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Research Article
1 September 2005

Phylogenetic Characterization of Virulence and Resistance Phenotypes of Pseudomonas syringae


Individual strains of the plant pathogenic bacterium Pseudomonas syringae vary in their ability to produce toxins, nucleate ice, and resist antimicrobial compounds. These phenotypes enhance virulence, but it is not clear whether they play a dominant role in specific pathogen-host interactions. To investigate the evolution of these virulence-associated phenotypes, we used functional assays to survey for the distribution of these phenotypes among a collection of 95 P. syringae strains. All of these strains were phylogenetically characterized via multilocus sequence typing (MLST). We surveyed for the production of coronatine, phaseolotoxin, syringomycin, and tabtoxin; for resistance to ampicillin, chloramphenicol, rifampin, streptomycin, tetracycline, kanamycin, and copper; and for the ability to nucleate ice at high temperatures via the ice-nucleating protein INA. We found that fewer than 50% of the strains produced toxins and significantly fewer strains than expected produced multiple toxins, leading to the speculation that there is a cost associated with the production of multiple toxins. None of these toxins was associated with host of isolation, and their distribution, relative to core genome phylogeny, indicated extensive horizontal genetic exchange. Most strains were resistant to ampicillin and copper and had the ability to nucleate ice, and yet very few strains were resistant to the other antibiotics. The distribution of the rare resistance phenotypes was also inconsistent with the clonal history of the species and did not associate with host of isolation. The present study provides a robust phylogenetic foundation for the study of these important virulence-associated phenotypes in P. syringae host colonization and pathogenesis.
Pseudomonas syringae is one of the preeminent model systems for the study of host specificity and virulence. This gram-negative plant-pathogenic bacterium is the causal agent of a variety of bacterial spot, speck, and blight diseases on a wide range of plant hosts, including (but not limited to) apples, beets, beans, cabbage, cucumbers, flowers, oats, olives, peas, tobacco, tomato, and rice (25). Isolates of P. syringae are taxonomically subdivided into pathogenic varieties known as pathovars, based largely on their host of isolation. The tremendous diversity of hosts and disease symptomatology found in this species presents a unique opportunity to investigate the factors that determine host specificity.
P. syringae uses an impressive variety of virulence-associated systems during the course of its host interactions. These systems produce toxins, ice nucleation proteins, antimicrobial resistance, and secreted effectors. The best-studied virulence-associated factors are the effector proteins secreted through the type III secretion system, which both restrict and promote specific pathogen-host interactions (1, 21, 26, 30, 45). Also well studied, although perhaps less well understood, are the systems that produce toxins, nucleate ice, and confer antimicrobial resistance.
P. syringae produces four primary toxins: coronatine, phaseolotoxin, syringomycin, and tabtoxin (5). All four contribute to chlorosis (yellowing of the leaf tissue typically as a result of chloroplast disruption) or necrosis, while phaseolotoxin has also been implicated in increased pathogen growth and spread in planta. The mode of actions of these toxins is as diverse as their chemical bases. Coronatine is a polyketide molecule that mimics methyl-jasmonate, a key host signaling molecule. Phaseolotoxin is a sulfodiaminophosphinyl peptide that disrupts the urea cycle, thereby causing arginine deficiencies. Syringomycin is one of a related class of lipodepsinonapeptides that causes electrolyte leakage via pores formed in the host plasma membrane. Tabtoxin is a β-lactam that inhibits glutamine synthesis. Although all of these toxins have been shown to modulate virulence, none are essential for the disease process (5), and there is still no consensus on their overall importance in pathogenesis (14).
Pseudomonads in general have a reputation for being highly resistant to antimicrobial compounds (37), and P. syringae is no exception. Antimicrobials such as copper (11) and streptomycin (15) have been used for decades to control P. syringae infections of crop plants. P. syringae strains also come into contact with medically important antibiotics, and their associated resistance genes, that spread in the natural environment (37).
P. syringae also produces an extremely effective protein that facilitates the formation of ice crystals at temperatures 2 to 8°C higher than would naturally occur on the leaf surface. This protein is responsible for millions of dollars in crop losses each year. An attempt to create a genetically modified P. syringae strain that would act as a biocontrol antagonist against naturally occurring, ice nucleating, conspecific bacteria was the impetus for the first field trial of a genetically modified organism (23). P. syringae and its ice-nucleating protein are also key components of the artificial snow-making industry (2). Despite this interest, we know very little about the actual role played by this protein in the disease process.
Despite P. syringae's reputation for toxin production, ice nucleation, and antimicrobial resistance, very few studies have investigated the distribution of these important virulence-associated phenotypes at the broad species level. Furthermore, there have been no studies of these phenotypes based on a rigorous phylogenetic framework. Volksch and Weingart (48) produced the most comprehensive survey of toxin production among P. syringae strains. They assayed 75 strains and found toxin production to be widespread among P. syringae pathovars but did not attempt to determine whether it was associated with the host of isolation or with the evolutionary history of the strains (48).
Sarkar and Guttman (40) recently used multilocus sequence typing (MLST) to characterize the core genome of P. syringae. The core genome consists of genes ubiquitously found among all strains of a bacterial species and typically includes housekeeping genes and RNAs that are essential for the survival of the organism. The core genome is less prone to horizontal gene transfer and therefore provides the best indication of the clonal evolutionary history of a bacterial species (22). In contrast to the core genome, the flexible genome consists of genes that vary among strains within a species. These genes typically encode proteins that are responsible for adaptation to specific niches and the many mobile elements that move in and out of genomes. The flexible genome largely evolves through horizontal genetic exchange (i.e., through gene acquisition and loss).
The MLST analysis of Sarkar and Guttman (40) used the DNA sequences of seven housekeeping genes to determine the evolutionary history of 60 P. syringae strains spanning the diversity of the species complex. These authors found the core genome of P. syringae to be highly clonal (not prone to recombination or horizontal gene transfer) and therefore an excellent indicator of the evolutionary history of the species. These authors also determined that the species has maintained roughly a constant population size through time, indicating that it is probably an endemic pathogen and that the genetic variation in the core genome is only weakly associated with the host of isolation. This phylogenetic analysis now provides us with a framework to assess the role and importance of horizontal gene transfer in facilitating the adaptation of P. syringae strains to their hosts.
In the present study, we used functional, phenotypic assays to survey the ability of 95 P. syringae strains to produce toxins, nucleate ice, and resist antimicrobial agents. We also refined the MLST protocol so that it could be used on nearly any fluorescent pseudomonad. Furthermore, we reduced the number of MLST loci used for typing from seven to four, which dramatically increased the rate at which strains could be typed, without any significant reduction in the phylogenetic resolution. We then mapped the toxin and resistance phenotypes onto the phylogeny of the core genome and showed that most of these phenotypes were distributed in a manner that was not consistent with a clonal evolutionary process. The present study provides a robust, phylogenetic foundation for studies of host adaptation and virulence in P. syringae.


