in the utilization of a number of carbon sources, while Clp mutations in X. campestris pv. campestris are not. However Clp xanthomonads are pleiotropically affected in extracellular enzyme, pigment and polysaccharide production, and they demonstrate a reduction in virulence. Whether the clp gene product directly or indirectly regulates enzyme and polysaccharide production remains unknown. However Clp has similar DNA-binding specificity to E. coli CAP, which points to the possibility of Clp playing a role in the protein–protein interaction with RNA polymerase in the transcriptional activation of X. campestris promoters [57]. Although the regulatory elements described above have been proposed to be responsible for genetic regulation of both enzymes and polysaccharide production, polysaccharide regulation could be substantially more complicated. The biosynthesis of xanthan and lipopolysaccharide requires precursors of carbohydrates and acyl groups [25,58,59]. Thus, the regulation of the polysaccharide synthesis pathway would be influenced by the formation of precursors that are under different and separate control. In addition, the export mechanisms of xanthan, unlike extracellular enzymes, have not been well elucidated. The secretion system for extracellular enzymes is expressed by a cluster of genes in Xanthomonas [8,9,60,61]. The deduced amino acid sequences of some of the Xanthomonas protein secretion (xps) gene products showed close relatedness to pul gene products from the Klebsiella spp. XpsE shares similarity to a number of nucleoside triphosphate-binding proteins and is largely hydrophilic [9]. This suggests that XpsE may be involved in the mediation of an energy tranduction event during the translocation of the protein across the membrane. Hydropathy profiles of XpsF suggest that it is a transmembrane protein with cytoplasmic and periplasmic domains, and may be involved as part of a membrane pore through which proteins are translocated. XpsD, XpsG, XpsH, XpsI and XpsJ are thought to be parts of this membrane pore as well, and XpsO is a type IV pre-pilin leader peptidase required for protein secretion and the processing of type IV pilin [9,62]. Although the mode of

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regulation of the xps gene cluster is unknown, it is possible that most of the gene products are transcribed as a polycistronic message as in the Klebsiella pul cluster [63]. In fact, alignment of similar secretory pathways from other plant pathogens indicates that there is high homology between the different protein export genes and the Xps proteins of the Xanthomonas type II secretion pathway.

4. Compatible and incompatible plant–bacterial interactions The interactions between a plant and pathogenic bacterium can be classified as either compatible or incompatible. In a compatible interaction, a bacterium is able to overcome plant host defenses, and ultimately manifest disease symptoms through the elaboration of toxins, extracellular enzymes and/or other factors. In contrast, an incompatible interaction between a bacterium and a nonhost or resistant plant results in the growth of the bacterium being severely attenuated. Rapid localized plant-cell death at the site of infection is often the hallmark of an incompatible interaction. This incompatible response, also known as hypersensitive response (HR), is due to a wide array of plant defense responses targeted against the invading bacterium resulting in localized necrosis of the plant cells [64]. The bacterial genes responsible for compatible and incompatible interactions between plant and bacteria are known as hrp (hypersensitivity response and pathogenicity) genes and avr (avirulence) genes. The former group of genes determine the outcome of the plant–bacterial interactions, whereas the latter group determine the specificity of the interaction. Plants have evolved defense mechanisms that are due to distinct resistance genes, resulting in the gene-for-gene interaction with a pathogen [65]. In the absence of either bacterial avr genes or plant resistance genes, the outcome is disease. Resistance occur when plant resistance gene products are able to detect bacterial avr gene products. In contrast, another form of resistance exists, which is termed nonhost resistance or general resistance, and is due to other mechanisms without the requirement of specific plant resistance genes. The cloning of the first plant disease resistance gene was reported in 1992, and since then many different resistance genes, which can be categorized into different classes based on the predicted features of the gene products, have been cloned [66]. For Xanthomonas, two resistance genes have been reported. The Xa1 gene from rice is specific to race 1 of X. oryzae pv. oryzae and is classified with resistance genes encoding cytoplasmic proteins with a leucinerich repeat domain and a nucleotide binding site [67]. In contrast, the Xa21 gene provides resistance to over 30 races of X. oryzae pv. oryzae and encodes a transmembrane protein with an extracellular leucine-rich repeat and a cytoplasmic protein kinase domain [68]. Based on just these two examples, it appears likely that resistance genes to xanthomonads will have a diversity of structures and belong to a variety of resistance gene classes.

5. Hypersensitivity response and pathogenicity hrp genes The first cluster of genes shown to play a role in both HR and pathogenicity was discovered in Pseudomonas syringae, and is now commonly referred to as hrp genes [69,70]. Mutation of the hrp genes would result in loss of pathogenicity in host (or compatible) interac-