Bacterial and yeast strains.

Ninety-five P. syringae strains were used in the present study (Table 1). (Supplementary information on these strains can be found at .) Our definition of strain is any bacterial isolate that was collected at a unique time and place. Efforts were made to avoid using isolates collected within a year of each other by the same individual. All P. syringae strains were grown in King's B (KB) medium (27) at 30°C. Escherichia coli N99, obtained from B. E. Funnell (University of Toronto, Toronto, Ontario, Canada), was grown in Luria-Bertani (LB) medium (3) at 37°C. Rhodotorula pilimanae MUCL3039, obtained from A. Bultreys (Ministère des Classes Moyennes et de l'Agriculture, Gembloux, Belgium), was grown in potato dextrose broth medium (3) at 30°C.


The four housekeeping genes sequenced were rpoD, encoding sigma factor 70; gyrB, encoding DNA gyrase B; gltA (also known as cts), encoding citrate synthase; and gapA, encoding glyceraldehyde-3-phosphate dehydrogenase. These loci are a subset of the seven used in the original P. syringae MLST paper (40) and were chosen because they consistently provide robust data, and their combined level of polymorphism is sufficient to reliably resolve evolutionary relationships.
The MLST primers used for DNA amplification and sequencing of the four loci (Table 2) were modified from the original MLST publication (40). These degenerate primers were designed based on global multiple sequence alignments from a wide range of fluorescent Pseudomonads. As such, they appear to be useful for MLST typing any fluorescent pseudomonad (P. W. Wang, R. L. Morgan, and D. S. Guttman, unpublished data).
The MLST protocol has been described by Sarkar and Guttman (40). Minor modifications to the PCRs include the use of only 50 ng of template genomic DNA and the addition of dimethyl sulfoxide (Fisher) at a final concentration of 5%. DNA sequencing was performed as described previously (40), except that Betaine (Sigma) was added to the sequencing mix at a final concentration of 1 M. Shared regions for each locus from all strains ranged in size from 494 to 529 bp of double-stranded sequence. Sequences from each locus were aligned by using CLUSTAL W (44) and were trimmed to their minimal shared length in GeneDoc ( ).
Neighbor-joining and maximum-likelihood phylogenetic analyses were performed on the individual and combined datasets by using MEGA version 2.1 (29), PAUP* version 4.0b10 for UNIX (43), and PHYLIP version 3.6.2 (18). The trees were rooted with orthologous sequences from Pseudomonas fluorescens Pf0-1 (U.S. Department of Energy, Joint Genome Institute), although this sequence is not presented in Fig. 1 to improve clarity. Analyses were performed as described by Sarkar and Guttman (40).

Tabtoxin determination.

Production of tabtoxin was determined by an agar plate diffusion test with E. coli N99 as the indicator strain (19, 46). E. coli N99 was grown overnight in LB medium at 37°C and harvested by centrifugation. The pellet was washed and resuspended in 10 ml of sterile 0.9% NaCl at an optical density at 600 nm (OD600) of 0.2. Two milliliters of 0.7% molten mineral salts-glucose (MG) (46) agar (maintained at 45°C) was mixed with 2 ml of E. coli and poured onto MG agar plates. MG-glutamine plates were made by overlaying the E. coli MG soft agar mixture with 17 μmol of glutamine (33). Next, 10 μl of an overnight culture of P. syringae grown in MG medium was spotted onto the MG-E. coli and MG-glutamine-E. coli plates, followed by incubation at room temperature for 48 h. Strains were scored as positive for tabtoxin when there was a zone of inhibition surrounding the P. syringae colonies on the MG plates but not surrounding the corresponding colonies on the MG-glutamine plates.

Phaseolotoxin production.

Phaseolotoxin production was determined by using a method modified from Staskawicz and Panopoulos (42). E. coli N99 was grown in Davis minimal medium (3) for 48 h at 37°C. A 2-ml portion of culture was mixed with 2 ml of 2% molten agar in water and overlaid on Davis minimal medium plates (42). P. syringae strains were grown in minimal A medium (36) for 48 h at 30°C, and 10 μl of the P. syringae culture was spotted onto the E. coli test plates. The presence of phaseolotoxin was characterized by a zone of inhibition surrounding the P. syringae colonies after 24 h.

Syringomycin (lipodepsipeptide) production.

We assayed for syringomycin production by using a general method for detecting lipodepsipeptides (9). Twenty microliters of a P. syringae overnight culture was spotted onto potato dextrose agar (3), followed by incubation for 48 h at 30°C. Subsequently, the plates were sprayed with an overnight culture of Rhodotorula pilimanae (9) and incubated for 24 h at room temperature. The presence of lipodepsipeptide was characterized by the development of a zone of inhibition surrounding the P. syringae colonies.

Coronatine production.

Production of coronatine was determined by a semiquantitative potato disk bioassay (47). Fifty microliters of an overnight P. syringae culture was added to 1 ml of Hoitink and Sinden medium (HSC) (24), and this was followed by incubation on a 250-rpm rotary shaker at 20°C for 4 days. One milliliter of this bacterial suspension was centrifuged at 2,000 × g for 10 min at room temperature, and 20 μl of the bacterial supernatant was spotted onto the potato tuber disk prepared as described in Volksch et al. (47). The presence of coronatine was characterized by a hypertrophic response (an obvious enlargement of tissue) on the potato disks.

Ice nucleation activity.