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tion and loss of HR in nonhost (or incompatible) interaction. These findings are significant in that they point to a regulatory system that exists at a higher level than the ones previously described for extracellular enzymes and polysaccharide production. To date, most of the discoveries in the hrp system have come from the genus Pseudomonas. The first published report of the hrp genes in Xanthomonas was made by Stall and Minsavage [71] for X. campestris pv. vesicatoria. It was argued that the presence of a hrp gene cluster in a xanthomonad strain provided strong evidence that the bacterium is capable of growing in the plant host. Efforts to detect the hrp genes in saprophytic xanthomonads through Southern hybridization proved that the hypothesis is true, and thus, the hrp genes do appear to play a critical role in determining the outcome of a plant–bacterial interaction [71]. However, weak hybridization signals were obtained under low-stringency washing conditions for almost half of the saprophytic xanthomonads. This suggest that the progenitor of the hrp genes may have served other roles and may exist for functions other than pathogenicity. Using hrp-conserved primers and the polymerase chain reaction, Leite et. al. [72] showed that hrp gene clusters are highly conserved in many X. campestris pathovars. Saprophytic xanthomonads did not yield any amplification products. However, some phytopathogenic xanthomonads, such as X. campestris pv. translucens, did not produce any amplified products either. There are at least two other weakly-homologous hrp gene clusters within the X. campestris pathovars [73,74]. Hrp clusters can be divided into two groups based on the similarity of the genes, operon structure and regulation, and for X. campestris the hrp cluster belongs to group 2 [7]. The 23-kb hrp cluster in X. campestris pv. vesicatoria is by far the best characterized among the xanthomonad hrp genes [73,75]. The hrp cluster is located on the chromosome, with six loci, designated hrpA to hrpF, identified by Tn5-mutagenesis and marker gene exchange experiments. The X. campestris pv. vesicatoria hrp cluster could complement the loss of HR and pathogenicity in a mutagenized X. campestris pv. phaseoli. The gene cluster could also be Tn5 mutagenized and transferred by marker exchange into a number of X. campestris pathovars resulting in a loss of HR and pathogenicity in their respective plant hosts and nonhosts. The entire sequence of the hrp cluster has been determined [75]. Similarities between several of the predicted Hrp proteins and secretion proteins from Yersinia, Shigella flexneri, Bacillus subtilis, Salmonella typhimurium and Caulobacter crescentus led researchers to postulate that the X. campestris pv. vesicatoria Hrp proteins are part of a type III secretion apparatus required for the export of virulence factors and elicitors of plant defenses, such as the avr gene products. The hrpA gene encodes a 64 kDa protein with an N-terminus that resembles a signal peptide sequence. Based on the presence of a hydrophobic core, the HrpA1 gene product is likely to be a transmembrane protein. The hrpB operon encodes 8 proteins [76]. HrpB3 is a 27.3-kDa protein with two possible hydrophobic membrane-spanning domains and a lipoprotein signal sequence cleavage site for signal peptidase II. The HrpB3 gene product is also likely to be localized in the membrane. In contrast, HrpB6 is a hydrophilic protein with a molecular weight of 47.7 kDa. The hrpC operon encodes at least three genes, one of which is a 69.9-kDa protein with six possible transmembrane domains, and is predicted to be an innermembrane protein. The hrpD locus encodes three gene products designated as HrpDI to HrpD3 that also conservation with components of the type III secretion system [77]. The

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hrpF locus encodes a 86.4-kDa protein that is predominantly hydrophilic, but has two hydrophobic domains. Sequence homology exists between HrpF and NolX from Rhizobium fredii NolX is involved in host specificity, and mutation in this protein would lead to host-range extension in Rhizobium. It is uncertain how this function could be translated into the pathogenicity of Xanthomonas but it is interesting that NolX belongs to cluster of genes that have shown some homology to the X. campestris pv. vesicatoria hrp genes. It is tempting to speculate that HrpF is also involved in protein translocation. The regulation of the 23 kb hrp region is under the control of hrpXv gene, which encodes a protein belonging to the AraC family of positive transciptional activators [46]. HrpXv may interact with plant inducible promoter regions in the hrp gene cluster. The expression of hrpXv is regulated by hrpG, which was recently discovered to be adjacent to the hrpXv gene [78]. Sequence data for hrpG has revealed that the gene product is a response regulator belonging to the OmpR subclass of two-component signal transducing systems. Therefore, it is conceivable that the hrp gene expression is the result of a cascade of events beginning with a sensor detecting a change in the bacterium’s environment. Specifically, the signal is the availability or change in concentration of unknown plant compounds that are smaller than 1000 Da, heat-stable, organic and hydrophilic [79]. The gene encoding the sensor for HrpG has not be found, and unlike most two-component signal transducing systems, it is not located in close proximity to the gene for the response regulator. Regulation of hrp genes by HrpXv may also be common in Xanthomonas, as homologs of hrpXv were found in other xanthomonads, and those from X. campestris pv. campestris, X. campestris pv. vesicatoria and X. oryzae pv. oryzae were interchangable [80]. Two hrp clusters have been cloned from X. campestris pv. campestris [81,82]. The gene cluster identified by Arlat et al. [81] spans 25 kb, and it was established that the loss of pathogenicity of the hrp mutants is not due to the failure to produce wild-type levels of extracellular enzymes. This provides direct evidence that the regulation and export of extracellular enzymes and the secretion of pathogenicity factors through the Hrp type III secretion apparatus are independent. Similar results have been obtained from another hrp cluster, designated as hrpXc, from X. campestris pv. campestris [82]. The sequences of hrpXc and the hrp cluster reported by Arlat et al. [81] are different [47]. The nucleotide sequences for four hrp clusters from Ralstonia solanacearum, Erwinia amylovora, Pseudomonas syringae pv. syringae and Xanthomonas campestris pv. vesicatoria have been determined [83]. Most of the genes from these clusters are highly conserved, they are all involved in a type III protein secretion pathway, and they show remarkable similarity to flagellar biogenesis components. It has been speculated that the hrp system derived from a flagellar system to permit protein secretion by an extracellular appendage and may have been acquired through horizontal gene transfer in pathogenic bacteria [7]. Recently, Bogdanove et al. [83] proposed harmonizing the nomenclature of hrp genes in phytopathogens with a new designation of ‘hrc’ denoting hypersensitivity response and conserved among pathogenic bacteria. These researchers also proposed standardizing the nomenclature of several known type III secretion pathway genes in Yersinia, as they have shown the highest degree of homology to the hrp genes of plant pathogens. The hrp genes also may function in preventing host recognition or suppressing plant defense responses. The hrp genes of xanthomonads can suppress papilla formation in leaf mes-

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ophyll cells and a hypersensitive response in vascular tissue [84,85]. One hypothesis of how this could occur is that the hrpX gene encodes protein myristoylation activity that could modify the bacterial surface to prevent host recognition [80].