P. syringae strains were grown on KB plates at room temperature for 4 to 5 days. A single colony of P. syringae was suspended in 100 μl of potassium phosphate buffer (10 mM, pH 7) (PPB) by gentle vortexing. Then, 10 μl of this suspension was added to 2 ml of PPB prechilled in a −10°C ethanol-ice water bath for 5 min. Strains were scored positive for ice nucleation activity if there was immediate ice formation in the tube.

Copper resistance.

Copper resistance was determined by using the method of Cazorla et al. (10). First, 50 μl of a P. syringae culture (OD600 = 0.5) was mixed with 50 μl of mannitol-glutamic-acid yeast extract medium (34) containing CuSO4 at the following final concentrations: 0, 0.5, 0.8, 1.0, 1.5, 2.0, 3.0, or 3.5 mM in 96-well microtiter plates (10). The plates were shaken at 30°C, and the OD600 of the mixture was taken immediately after inoculation and at 48 h postinoculation using a Tecan GENios microplate reader. Bacterial growth from the two time points was compared, and the MIC was determined. MIC is defined as the point where the OD of the bacterial culture at 48 h was the same or less than it was at 0 h. Strains with MICs of ≤0.8 mM CuSO4 were scored as copper sensitive (10).


P. syringae strains were streaked onto KB plates containing ampicillin (100 μg/ml), chloramphenicol (25 μg/ml), kanamycin (50 μg/ml), rifampin (50 μg/ml), streptomycin (100 μg/ml), or tetracycline (15 μg/ml). Bacterial growth was checked after 24 h and 48 h.

Statistical analyses.

We tested for associations between the MLST data and functional data by using an analysis of molecular variance (AMOVA) (17) as implemented in Arlequin, version 2.0 (41). The analysis determines how the genetic variation found among housekeeping genes is partitioned within and among populations. Populations are defined as strains that were either positive or negative for a specific phenotype (e.g., one population would be strains that produced coronatine, whereas the other population would be strains that did not). Pairwise distances were computed by using the Tamura and Nei distance measure with a gamma correction of 0.18. One thousand permutations of the data were used to create the null distribution. Other statistical tests were performed with StatView version 5.0.1 (SAS Institute).



The original MLST analysis of P. syringae (40) was performed with seven housekeeping genes on 60 strains. The current analysis expands on the number of strains to 95, but reduces the number of loci examined, and refines the PCR and sequencing primers. We were able to reduce the number of loci typed because of the extremely high level of phylogenetic congruence among the original set (40). This reduction dramatically increased the throughput of the analysis with almost no loss of phylogenetic resolution.
All of the basic population genetic and phylogenetic analyses of the current data set are in agreement with the original analyses of Sarkar and Guttman (40). Four primary clades of P. syringae were identified in both analyses (Fig. 1). Group 1 is largely composed of pathogens of tomatoes and brassicaceous crops (pathovars tomato and maculicola, respectively). Group 2 shows the greatest host diversity, and is the home to pea pathogens (pathovar pisi), and most of the pathovar syringae strains. Group 3 holds most of the bean (pathovars glycinea and phaseolicola) and cucumber (pathovar lachrymans) pathogens. Finally, group four strictly contains monocot pathogens. The original analysis also identified two identical radish pathogens (pathovar maculicola) that diverged from the rest of the P. syringae strains early on. We have since identified two additional strains that cluster with this group (only one shown in this analysis) and are now referring to this clade as group 5. The one group 5 strain not shown in the present study is the radish pathogen P. syringae pv. maculicola M4 (Pma M4), which has been used in a number of important studies (13, 31, 38, 39). The strain originally described as Pma M4 in Sarkar and Guttman (40) was misidentified prior to receipt by the DSG laboratory and has now been renamed Pma M4a.


Tabtoxin is a monocyclic β-lactam in which the dipeptide toxin is linked by a peptide bond to threonine (5). It produces chlorosis in the host plant cell after cleavage of the peptide bond, which releases the toxic tabtoxinine-β-lactam moiety (TβL) (5). TβL is believed to inhibit glutamine synthesis or the detoxification of ammonia by irreversibly inhibiting glutamine synthetase. Tabtoxin is historically associated with pathovars tabaci (tobacco), coronafaciens (oats), and garcae (coffee) (5).
Escherichia coli N99 was used as an indicator strain for tabtoxin production. The production of tabtoxin by a P. syringae colony resulted in a zone of inhibition around the colony due to the localized killing of the surrounding E. coli lawn. The addition of exogenous glutamine to the assay plates suppressed tabtoxin-mediated toxicity to E. coli.
Four strains in our collection tested positive for tabtoxin production (Fig. 1). Only one of two pathogens of both tobacco and oats were tabtoxin positive. The other positive strains were both group 1 tomato pathogens. The acquisition of tabtoxin appears to have occurred independently in the two genetically divergent tomato pathogens since all other members of this clade lack the phenotype.


Phaseolotoxin is a chlorosis-inducing phytotoxin, which has largely been found in strains causing disease in beans (pathovar phaseolicola) and kiwi (pathovar actinidiae) (5). This toxin inhibits ornithine carbamoyltransferase, a central enzyme in the urea cycle, thereby resulting in arginine deficiency (5).
E. coli strain N99 is sensitive to phaseolotoxin, and was used as an indicator of phaseolotoxin production. Phaseolotoxin-positive P. syringae colonies grown on a lawn of E. coli N99 produced a zone of inhibition around the colony. Of the 95 strains assayed, only 5 were phaseolotoxin positive (Fig. 1). Surprisingly, only three of eight phaseolicola strains in our collection produced phaseolotoxin (Pph KN86, NS368, and Y5_2). Notably, strains Pph NS368 and Pph Y5_2 have MLST haplotypes identical to that of strain Pph 1448A, which is currently being sequenced by The Institute for Genomic Research, although Pph 1448A was not included in the present study. The single pathovar actinidiae strain in our study collection (Pan FTRS_L1) did not produce phaseolotoxin. The only other two phaseolotoxin-positive strains were the Chinese cabbage pathogen P. syringae pv. maculicola H7608 and the tomato pathogen P. syringae pv. tomato KN10. Phaseolotoxin production could not be determined in P. syringae pv. glycinea KN44 since this strain could not grow on the minimal media, perhaps due to an auxotrophic mutation.