6. Avirulence (avr) genes The discovery that the secretion of extracellular enzymes and polysaccharide is independent of the hrp secretion pathway has led researchers in search of factors interacting with the type-III secretion apparatus. The most obvious candidates are the avirulence (avr) gene products, which determine the specificity of the HR. The cloning of an avr gene from Pseudomonas syringae pv. glycinea and converting compatible strains to elicit HR in a number of soybean cultivars was the first experiment to discover the role of avirulence genes as host range determinants [86,87]. Because the presence of an avirulence gene in a pathogen is related to the HR in the plant host, it follows that avr genes should have a negative effect on the fitness of a bacterium as a pathogen and therefore should be lost over time. However, the continual persistence and widespread occurrence of avirulence genes in bacterial pathogens appears to contradict this hypothesis. Thus, one can speculate that avirulence genes must serve a positive physiological function. It is tempting to speculate that all avr genes fulfill a virulence function, but functional analysis based on marker-exchange experiments in Pseudomonas syringae pathovars have indicated that eight out of ten avr genes examined thus far appear to have no virulence role [88]. Until the physiological functions of avirulence genes have been clearly assigned, the evolutionary significance of avr genes will remain unresolved. Within the Xanthomonas genus, at least 16 avirulence genes have been cloned [88]. With the exception of one avr gene on the chromosome, all of the remaining are plasmid-borne. This is an interesting feature of avirulence genes, as it potentially makes horizontal gene transfer a primary mode of transmission of avr genes in the field under selective pressure, especially when some of the plasmids also encode resistance to chemical bactericides such as copper [89]. In contrast to P. syringae, four out of six avr genes of xanthomonads examined thus far also carry a virulence function, but the rest of the genes remain to be characterized. Evidence of a positive physiological role for an avirulence gene can be observed with avrBs2. The avrBs2 from X. campestris pv. vesicatoria has been found to be highly conserved in the strains of this pathovar and in many other pathovars of X. campestris that do not cause diseases in pepper or tomato [90,91]. Thus far, the avrBs2 gene is also the only Xanthomonas avirulence gene found on the chromosome. Clones from DNA libraries with homology to avrBs2 were identified in X. campestris pv. alfalfae and X. campestris pv. phaseoli, and these were able to complement an avrBs2 mutation in X. campestris pv. vesicatoria. Furthermore, mutation in the avrBs2 of X. campestris pv. vesicatoria had a 100-fold reduction in the virulence toward the susceptible host. Hence, a highly conserved virulence-related mechanism appears to exist in the X. campestris pathovars that also act as a host-range determinant. AvrBs2 has homology to enzymes that synthesize or hydrolyze phosphodiester bonds, and this activity may modify bacterial glucans to help adapt the pathogen to the environment of plant intercellular spaces [92]. Cell-associated glucans have been isolated from X. campestris

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pv. citri [93]. Four other avirulence genes, designated as avrBs1, avrBs3, avrBsT and avrBsT, were cloned from X. campestris pv. vesicatoria. Each avirulence gene-resistant gene interaction produced different HR phenotypes varying in the timing of HR appearance, the intensity of browning, the degree of confluence and the timing and rate of electrolyte leakage in leaves [89]. Research on X. campestris pv. malvacearum also points to an avirulence gene designated as avrb6, which may confer a positive selective advantage to the pathogen [94]. Unlike the other avirulence genes from this pathovar, the presence of avrb6 seems to allow increase water-soaking symptoms during infection. The avirulence genes from this pathovar are all highly related, encoded on a single 90-kb plasmid, and perhaps are members of the same avr gene family. All show strong homology to avrBs3 of X. campestris pv. vesicatoria [95]. The presence of terminal inverted repeats at the boundaries of these avr genes may reflect some form of transposition function, but this is highly speculative. Genes in the avrBs3 family have other common features, such as nucleotide localization sites and heptad repeats similar to leucine zippers, which could be involved in protein–protein or protein–DNA interactions [96]. The interaction of the resistance genes from cotton and avirulence genes of X. campestris pv. malvacearum does not entirely follow the gene-for-gene interaction model. Some of the cotton resistance genes are able to react with multiple avr genes, while others do not. The harbouring of multiple avirulence genes is difficult to explain but may have resulted from horizontal gene transfer events where duplication had taken place. In addition, the involvement of the X. campestris pv. malvacearum avr genes in watersoaking is additive and quantitative [97]. Mutation by marker-exchange experiments in a number of avr genes led to a loss in watersoaking symptoms, while the growth of the pathogen in the host was unaffected. Therefore, because the selective value may be cumulative, the inherent positive selection advantage conferred by avirulence genes may be difficult to realize, especially when each gene contributes differently to the trait. The dual role of pthA as a virulence/avirulence factor from X. citri has proven to be quite complicated [98,99]. pthA was originally considered as a pathogenicity factor, but sequence analysis has revealed that this gene is homologous to the avrBs3 gene from X. campestris pv. vesicatoria [88]. The pthA gene is intriguing, as its product can not only increase the virulence of an opportunistic pathogen of citrus, X. campestris pv. citrumelo, but it could make two avirulent pathovars on citrus, X. campestris pv. phaseoli and X. campestris pv. malvacearum into weak pathogens in citrus. Mutation of pthA in X. citri rendered the pathogen avirulent by affecting its growth in planta.

7. The interaction of hrp and avr genes The recent work of Van den Ackerveken et al. [100] on avrBs3 from X. campestris pv. vesicatoria has advanced the understanding of the interactions between the Hrp-dependent secretion pathway, avr genes and plant-resistance genes. Various working models have been proposed for plant host recognition of avirulence genes, where the plant cell receptors for Avr proteins are presumably located on the plant cell surface [101,102]. However, when AvrBs3 protein was infiltrated into the the intercellular space of pepper leaves, HR symptoms were not apparent. The findings currently point to a mode of protein transport whereby

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Hrp-dependent type III secretion apparatus may be responsible for translocating the AvrBs3 protein into the plant host cell by forming a pilus in an analogous fashion as in Yersinia [100]. This is consistent with the idea that type III secretion occurs only when a pathogen is in close contact with host cells [103]. The AvrBs3 has nuclear localization signals and the product of the plant Bs3 resistance gene is responsible for interacting with the bacterial AvrBs3 protein in the nucleus. The interaction subsequently induces a cascade of effects leading to the HR. This transport pathway may even hold true for most, if not all, Avr proteins, as they all require a Hrp-dependent pathway [104]. A major breakthrough in the understanding of the hrp-dependent type III secretion apparatus was made in Pseudomonas syringae pv. tomato [105]. Using hrp-inducing medium, it was shown that the hrp genes are responsible for the production of a hrp-dependent pilus. The pilus structure provides the first physical link between the plant cell surface and the pathogen, and may serve as the conduit for the translocation of Avr proteins from the pathogen to the plant cytoplasm, not unlike those observed in the T-DNA transmission by the crown gall causal agent, Agrobacterium tumefaciens. The discovery of hrp and avr genes in other bacteria begs the question of whether the biochemical functions of all bacterial elicitation systems are similar. Although Avr proteins are likely to be the main candidates for being the HR elicitors in Xanthomonas, this may not always be the case. The cloning of hrpN from Erwinia amylovora, which encodes a protein called harpinEa, has shown that other HR elicitors are also present in bacterial plant pathogens [106]. Biochemically similar proteins have also found in Pseudomonas syringae pv. syringae (hrpZ for harpinPss) and P. solanacearum (popA1 for PopA1) [107,108]. The primary sequences of these harpin-like HR elicitors have been found to be different. The harpinEa is likely to be the sole HR elicitor present in E. amylovora, as mutations in hrpN eliminated all HR-eliciting ability in the pathogen. In contrast, mutations in hrpZ or popA1 either reduced the HR-eliciting function or did not affect HR-eliciting ability suggesting that other HR elicitors, such as Avr proteins, are also present in Pseudomonas. Thus far, no harpin or harpin-like proteins have been found in Xanthomonas [109]. However, a weak hybridization signal was reported in Southern hybridization blots of X. oryzae pv. oryzae probed with hrpZ raising the possibility that there are proteins similar to harpins in Xanthomonas [110].

8. Conclusions A significant number of Xanthomonas genes have now been shown to be important in virulence. They are all related to compounds that are either surface components or are secreted by the bacterial cell. This is not surprising, as most of the growth of Xanthomonas pathogens is in plant intercellular spaces between parenchyma cells or inside xylem vessels. Therefore, virulence determinants must have some means of contacting host cells. Mutations in xanthomonad secretory pathways have clearly shown the importance of these exported compounds. However, it has been much more difficult to identify the important individual exported components. This is particularly so for the extracellular enzymes. In some cases, this may occur because there are as yet unknown compounds that are exported by these secretion

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pathways, or it may be that a particular combination of secreted factors is required, such as the case of the cumulative effect of some Avr proteins in causing water soaking. One approach to group some of these virulence determinants is to divide them based on the type of secretory system involved. Determinants would be described as being either type I, II or III secretion pathway-dependent. A separate group of virulence determinants would be certain bacterial surface components, such as LPS and EPS, and finally another group of virulence determinants would be composed of those regulatory genes controlling expression of the virulence genes mentioned above. One can hypothesize that the virulence determinants excreted or on the cell surface of xanthomonads could act in one of two possible ways to promote bacterial growth. One would be to obtain nutrients by either altering host metabolism to enhance the release of nutrients from intact host cells into intercellular spaces or by scavenging nutrients through the degradation of polymers associated with plant cells. Another possible function would be to alter host metabolism to prevent recognition or suppress host defense responses. Thus far, there is more supporting evidence for the latter role, but further evidence is needed to determine which is more crucial to the growth of xanthomonads in host tissues. More research is needed to unravel the positive functions of avr genes, as there is accumulating evidence that a significant number of xanthomonad avirulence genes have dual functions in both avirulence and virulence. Avirulence genes have received considerable attention because of their role in determining host specificity. Plants have evolved to respond to these avirulence gene products apparently because these products are convenient chemical signatures to recognize the presence of pathogenic microorganisms. For xanthomonads, all of the avirulence gene products identified thus far are associated with the type III secretory pathway. Perhaps this relationship between the type III pathway and Avr proteins occurred because some of these secreted compounds were always necessary for xanthomonad growth inside plants, and plants could also readily modify cytoplasmic proteins to recognize these bacterial compounds. However, the discovery of the Xa21 resistance gene implies that it is also possible for plants to develop membrane proteins that can recognize xanthomonad extracellular products. With our greater knowledge of the type III secretion pathway and the compounds exported by it, a rethinking of the these exported proteins is perhaps needed. Are all proteins exported by the type III pathway recognized by plant resistance genes? Why are some important for virulence and others are not? Could some of these proteins be secreted into other microorganisms? Could their role in virulence vary between hosts or between varieties of a particular host? One suggestion that would help change our thinking about these proteins is the proposal of Alfano and Collmer [111] to rename the proteins secreted by the type III pathway from Avr to Hop (Hrp-dependent outer protein), which would be analogous to the term Yop (Yersinia outer protein), which are the proteins secreted by a type III pathway of this pathogen. The advantage of the term Hop is that it would include all proteins exported by the type III secretion pathway, including those that would not be recognized by a particular plant and therefore have no avirulence activity. Our knowledge of the virulence of xanthomonads has expanded rapidly in recent years. Many of these virulence factors were completely unsuspected prior to the application of molecular biology techniques, and it has been particularly surprising to find similarities between

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xanthomonads and animal bacterial pathogens, especially as Xanthomonas species are all plant associated. However, significant gaps in our understanding remain. Considering the wide diversity of diseases involved and the long period of coevolution between xanthomonads and plants, it is not surprising that the interaction between these pathogens and their hosts has proven to be so complex.