Syringomycin is a member of the cyclic lipodepsipeptide class of phytotoxins, which induce necrosis by forming pores in the plasma membrane of the host plant cell (5). The secretion of syringomycin promotes passive transmembrane influx of H+ and Ca2+ ions, acidifying the cytoplasm, resulting in cell death and the induction of a calcium related cellular signaling cascade (5, 9).
The basidiomycete yeast R. pilimanae was used as an indicator for syringomycin production. P. syringae strains were grown on a lawn of R. pilimanae. Strains that produced syringomycin produced a zone of inhibition around their colonies due to the antifungal activity of the toxin.
Syringomycins have classically been found in syringae pathovars (5). Twenty-one (22%) of our strains produced syringomycin, with a disproportionate number being in pathovar syringae strains (9 of 14 [64.3%], Fig. 1). Group 2 strains were strongly correlated with syringomycin production, with 76.2% of syringomycin-positive strains in this clade (P < 0.001 [Fisher exact test]). Within group 2, two independent clades of pathovar pisi strains (pea pathogens) have lost the ability to produce syringomycin. The only pea isolate that produced syringomycin is a pathovar syringae strain (Psy 1212R) that does not cluster with the pathovar pisi strains. An AMOVA analysis of syringomycin reveals that 89.74% of core genome genetic variation is found within populations (syringomycin producers versus nonproducers), while 10.26% of variation is found among populations (see supplementary Table S1).


Coronatine is a polyketide phytotoxin secreted by P. syringae, which induces chlorosis and lesions in host cells (5). The structure of coronatine mimics that of methyl jasmonate, an important growth regulator and signaling molecule that is synthesized by plants under biological stress (4). Coronatine production was found to be associated with the induction of 50 jasmonate and wound responsive genes and the suppression of pathogenesis-related genes during P. syringae infection of A. thaliana (49), indicating an important role for coronatine in plant virulence. The coronatine synthetic genes are commonly plasmid localized (6, 7). We assayed for coronatine production by scoring for a hypertrophic response or rotting of potato disks (48).
Coronatine production has been documented in pathogens of ryegrass (pv. atropurpurea), soybean (pv. glycinea), stone fruit (pv. morsprunorum), and tomato (pv. tomato) (7). Fourteen (14.7%) of our strains produced coronatine, with production predominantly found in tomato pathogens (7 of 11 tomato strains tested positive, Fig. 1). All of the other coronatine-positive strains were pathogens of either brassicaceous crops (radish or cabbage) or beans (kidney or soybean). Surprisingly, one of the strains most intensively used in coronatine studies (Pto PT23) did not test positive for coronatine production (discussed below). The AMOVA analysis found 81.72% of genetic variation was found within populations, while 18.28% was distributed among populations (see supplementary Table S1).

Ice nucleation activity.

Water on leaf surfaces typically supercools to temperatures below −5°C before forming ice nuclei and freezing. The P. syringae gene ina produces a protein (INA) that acts as a heterogeneous nuclei for ice crystal formation, raising the temperature of ice formation to as high as −1.2°C, thereby causing increased frost damage (32). The INA protein has a repeated octapeptide motif highly conserved in at least four diverse bacterial species known to carry ina orthologs (16). This protein forms aggregates that associate with the bacterial outer membrane, mimicking the structure of an ice crystal (16).
We surveyed for ice nucleation activity by monitoring the ability of bacteria to freeze supercooled buffer. A total of 80% (76 of 95 strains) of our strains produced the ice nucleation phenotype (Fig. 1). All of our tomato pathogens tested positive for ice nucleation activity, although historically pathovar tomato strains have not been known to exhibit this phenotype (23). All of our cucumber pathogens (pv. lachrymans) were also INA positive. Interestingly, all of the soybean and pea pathogens were INA positive, while there was a slight, yet statistically significant, negative association for ice nucleation activity in kidney bean pathogens (pv. phaseolicola, P = 0.025 [Fisher exact test]). This deficiency in ice nucleation activity among phaseolicola strains was most apparent in the very closely related phaseolicola clade found in group 3, with six of eight strains being INA negative (P = 0.002). Finally, pathogens of brassicaceous crops were also negatively associated with INA activity (P = 0.007). All group four strains and all but one strain of the group 2 strains have ice-nucleating activity. As expected, an AMOVA finds that the vast majority of genetic variation (95.60%) is found within rather than among populations defined by the presence or absence of INA activity (see supplementary Table S1).

Copper resistance.

Copper sulfate has been used as a potent bactericide for the control of phytopathogenic P. syringae for more than a century (8). Recently, several P. syringae copper resistant determinants have been identified and characterized. One of the determinants is the plasmid-encoded cop operon (11), which encodes membrane and periplasmic proteins that are believed to sequester and compartmentalize copper in the periplasmic space and outer membrane of the bacteria (11).
We used the MIC (MIC) of copper to quantified resistance. Those strains with MICs of greater than 0.8 mM CuSO4 were considered resistant as per Cazorla et al. (10). 75% of our strains (72 out of 95) were resistant to copper (Fig. 1). The sensitive strains were scattered throughout the MLST tree, although a slight excess of sensitive strains was found among pathovar phaseolicola bean pathogens (P = 0.018), particularly within the closely related group 3 phaseolicola clade (P = 0.005), where only two of the eight pathovar phaseolicola strains were resistant. This is the same clade that was deficient in ice nucleation activity. An AMOVA analysis indicates that 97.14% of genetic variation is found within populations (see supplementary Table S1).