References [1] Hayward AC. The hosts of Xanthomonas. In: Swings G, Civerolo EL, editors. Xanthomonas. London: Chapman and Hall, 1993. pp. 1–119. [2] Dye DW, Bradbury JF, Goto M, Hayward AC, Lelliott RA, Schroth MN. International standards for naming pathovars of phytopathogenic bacteria and a list of pathovar names and pathotypes strains. Rev Plant Pathol 1980;59:153–68. [3] Vauterin L, Hoste B, Kersters K, Swings J. Reclassification of Xanthomonas. Int J Syst Bacteriol 1995;45:472–89. [4] Vauterin L, Swings J. Are classification and phytopathological diversity compatible in Xanthomonas? J Ind Microbiol Biotechnol 1997;19:77–82. [5] Daniels MJ. Molecular genetics of host-pathogen interactions. In: Palacios R, Vema DPS, editors. Molecular Genetics of Plant Microbe Interactions. St. Paul: APS Press, 1988. pp. 229–34. [6] Rudolph K. Infection of the plant by Xanthomonas. In: Swings JG, Civerolo EL, editors. Xanthomonas. London: Chapman and Hall, 1993. pp. 193–264. [7] Alfano JR, Collmer A. Bacterial pathogens in plants: life up against the wall. Plant Cell 1996;8:1683–98. [8] Dow JM, Scofield G, Trafford K, Turner PC, Daniels MJ. A gene cluster in Xanthomonas campestris pv. campestris required for pathogenicity controls the excretion of polygalacturonase lyase and other enzymes. Physiol Mol Plant Pathol 1987;31:261–71. [9] Dums F, Dow JM, Daniels MJ. Structural characterization of protein secretion genes of the bacterial phytopathogen Xanthomonas campestris pathovar campestris: relatedness to secretion systems of other gramnegative bacteria. Mol Gen Genet 1991;229:357–64. [10] Gough CL, Dow JM, Barber CE, Daniels MJ. Cloning of two endoglucanase genes of Xanthomonas campestris pv. campestris: analysis of the role of the major endoglucanase in pathogenesis. Mol PlantMicrobe Interact 1988;1:275–81. [11] Gough CL, Dow JM, Keen J, Henrissat B, Daniels MJ. Nucleotide sequence of the engXCA gene encoding the major endoglucanase of Xanthomonas campestris pv. campestris. Gene 1990;89:53–9. [12] Beaulieu C, Minsavage GV, Canteros BI, Stall RE. Biochemical and genetic analysis of a pectate lyase gene from Xanthomonas campestris pv. vesicatoria. Mol Plant-Microbe Interact 1991;4:446–51. [13] Dow JM, Mulligan DE, Jamieson L, Barber CE, Daniels MJ. Molecular cloning of a polygalacturonase lyase gene form Xanthomonas campestris pv. campestris and role of the gene product in pathogenicity. Physiol Mol Plant Pathol 1989;35:113–20. [14] Liao CH, Gaffney TD, Bradley SP, Wong LJ. Cloning of a pectate lyase gene from Xanthomonas campestris pv. malvacearum and comparison of its sequence relationship and pel genes of soft rot Erwinia and Pseudomonas. Mol Plant-Microbe Interact 1996;9:14–21. [15] Dow JM, Clarke BR, Milligan DE, Tang JL, Daniels MJ. Extracellular proteases from Xanthomonas campestris pv. campestris, the black rot pathogen. Appl Environ Microbiol 1990;56:2994–8. [16] Liu Y-N, Tang J-L, Clarke BR, Dow JM, Daniels MJ. A multipurpose broad host range cloning vector and its use to characterize an extracellular protease gene of Xanthomonas campestris pathovar campestris. Mol Gen Genet 1990;220:433–40. [17] Tang JL, Gough CL, Barber CE, Dow JM, Daniels MJ. Molecular cloning of protease gene(s) from Xanthomonas campestris pv. campestris expression in Escherichia coli and role in pathogenicity. Mol Gen Genet 1987;210:443–8.

504

J.W.Y.F. Chan, P.H. Goodwin / Biotechnology Advances 17 (1999) 489–508

[18] Hwang I, Lim SM, Shaw PD. Cloning and characterization of pathogenicity genes from Xanthomonas campestris pv. glycines. J Bacteriol 1992;174:1923–31. [19] Hu NT, Hung MN, Huang AM, Tsai HF, Yang BY, Chow TY, Tseng H. Molecular cloning, characterization and nucleotide sequence of the gene for secreted a-amylase from Xanthomonas campestris pv. campestris. J Gen Microbiol 1992;138:1647–55. [20] Newman MA, Conrads-Strauch J, Schofield G, Daniels MJ, Dow JM. Defense-related gene induction in Brassica campestris in response to defined mutants of Xanthomonas campestris with altered pathogenicity. Mol Plant-Microbe Interact 1994;4:553–63. [21] Zhang L, Birch RG. The genes for albicidin detoxification from Pantoea dispersa encodes an esterase and attenuates pathogenicity of Xanthomonas albilineans to sugarcane. Proc Natl Sci USA 1997;94:9984–9. [22] Andrew TR. Application of xanthan gum in foods and related products. In: Sandford PA, Laskin A, editors. Extracellular Microbial Polysaccharides. Washington, DC: American Chemical Society, 1977. pp. 231–41. [23] Baird JK, Sandford PA, Cottrell IW. Industrial applications of some new microbial polysaccharides. Bio/ Technology 1983;1:778–83. [24] Kennedy JF, Bradshaw IJ. Production, properties and applications of xanthan. In: Bushell ME, editor. Progress in Industrial Microbiology: Modern Applications of Traditional Biotechnologies, vol. 19. New York: Elsevier Science Publishing Company, Inc, 1984. pp. 319–71. [25] Sutherland IW. Xanthan. In: Swings JG, Civerolo EL, editors. Xanthomonas. London: Chapman and Hall 1993. pp. 363–88. [26] Sutherland IW. Biosynthesis and composition of gram-negative bacterial extracellular and wall polysaccharides. Annu Rev Microbiol 1985;39:243–70. [27] Hötte B, Rath-Arnold I, Pühler A, Simon R. Cloning and analysis of a 35.3-kilobase DNA region involved in exopolysaccharide production by Xanthomonas campestris pv. campestris. J Bacteriol 1990;172:2804–7. [28] Vanderslice RW, Doherty DW, Capage MA, Betlach MR, Hasller RA, Henderson NM, Ryan-Graniero J, Techlenburg M. Genetic engineering of polysaccharide in Xanthomonas campestris. In: Crescenzi V, Dea ICM, Stivola SS, editors. Recent Development in Industrial Polysaccharides: Biomedical and Biotechnological Advances. New York: Gordon and Breach Science Publishers, 1989. pp. 145–56. [29] Barrère GC, Barber CE, Daniels MJ. Molecular cloning of genes involved in the production of the extracellular polysaccharide xantha by Xanthomonas campestris pv. campestris. Int J Biol Macromol 1986; 8:372–4. [30] Harding NE, Cleary JM, Cabanas DK, Rosen IG, Kang KS. Genetic and physical analysis of a cluster of genes essential for xanthan gum biosynthesis in Xanthomonas campestris. J Bacteriol 1987;169:2854–61. [31] Thorne L, Tansey L, Pollock TJ. Clustering of mutations blocking synthesis of zanthan gum by Xanthomonas campestris. J Bacteriol 1987;169:3593–600. [32] Katzen F, Becker A, Zorreguieta A, Pühler A, Ilepi L. Promoter analysis of Xanthomonas campestris pv. campestris gum operon directing biosynthesis of the xanthan polysaccharide. J Bacteriol 1996;178:4313–8. [33] Katzen F, Ferreiro DU, Oddo CG, Ielmini V, Becker A, Pühler A, Ielpi L. Xanthomonas campestris pv. campestris gum mutants: effects on xanthan biosynthesis and plant virulence. J Bacteriol 1998;180:1607–17. [34] Chou FL, Chou HC, Lin YS, Yang BY, Lin NT, Weng SF, Tseng YH. The Xanthomonas campestris gumD gene required for synthesis of xanthan gum is involved in normal pigmentation and virulence in causing black rot. Biochem Biophys Res Commun 1997;233:265–9. [35] Ramírez ME, Fuckiovsky L, Garcia-Jiménez F, Quintero R, Galindo E. Xanthan gum production by altered pathogenicity variants of Xanthomonas campestris. Appl Microbiol Biotechnol 1988;29:5–10. [36] Tang JL, Feng JZ, Li QQ, Wen HX, Zhou DL, Wilson TJG, Dow JM, Ma QS, Daniels MJ. Cloning and characterization of the rpfCi gene of Xanthomonas oryzae pv. oryzae: involvement in expopolysaccharide production and virulence to rice. Mol Plant-Microbe Interact 1996;9:664–6. [37] Tang JL, Lui Y-N, Barber CE, Dow JM, Wootton JC, Daniels MJ. Genetic and molecular analysis of a cluster of rpf genes involved in positive regulation of synthesis of extracellular enzymes and polysaccharide in Xanthomonas campestris pathovar campestris. Mol Gen Genet 1991;226:409–17. [38] Coplin DL, Cook D. Molecular genetics of extracellular polysaccharide biosynthesis in vascular phytopathogenic bacteria. Molecular Plant-Microbe Interact 1990;3:271–9. [39] Newman M-A, Daniels MJ, Dow JM. The activity of Lipid A and core components of bacterial li-