Six different antimicrobial agents were used in the present study: (i) ampicillin, a β-lactam, which inactivates penicillin-binding proteins, thereby inhibiting cell wall biosynthesis; (ii) rifampin, which targets the β subunit of RNA polymerase II, thereby inhibiting transcription initiation; (iii) chloramphenicol, which binds to the 70S ribosome and inhibits the peptidyl transferase reaction during translation; (iv) kanamycin, which inhibits protein synthesis by targeting the 30S ribosome; (v) streptomycin, which also inhibits protein synthesis by inactivating the 30S ribosome; and (vi) tetracycline, which inhibits chain elongation during protein synthesis by blocking aminoacyl tRNA binding at the A site (12, 35).
Of our entire MLST-typed strain collection, only one strain, P. syringae pv. syringae NCPPB281 (PsyNCPPB28, alternatively ATCC 19310 ), was resistant to kanamycin and tetracycline (Fig. 1). This strain is a weak pathogen of lilacs and has been used in a number of published studies. It is possible that the atypical resistance pattern seen in this strain is due to the genetic modification of the particular isolate provided to the DSG laboratory. Streptomycin resistance was only found in eight strains. Four of these isolates are cucumber pathogens (pathovar lachrymans) that form a tight clade in group 3. Rifampin resistance, which is readily selected for in the laboratory, was found in 16.8% of strains (16 of 95), while 37.9% of strains (36 of 95) were resistant to chloramphenicol. These two resistance phenotypes are scattered throughout the tree with no apparent phylogenetic or host-specific bias. A total of 57.9% (55 of 95) of strains are resistant to ampicillin. Most of the ampicillin-sensitive strains are found in group 1, particularly the cabbage, cauliflower, and radish pathogens (pv. maculicola). All eight group 1 maculicola strains are sensitive to ampicillin, while one of the two group 5 maculicola strains is ampicillin sensitive. The sole remaining maculicola strain in the collection, which is found in group 3, is resistant to ampicillin. All three of the resistant maculicola strains are radish pathogens.
It is notable that there are strains that are identical by our MLST typing but which differ in their antibiotic resistance profiles. For example, strain Pph R6a, which is resistant to both rifampin and chloramphenicol, and strain Pph SG44, which is sensitive to both antibiotics, are identical at all of the MLST loci.