J.W.Y.F. Chan, P.H. Goodwin / Biotechnology Advances 17 (1999) 489–508

[40]

[41] [42] [43] [44] [45]

[46] [47]

[48] [49]

[50]

[51] [52]

[53] [54]

[55] [56]

[57] [58]

[59] [60]

505

popolysaccharides in the prevention of the hypersensitive response in pepper. Mol Plant-Microbe Interact 1997;10:926–8. Kingsley MT, Gabriel DW, Marlow GC, Roberts PD. The opsX locus of Xanthomonas campestris affects host range and biosynthesis of lipopolysaccharide and extracellular polysaccharide. J Bacteriol 1993;175:5839–50. Dow JM, Osbourn AE, Greer-Wilson TJ, Daniels MJ. A locus determining pathogenicity of Xanthomonas campestris is involved in lipopolysaccharide biosynthesis. Mol Plant-Microbe Interact 1995;8:768–77. Boher B, Nicole M, Potin M, Geiger JP. Extracellular polysccharides from Xanthomonas axonopodis pv. manihotis interact with cassava cell walls during pathogenesis. Mol Plant-Microbe Interact 1997;10:803–11. Albright LM, Huala E, Ausubel FM. Prokaryotic signal tranduction mediated by sensor and regulator protein pairs. Annu Rev Genet 1989;23:311–36. Alex LA, Simon ML. Protein histidine kinases and signal transduction in prokaryotes and eukaryotes. Trends Genet 1994;10:133–8. Osbourne AE, Clarke BR, Stevens BJH, Daniels MJ. Use of oligonucleotide probes to identify members of two-component regulatory systems in Xanthomonas campestris pv. campestris. Mol Gen Genet 1990;222:145–51. Wengelnik K. Bonas U. HrpXv, an AraC-type regulator activates expression of five out of six loci in the hrp cluster of Xanthomonas campestris pv. vesicatoria. J Bacteriol 1996;178:3462–9. Dow JM, Daniels MJ. Pathogenicity determinants and global regulation of pathogenicity of Xanthomonas campestris pv. campestris. In: Dangl JL, editor. Current Topics in Microbiology and Immunology. Heidelberg, Germany: Springer Verlag, 1994. pp. 29–41. Turner P, Barber C, Daniels M. Evidence of clustered pathogenicity genes in Xanthomonas campestris pv. campestris. Mol Gen Genet 1985;199:338–43. Sawczyz MK, Barber CE, Daniels MJ. The role in pathogenicity of some related genes in Xanthomonas campestris pathovars campestris and translucens: a shuttle strategy for cloning genes required for pathogenicity. Mol Plant-Microbe Interact 1989;2:249–55. Barber CE, Tang JL, Feng JX, Pan MQ, Wilson TJG, Slater H, Dow JM, Williams P, Daniels MJ. A novel regulatory system required for pathogenicity of Xanthomonas campestris is mediated by a small diffusible signal molecule. Mol Microbiol 1997;24:555–66. Poplawsky AR, Chun W. pigB determines a diffusible factor needed for extracellular polysaccharide slime and xanthomonadin production in Xanthomonas campestris pv. campestris. J Bacteriol 1997;179:439–44. Poplawsky AR, Chun W, Slater H, Daniels MJ, Dow JM. Synthesis of extracellular polysaccharide, extracellular enzymes, and xanthomonadin in Xanthomonas campestris: evidence for the involvement of two intercellular regulatory signals. Mol Plant-Microbe Interact 1998;11:68–70. Chun W, Cui H, Poplawsky AR. Purification, characterization and biological role of a pheromone produced by Xanthomonas campestris pv. campestris. Physiol Mol Plant Pathol 1997;51:1–14. Tang JL, Gough CI, Daniels MJ. Cloning of genes involved in negative regulation of production of extracellular enzymes and polysaccharide of Xanthomonas campestris pathovar campestris. Mol Gen Genet 1990;222:157–60. Osbourne AE, Barber CE, Daniels MJ. Identification of plant-induced genes of the bacterial pathogen Xanthomonas campestris pathovar campestris using a promoter-probe plasmid. EMBO J 1987;82:23–8. De Crécy-Lagard V, Glaser P, Lejeune P, Sismeiro O, Barber CE, Daniels MJ, Danchin A. A Xanthomonas campestris pv. campestris protein similar to catabolite activation factor is involved in regulation of phytopathogenicity. J Bacteriol 1990;172:5877–83. Dong Q, Ebright RH. DNA binding specificity and sequence of Xanthomonas campestris catabolite gene activator protein-like protein. J Bacteriol 1992;174:5457–61. Köplin R, Arnold W, Hötte B, Simon R, Wang G, Pühler A. Genetics of xanthan production in Xanthomonas campestris: the xanA and xanB genes are involved in UPD-glucose and GPD-mannose biosynthesis. J Bacteriol 1992;174:191–9. Marzocca MP, Harding NE, Petroni A, Cleary JM, Ielpi L. Location and cloning of ketal pyruvate transferase gene of Xanthomonas campestris. J Bacteriol 1991;173:519–24. Hu NT, Hung MN, Chiou SJ, Tang F, Chiang DC, Huang HY, Wu CY. Cloning and characterization of a

506

[61] [62] [63] [64] [65] [66] [67]

[68] [69] [70] [71]

[72]

[73]

[74]

[75]

[76] [77] [78]

[79] [80]

[81]

J.W.Y.F. Chan, P.H. Goodwin / Biotechnology Advances 17 (1999) 489–508 gene required for the secretion of extracellular enzymes across the outer membrane by Xanthomonas campestris pv. campestris. J Bacteriol 1992;174:2679–87. Thorne L, Gosink KK, Pollock TJ. Mutants of Xanthomonas campestris defective in secretion of extracellular enzymes. J Ind Microbiol 1989;4:135–44. Hu NT, Lee PF, Chen C. The type IV pre-pilin leader peptidase of Xanthomonas campestris pv. campestris is functional without conserved cysteine residues. Mol Microbiol 1995;18:769–77. Salmond GPC. Secretion of extracellular virulence factors by plant pathogenic bacteria. Annu Rev Phytopathol 1994;32:181–200. Dixon RA, Harrison MJ, Lamb CJ. Early events in the activation of plant defense responses. Annu Rev Phytopathol 1994;32:479–501. Gabriel DW, Rolfe BG. Working models of specific recognition in plant-microbe interactions. Annu Rev Phytopathol 1990;28:365–91. Hammond-Kosack KE, Jones JDG. Plant disease resistance genes. Annu Rev Plant Physiol Mol Biol 1997;48:575–607. Yoshimura S, Yamanouchi U, Katayose Y, Toki S, Wang Z-W, Kono I, Kurata N, Yano M, Iwata N, Sasaki T. Expression of Xa1, a bacterial blight-resistance gene in rice is induced by bacterial inoculation. Proc Natl Acad Sci USA 1998;95:1663–8. Song WY, Wang GL, Chen LL, Kin HS, Holsten T, Wang B, Zhai WZ, Zhu LH, Fauquet C, Ronald P. The rice disease resistance gene, Xa21, encodes a receptor-like protein. Science 1995;270:1804–6. Lindgren PB, Peet RC, Panopoulos NJ. Gene cluster of Pseudomonas syringae pv. phaseolicola controls pathogenicity on bean plants and hypersensitivity on nonhost plants. J Bacteriol 1986;168:512–22. Niepold F, Anderson D, Mills D. Cloning determinants of pathogenesis from Pseudomonas syringae pathovar syringae. Proc Natl Acad USA 1985;82:406–10. Stall RE, Minsavage GV. The use of hrp genes to identify opportunistic xanthomonads. In: Klement Z, editor. Proceedings of the 7th International Conference of Plant Pathogenic Bacteria. Budapest, Hungary: Académia Kiadó, 1989. pp. 369–74. Leite RPJ, Minsavage GV, Bonas U, Stall RE. Detection and identification of phytopathogenic Xanthomonas strains by amplification of DNA sequences related to the hrp gene of Xanthomonas campestris pv. vesicatoria. Appl Environ Microbiol 1994;60:1068–77. Bonas U, Schulte R, Fenselau S, Minsavage GV, Staskawicz BJ, Stall RE. Isolation of a gene cluster from Xanthomonas campestris pv. vesicatoria that determines pathogenicity and the hypersensitive response on pepper and tomato. Mol Plant-Microbe Interact 1991;4:81–8. Waney VR, Kingsley MT, Gabriel DW. Xanthomonas campestris pv. translucens genes determining hostspecific virulence and general virulence on cereals identified by Tn5-gusA insertion mutagenesis. Mol Plant-Microbe Interact 1991;4:623–7. Fenselau S, Balbo I, Bonas U. Determinants of pathogenicity in Xanthomonas campestris pv. vesicatoria are related to proteins involved in secretion in bacterial pathogens of animals. Mol Plant-Microbe Interact 1992;5:390–6. Bonas U. hrp genes of phytopathogenic bacteria. Curr Top Microbiol Immunol 1994;192:79–98. Huguet E, Bonas U. hrpF of Xanthomonas campestris pv. vesicatoria encodes an 87-KDa protein with homology to NolX of Rhizobium fredii. Mol Plant-Microbe Interact 1997;10:488–98. Wengelnik K, Van den Ackerveken V, Bonas U. HrpG, a key hrp regulatory protein in Xanthomonas campestris pv. vesicatoria is homologous to two-component response regulators. Mol Plant-Microbe Interact 1996;9:704–12. Schulte R, Bonas U. Expression of the Xanthomonas campestris pv. vesicatoria hrp gene cluster, which determines pathogenicity and hypersensitivity on pepper and tomato, is plant inducible. J Bacteriol 1992;174:815–23. Kamdar HV, Kamoun S, Kado CI. Restoration of pathogenicity of avirulent Xanthomonas oryzae pv. oryzae and X. campestris pathovars by reciprocal complementation with the hrpXo and hrpXc genes and identification of HrpX function by sequence analysis. J Bacteriol 1993;175:2017–25. Arlat M, Gough CL, Barber CE, Boucher C, Daniels MJ. Xanthomonas campestris contains a cluster of hrp genes related to the larger hrp cluster of Pseudomonas solanacearum. Mol Plant-Microbe Interact 1991;4:593–601.