There has been a tremendous explosion of interest in the role played by virulence-associated molecules in P. syringae host interactions. Most of the current studies focus on the widely conserved type III secretion system and its effector proteins. Nevertheless, strains of P. syringae also produce an impressive array of toxins, which typically act in a non-host-specific manner. Although toxins are not required for pathogenesis, they have been found to enhance virulence by increasing the severity of lesions and by contributing to increased growth and movement of bacteria inside the plant tissue (5). Pseudomonads in general are also widely recognized as being highly resistant to a broad range of medically and agriculturally important antimicrobial compounds. P. syringae is no exception to this rule.
To date, no study of toxin production or antimicrobial resistance in P. syringae has used a precise phylogenetic framework. We have used an MLST approach to characterize the P. syringae core genome, which provides the most accurate reflection of the clonal evolutionary history of the species. By mapping the distribution of toxin production and antimicrobial resistance onto the MLST phylogeny, we can more precisely determine the evolutionary origin of these phenotypes.
Four conclusions emerge from our study of toxin production. First, toxin production is surprisingly rare in P. syringae. Of 95 strains assayed, at least 54 (56.8%) did not produce any of the four toxins tested. None of the six cucumber pathogens or three wheat pathogens produced any toxins. Only one of the eight pea pathogens produced a toxin, with the sole exception being a pea isolate that was not originally given a pathovar pisi designation (Psy 1212R, which produced syringomycin).
The conclusion that toxin production is relatively rare assumes (i) that our assays are robust in their ability to identify the four toxins in all strains, (ii) that strains do not lose their ability to produce toxins when stored in the laboratory, and (iii) that there are no other significant toxins produced by this species. We do not believe the first issue is significant since we used standard toxin assays that have been used widely in other studies. The second point is very important. Some strains of P. syringae have been known to lose toxin production when stored for long periods. These phenotypes can sometimes be recovered by passaging the strains in planta prior to the toxin assays. Unfortunately, this procedure was impossible in the present study given the very large diversity of hosts. We do not believe this issue is a significant problem since our date is consistent with published results in all cases except the loss of coronatine production in strain Pto Pt23. With respect to the last issue, it is very unlikely that there are significant numbers of undescribed toxins given their extensive study in P. syringae. Furthermore, most of the assays performed in the present study were of limited specificity, identifying any representative of a class of toxins. For example, the syringomycin assay would detect any lipodepsipeptide toxin. Given the lack of specificity, these assays should provide a conservative estimate of the frequency of toxin production.
The second conclusion comes from the observation of a surprising negative association among toxin. P. syringae strains are very unlikely to produce more than one toxin. Only 2 of the 95 strains produced two toxins, and only 1 strain produced three toxins (the tomato pathogen Pto KN10, which produced tabtoxin, phaseolotoxin, and coronatine). Of the 21 strains producing syringomycin, only one strain produced coronatine, and none produced phaseolotoxin. Of the 14 strains producing coronatine, only 1 produced phaseolotoxin, while 1 produced syringomycin. Of the six strains producing phaseolotoxin, only one produced coronatine, while none produced syringomycin. These negative associations are statistically significant as determined by Fisher exact test (P < 0.0001 [syringomycin-coronatine]; P < 0.0001 [syringomycin-phaseolotoxin]; and P = 0.003 [phaseolotoxin-coronatine]). In addition, three of the four tabtoxin-producing strains did not produce any other toxins. Is this negative association due to a cost associated with the production of multiple toxins or simply the by-product of a relatively low rate of toxin production in P. syringae? Based on our empirical determination of the frequency of each toxin, at least 7 strains of a collection of 95 should produce two or more toxins. Our observation of only three multiple-toxin-producing strains indicates that these strains are less frequent in the population than they should be if toxins were produced independently of each other. This suggests a cost to the production of multiple toxins in P. syringae.
Our third conclusion is that toxin production is very poorly associated with the host of isolation (Fig. 1). The best evidence for host association comes from a cluster of tomato pathogens that produce coronatine. However, this case is just as readily explained by nonindependent evolutionary histories. Furthermore, only 7 of the 11 tomato pathogens produced coronatine. It is perhaps easier to make the case for a correlation between the lack of production of a particular toxin and a specific host. This appears to be the case with respect to the two divergent clades of pea pathogens (pv. pisi) in group 2. As discussed above, both of these clades lack syringomycin production, while this toxin is relatively common throughout the rest of the group 2 strains.
Finally, we conclude that toxin production is generally distributed in a manner inconsistent with clonal (vertical) evolution (Fig. 1); therefore, the evolution of toxin producing genes is very likely driven by horizontal gene transfer. This conclusion is also supported by similarity analysis of the genes and proteins responsible for toxin production. For example, the Cma proteins, which are necessary for coronatine biosynthesis, are similar to proteins distributed throughout the bacterial and eukaryota domains, including other pseudomonads. However, the only similar nucleotide sequence found outside the highly conserved orthologs in the P. syringae complex is in Burkholderia pseudomallei (E value = 2e-37, NCBI discontiguous megablast). Either the P. syringae cma genes were acquired vertically and diverged to such an extent that they no longer show any nucleotide similarity to homologs in other pseudomonads or, more likely, they were acquired horizontally from an as-yet-unsequenced organism.
The tabtoxin and phaseolotoxin biosynthetic genes have no homologs outside of the P. syringae complex. The small tabtoxin biosynthetic cluster is sufficient for tabtoxin synthesis and was observed by Kinscherf et al. (28) to excise from the chromosome at frequencies as high as 10−3/CFU. The specific mechanism driving this process is not known.
The interpretation of the history of the syringomycin biosynthetic genes is less clear. Syringomycin production is heavily concentrated in group 2. It appears that a common ancestor of this group may have acquired the syringomycin regulon, and it has since passed vertically to the descendants of this ancient bacterium. In fact, syringomycin is the only toxin produced by the group 2 strains. Nevertheless, during the evolution of this group, the syringomycin system appears to have been lost or disabled in a substantial number of strains, most notably, the ancestor of the clade of six pathovar pisi strains. Is syringomycin production ancestral in the P. syringae complex? The most parsimonious answer to this question is no, since syringomycin production is very rare in the other four syringae groups. This is further supported by the fact that three of these four groups branched off basal to group 2.
This syringomycin biosynthesis operon is dominated by the enormous syrE gene, which is 28.1 kb in length, encoding a protein of 1,039 kDa, making it the largest prokaryotic protein discovered (20). The SyrE synthetase protein contains eight conserved modules that show high similarity (typically ca. 75% nucleotide identity, but often as high as 90%) to homologous modules in a wide range of related gram-negative bacteria and even in species as distant as high-GC gram-positive bacteria such as Streptomyces spp. Given the common occurrence of syringomycin production in the group 2 strains and the wide distribution of the biosynthetic genes throughout the bacterial domain, further comparative analysis would have to be completed before we can conclusively determine whether the distribution of this gene cluster is due to horizontal genetic exchange or simply rampant gene loss after vertical descent.
Unlike toxin production, the ability to nucleate ice and detoxify copper appears to be an ancestral trait in P. syringae (Fig. 1). With 80% of strains being able to nucleate ice, and 75% of strains being copper resistant, perhaps the more interesting observations come from those strains that have lost this ability. Although all soybean and pea pathogens are INA positive, there was a negative association between INA activity and the bean pathogens. Alternatively, this pattern may simply be the result of nonindependent evolutionary histories, a circumstance analogous to the situation in the tomato pathogens with respect to coronatine production. One way to answer this question is to determine whether the three strains that are INA positive in the closely related bean pathogen clade gained (or retained) the phenotype independently. A much stronger case can be made for pathogens of brassicaceous crops, which are also negatively associated with INA activity. These strains are scattered throughout groups 1 and 5. It would be much more difficult to invoke nonindependent evolutionary histories as the cause for the correlation. No clear phylogenetic or host-specific pattern was seen with copper resistance, except for the observation that the group 3 bean pathogen clade that was deficient in its ability to nucleate ice nucleation also had an excess of copper-sensitive strains. The meaning of the correlation is unclear.
Resistance to ampicillin appears to be ancestral in P. syringae, although the relatively high rate of ampicillin sensitivity in groups 1 and 5 (39 and 33% resistant, respectively, versus 70% resistance among the three remaining groups) presents the possibility that ampicillin resistance was lost in the ancestral lineage that gave rise to groups 1 and 5 or acquired in the ancestral lineage that gave rise to groups 2, 3, and 4. It is impossible to judge whether one hypothesis is more valid than the other based on the current data, since the phylogenetic support for these basal branches is very weak. Resistance to the other five antibiotics is generally quite rate, and there is absolutely no association between antibiotic resistance, the core genome phylogeny, or host of isolation. These findings are not surprising given the well-known propensity of antibiotic resistance genes to be transferred horizontally.
The interaction between a pathogen and its host is complex and multifaceted. To date, only type III effectors have been conclusively shown to both qualitatively limit and enable pathogenesis on specific hosts (1, 26, 45). We are unable to identify a similar role for toxins in the present study. No specific host association was found to be strictly associated with the production of a particular toxin or resistance to a specific antimicrobial agent. Despite the reputation of P. syringae as a copious toxin producer and the well-established role toxins play in modulating virulence, the present study clearly shows that no toxin is common or definitively ancestral in this species.
FIG.1. Phylogenetic tree and distribution of toxin and resistance phenotypes. A linearized neighbor-joining MLST tree from combined rpoD, gyrB, gltA, and gapA data set, constructed using the K2P-γ (α = 0.2) substitution model, is shown. Numbers above the nodes are bootstrap scores based on 1,000 pseudoreplicates. The genetic distance scale is presented below the tree. The tree is identical in its gross topology to one produced by maximum likelihood. Strain designations are presented on the right, along with the host of isolation. The black and white grid represents the presence or absence, respectively, of the assayed phenotypes. Tab, tabtoxin; Phas, phaseolotoxin; Cor, coronatine; Syr, syringomycin; INA, ice nucleation; Kan, kanamycin; Tet, tetracycline; Str, streptomycin; Rif, rifampin; Chl, chloramphenicol; Amp, ampicillin; Cu, copper.
TABLE 1. Strains
PathovarDesignationStrain namePlace of isolationbYr of isolationHostSourceaAccession no.
acerisATCC10853Pac A10853  MapleJ. DanglATCC 10853
actinidiaeFTRS_L1Pan FTRS_L1Japan1984KiwiMAFFMAFF302091
aesculi0893_23Pae 0893_23US Horse chestnutD. Cooksey 
apii1089_5Pap 1089_5US CeleryD. Cooksey 
aptata601Ptt 601 1966Sugar beetMAFFMAFF301008
aptataDSM50252Ptt DSM5022  WheatJ. Dangl 
aptataG733Ptt G733 1976Brown riceMAFFMAFF302831
atrofaciensDSM50255Paf DSM5025  WheatJ. Dangl 
broussonetiaeKOZ8101Pbr KOZ8101Japan1980Paper mulberryMAFFMAFF810036
cilantro0788_9Pci 0788_9US CilantroD. Cooksey 
coronafaciens3113Pcn 3113UK1958OatsD. ArnoldICMP3113
coronafaciensKN221Pcn KN221 1984OatsMAFFMAFF302787
glycineaBR1Pgy BR1 1989SoybeanMAFFMAFF210373
glycineaKN127Pgy KN127 1982SoybeanMAFFMAFF302751
glycineaKN166Pgy KN166 1982SoybeanMAFFMAFF302770
glycineaKN28Pgy KN28 1981SoybeanMAFFMAFF302676
glycineaKN44Pgy KN44Japan1981SoybeanMAFFMAFF301683
glycineaLN10Pgy LN10 1989SoybeanMAFFMAFF210389
glycineaMAFF301765Pgy M301765 1982SoybeanMAFFMAFF301765
glycineaMOC601Pgy MOC601 1994SoybeanMAFFMAFF311113
glycineaR4aPgy R4a  SoybeanJ. Dangl 
glycineaUnB647Pgy UnB647  Kidney beanMAFFMAFF210405
japonicaMAFF301072Pja M301072Japan1951BarleyMAFFMAFF301072
lachrymans107Pla 107  CucumberJ. DanglMAFF301315
lachrymans3988Pla 3988US1935CucumberD. ArnoldICMP3988
lachrymans1188_1Pla 1188_1US ZucchiniD. Cooksey 
lachrymansATCC7386Pla A7386  CucumberJ. DanglMAFF302278
lachrymansN7512Pla N7512Japan1975CucumberMAFFMAFF301315
lachrymansYM7902Pla YM7902 1979CucumberMAFFMAFF730057
lachrymansYM8003Pla YM8003 1980CucumberMAFFMAFF730069
maculicola4981Pma 4981Zimbabwe CauliflowerD. Cuppels 
maculicolaAZ85297Pma AZ85297 1985Chinese cabbageMAFFMAFF302539
maculicolaES4326Pma ES4326 1965RadishJ. T. Greenberg 
maculicolaH7311Pma H7311Japan1973Chinese cabbageMAFFMAFF301174
maculicolaH7608Pma H7608 1976Chinese cabbageMAFFMAFF301175
maculicolaKN203Pma KN203 1983Chinese cabbageMAFFMAFF302783
maculicolaKN84Pma KN84 1982RadishMAFFMAFF302724
maculicolaKN91Pma KN91 1982RadishMAFFMAFF302731
maculicolaM4aPma M4a  RadishJ. Dangl 
maculicolaM6Pma M6UK1965CauliflowerJ. Dangl 
maculicolaYM7930Pma YM7930 1979RadishMAFFMAFF301419
melleaN6801Pme N6801 1968TobaccoMAFFMAFF302303
moriMAFF301020Pmo M301020Japan1966MulberryMAFFMAFF301020
morsprunorum19322Pmp 19322  European plumJ. DanglATCC 19322
morsprunorumFTRS_U7805Pmp FTRS_U7Japan1978Japanese apricotMAFFMAFF301436
myricaeAZ84488Pmy AZ84488 1984BayberryMAFFMAFF302460
myricaeMAFF302941Pmy M302941 1989BayberryMAFFMAFF302941
oryzae36_1Por 36_1 1983RiceMAFFMAFF301538
oryzae1_6Por 1_6 1991RiceMAFFMAFF311107
phaseolicola1302APph 1302AEthiopia1984Kidney beanD. Arnold 
phaseolicola1449BPph 1449BEthiopia1985Hyacinth beanD. Arnold 
phaseolicolaHB10YPph HB10Y  Snap beanP. TurnerATCC 21781
phaseolicolaKN86Pph KN86Japan1982Kidney beanMAFFMAFF301673
phaseolicolaNPS3121Pph NPS3121  Kidney beanJ. T. Greenberg 
phaseolicolaNS368Pph NS368 1992Kidney beanMAFFMAFF311004
phaseolicolaR6aPph R6a  Kidney beanJ. Dangl 
phaseolicolaSG44Pph SG44US1980Snap beanS. Hirano 
phaseolicolaY5_2Pph Y5_2  KudzuMAFFMAFF311162
pisi895APpi 895A  PeaD. Arnold 
pisiH5E1Ppi H5E1 1993PeaMAFFMAFF311141
pisiH5E3Ppi H5E3 1993PeaMAFFMAFF311143
pisiH6E5Ppi H6E5 1994PeaMAFFMAFF311144
pisiH7E7Ppi H7E7 1995PeaMAFFMAFF311146
pisiPP1Ppi PP1Japan1978PeaMAFFMAFF301208
pisiR6aPpi R6a  PeaJ. Dangl 
savastanoi4352Psv 4352Yugoslavia OliveA. Colmer 
sesamiHC_1Pse HC_1  SesameMAFFMAFF311181
syringae1212RPsy 1212R  PeaD. Arnold 
syringaeA2Psy A2  Ornamental pearC. Bender 
syringaeB48Psy B48US PeachT. Denny 
syringaeB64Psy B64US WheatT. Denny 
syringaeB728APsy B728AUS Snap beanS. Hirano 
syringaeB76Psy B76US TomatoT. Denny 
syringaeFF5Psy FF5US1998Ornamental pearC. Bender 
syringaeFTRS_W6601Psy FTRS_W6 1966Japanese apricotMAFFMAFF301429
syringaeFTRS_W7835Psy FTRS_W7 1978Japanese apricotMAFFMAFF301430
syringaeL177Psy L177 1983LilacMAFFMAFF302085
syringaeLOB2_1Psy LOB2_1Japan1986LilacMAFFMAFF301861
syringaeNCPPB281Psy NCPPB28UK LilacB. HancockATCC 19310
syringaePs9220Psy Ps9220 1992Spring onionMAFFMAFF730125
syringaePSC1BPsy PSC1BUS CornT. Denny 
tabaci6606Pta 6606Japan1967TobaccoMAFFMAFF301612
theaK93001Pth K93001 1993TeaMAFFMAFF302851
tomato487Pto 487Greece TomatoD. Cuppels 
tomato1318Pto 1318Switzerland TomatoD. Cuppels 
tomato2170Pto 2170 1984TomatoMAFFMAFF301591
tomatoDC3000Pto DC3000UK TomatoJ. T. Greenberg 
tomatoDC84_1Pto DC84_1Canada TomatoD. Cuppels 
tomatoDC89_4HPto DC89_4HCanada TomatoD. Cuppels 
tomatoDCT6D1Pto DCT6D1Canada TomatoD. Cuppels 
tomatoKN10Pto KN10 1981TomatoMAFFMAFF302665
tomatoPT23Pto PT23US1986TomatoN. T. Keen 
tomatoTF1Pto TF1US1997TomatoS. Hirano 
cCit7Ps Cit7  Navel orangeS. Lindow 
cTLP2Ps TLP2  PotatoS. Lindow 
MAFF, Japanese Ministry of Agriculture, Forestry, and Fisheries; ATCC, American Type Culture Collection; ICMP, International Collection of Micro-organisms from Plants (New Zealand).
US, United States; UK, United Kingdom.
—, no pathovar designation.
TABLE 2. MLST primers
PrimeraTm (°C)bLength (bp)Sequence
Forward-strand primers (+), reverse-strand primers (−), PCR primers (p), and sequencing primers (s) are as indicated.
Tm, Melting temperature.