J.W.Y.F. Chan, P.H. Goodwin / Biotechnology Advances 17 (1999) 489–508

507

[82] Kamoun S, Kado CI. A plant inducible gene of Xanthomonas campestris pv. campestris encodes an exocellular component required for growth in the host and hypersensitivity on nonhosts. J Bacteriol 1990;172:5165–72. [83] Bogdanove AJ, Beer SV, Bonas U, Boucher CA, Collmer A, Coplin DL, Cornelis GR, Huange HC, Hutcheson SW, Panopoulos NJ, Van Gijsegem F. Unified nomenclature for broadly conserved hrp genes of phytopathogenic bacteria. Microbiology 1996;20:681–3. [84] Brown I, Mansfield J, Bonas U. hrp gene in Xanthomonas campestris pv. vesticatoria determine ability to suppress papilla deposition in pepper mesophyll cells. Mol Plant-Microbe Interact 1995;8:825–36. [85] Kamoun S, Kamdar HV, Tola E, Kado CI. Incompatible interactions between crucifers and Xanthomonas campestris involve a vascular hypersensitive response role of the hrpX locus. Mol Plant-Microbe Interact 1992;5:22–33. [86] Staskawicz BJ, Dahlbeck D, Keen NT. Cloned avirulence gene of Pseudomonas syringae pv. glycinea determines race-specific incompatibility on Glycine max (L). Proc Natl Acad Sci USA 1984;81:6024–8. [87] Staskawicz B, Dahlbeck D, Keen N, Napoli C. Molecular characterization of cloned avirulence genes from race 0 and race 1 of Pseudomonas syringae pv. glycinea. J Bacteriol 1987;169:5789–94. [88] Dangl JL. The enigmatic avirulence genes of phytopathogenic bacteria. In: Dangl JL, editor. Bacterial Pathogenesis of Plants and Animals: Molecular and Cellular Mechanisms. Berlin: Springer Verlag, 1994. pp. 99–117. [89] Minsavage GV, Dahlbeck D, Whalen MC, Kearney B, Bonas U, Staskawicz BJ, Stall RE. Gene-for-gene relationships specifying disease resistance in Xanthomonas campestris pv. vesticatoria-pepper interactions. Mol Plant-Microbe Interact 1990;3:41–7. [90] Kearney B, Staskawicz BJ. Widespread distribution and fitness contribution of Xanthomonas campestris avirulence gene avrBs2. Nature 1990;346:385–6. [91] Mazzola M, Leach JE, Nelson R. Analysis of the interaction between Xanthomonas oryzae pv. oryzae and the rice cultivars IR24 and IRBB21. Phytopathology 1994;84:392–7. [92] Swords KMM, Dahlbeck D, Kearney B, Roy M, Staskawicz BJ. Spontaneous and induced mutations in a single open reading frame alter both virulence and avirulence in Xanthomonas campestris pv. vesicatoria avrBs2. J Bacteriol 1996;178:4661–9. [93] Talaga P, Stahl B, Wieruszeski JM, Hillenkamp F, Tsuyumu S, Lippens G, Bohin JP. Cell-associated glucans of Burkholderia solonacearum and Xanthomonas campestris pv. citri: a new family of periplasmic glucans. J Bacteriol 1996;178:2263–71. [94] De Feyter R, Gabriel DW. At least six avirulence genes are clustered on a 90-kilobase plasmid in Xanthomonas campestris pv. malvacearum. Mol Plant-Microbe Interact 1991;4:423–32. [95] De Feyter R, Yang Y, Gabriel DW. Gene-for-genes interactions between cotton R genes and Xanthomonas campestris pv. malvacearum avr genes. Mol Plant-Microbe Interact 1993;6:225–37. [96] Vivian A, Gibbon MJ. Avirulence genes in plant-pathogenic bacteria: signals or weapons? Microbiology 1997;143:693–704. [97] Yang YO, Yuan QP, Gabriel DW. Waterlogging function(s) of XCM1005 are redundantly encoded by members of the Xanthomonas avr/pth gene family. Mol Plant-Microbe Interact 1996;9:105–13. [98] Swarup S, De Feyter R, Brlansky RH, Gabriel DW. A pathogenicity locus from Xanthamonas citri enables strains from several pathovars of X. campestris to elicit cankerlike lesions on citrus. Phytopathology 1991;81:802–9. [99] Swarup S, Yang Y, Kingsley MT, Gabriel DW. A Xanthomonas citri pathogenicity gene, pthA, pleiotropically encodes gratuitous avirulence on nonhosts. Mol Plant-Microbe Interact 1992;5:204–13. [100] Van den Ackerveken G, Marois E, Bonas U. Recognition of the bacterial avirulence protein AvrBs3 occurs inside the host plant cell. Cell 1996;87:1307–16. [101] Gopalan S, He SY. Bacterial genes involved in the elicitation of hypersensitive response and pathogenesis. Plant Dis 1996;80:604–10. [102] He SY. Elicitation of plant hypersensitive response by bacteria. Plant Physiol 1996;112:865–9. [103] Mesaca J, Strauss EJ. Molecular mechanisms of bacterial virulence: type III secretion and pathogenicity islands. Emerg Infect Dis 1996;2:271–88. [104] Parker JE, Coleman MJ. Molecular intimacy between protein specifying plant-pathogen recognition. Trends Biotechnol 1997;22:291–6.

508

J.W.Y.F. Chan, P.H. Goodwin / Biotechnology Advances 17 (1999) 489–508

[105] Roine E, Wei W, Yuan J, Nurmiaho-Lassila EL, Kalkkinen N, Romantschuk M, He SY. Hrp pilus: an hrpdependent bacterial surface appendage produced Pseudomonas syringae pv. tomato DC3000. Proc Natl Acad Sci USA 1997;94:3459–64. [106] Wei ZM, Laby RJ, Zumoff CH, Bauer DW, He SY, Collmer A, Beer SV. Harpin, elicitor of the hypersensitive response produced by the plant pathogen Erwinia amylovora. Science 1992;257:85–8. [107] Arlat M, van Gijsegem F, Heut JC, Pernollet JC, Boucher CA. PopA1, a protein which induces a hypersensitivity-like response on specific Petunia genotypes, is secreted via the Hrp pathway of Pseudomonas solanacearum. EMBO J 1994;13:543–53. [108] He SY, Huang HC, Collmer A. Pseudomonas syringae pv. syringae harpinPss: a protein that is secreted via the Hrp pathway and elicits the hypersensitive response in plants. Cell 1993;73:1255–66. [109] Gabriel DW. Targeting of protein signals from Xanthomonas to the plant nucleus. Trends Plant Sci 1997;2:204–6. [110] Jin AL, Liu NZ, Qui JL, Li BD, Wang J. A truncated fragment of harpinpss induces systemic resistance to Xanthomonas campestris pv. oryzae in rice. Physiol Mol Plant Pathol 1997;51:243–57. [111] Alfano JR, Collmer A. The type III (hrp) secretion pathway of plant pathogenic bacteria: trafficking harpins, Avr proteins, and death. J Bacteriol 1997;179:5655–62.