D.S.G. is a Canada Research Chair in Comparative Genomics and is supported by grants from the Natural Sciences and Engineering Research Council of Canada, the Canadian Foundation for Innovation, and Performance Plants, Inc., of Kingston, Ontario.
The D.S.G. laboratory is deeply indebted to those individuals who graciously provided strains.


Abramovitch, R. B., Y. J. Kim, S. Chen, M. B. Dickman, and G. B. Martin. 2003. Pseudomonas type III effector AvrPtoB induces plant disease susceptibility by inhibition of host programmed cell death. EMBO J.22:60-69.
Archer, G. P., C. J. Kennedy, and M. J. W. Povey. 1996. Investigations of ice nucleation in water-in-oil emulsions using ultrasound velocity measurements. Cryo-Letters17:391-396.
Atlas, R. M., and L. C. Parks. 1993. Handbook of microbiological media. CRC Press, Inc., Boca Raton, Fla.
Bender, C., D. Palmer, A. PenalozaVazquez, V. Rangaswamy, and M. Ullrich. 1996. Biosynthesis of coronatine, a thermoregulated phytotoxin produced by the phytopathogen Pseudomonas syringae. Arch. Microbiol.166:71-75.
Bender, C. L., F. Alarcon-Chaidez, and D. C. Gross. 1999. Pseudomonas syringae phytotoxins: mode of action, regulation, and biosynthesis by peptide and polyketide synthetases. Microbiol. Mol. Biol. Rev.63:266-292.
Bender, C. L., D. K. Malvick, and R. E. Mitchell. 1989. Plasmid-mediated production of the phytotoxin coronatine in Pseudomonas syringae pv. tomato. J. Bacteriol.171:807-812.
Bender, C. L., S. A. Young, and R. E. Mitchell. 1991. Conservation of plasmid DNA sequences in coronatine-producing pathovars of Pseudomonas syringae. Appl. Environ. Microbiol.57:993-999.
Bruins, M. R., S. Kapil, and F. W. Oehme. 2000. Microbial resistance to metals in the environment. Ecotoxicol. Environ. Safety45:198-207.
Bultreys, A., and I. Gheysen. 1999. Biological and molecular detection of toxic lipodepsipeptide-producing Pseudomonas syringae strains and PCR identification in plants. Appl. Environ. Microbiol.65:1904-1909.
Cazorla, F. M., E. Arrebola, A. Sesma, A. Perez-Garcia, J. C. Codina, J. Murillo, and A. de Vicente. 2002. Copper resistance in Pseudomonas syringae strains isolated from mango is encoded mainly by plasmids. Phytopathology92:909-916.
Cooksey, D. A. 1994. Molecular mechanisms of copper resistance and accumulation in bacteria. FEMS Microbiol. Rev.14:381-386.
Davies, J., and G. D. Wright. 1997. Bacterial resistance to aminoglycoside antibiotics. Trends Microbiol.5:234-240.
Debener, T., H. Lehnackers, M. Arnold, and J. L. Dangl. 1991. Identification and molecular mapping of a single Arabidopsis thaliana locus determining resistance to a phytopathogenic Pseudomonas syringae isolate. Plant J.1:289-302.
Durbin, R. D. 1982. Toxins and pathogenesis, p. 423-441. In M. S. Mount and G. H. Lacy (ed.), Phytopathogenic prokaryotes,vol. 1. Academic Press, Inc., New York, N.Y.
Dye, D. W. 1953. Control of Pseudomonas syringae with streptomycin. Nature172:683-684.
Edwards, A. R., R. A. Vandenbussche, H. A. Wichman, and C. S. Orser. 1994. Unusual pattern of bacterial ice nucleation gene evolution. Mol. Biol. Evol.11:911-920.
Excoffier, L., P. E. Smouse, and J. M. Quattro. 1992. Analysis of molecular variance inferred from metric distances among DNA daplotypes: application to human mitochondrial-DNA restriction data. Genetics131:479-491.
Felsenstein, J. 1993. PHYLIP (Phylogeny Inference Package), 3.5c ed. Department of Genetics, University of Washington, Seattle.
Gasson, M. J. 1980. Indicator technique for antimetabolic toxin production by phytopathogenic species of Pseudomonas. Appl. Environ. Microbiol.39:25-29.
Guenzi, E., G. Galli, I. Grgurina, D. C. Gross, and G. Grandi. 1998. Characterization of the syringomycin synthetase gene cluster: a link between prokaryotic and eukaryotic peptide synthetases. J. Biol. Chem.273:32857-32863.
Guttman, D. S., and J. T. Greenberg. 2001. Functional analysis of the type III effectors AvrRpt2 and AvrRpm1 of Pseudomonas syringae with the use of a single-copy genomic integration system. Mol. Plant-Microbe Interact.14:145-155.
Hacker, J., and E. Carniel. 2001. Ecological fitness, genomic islands and bacterial pathogenicity: a Darwinian view of the evolution of microbes. EMBO Rep.2:376-381.
Hirano, S. S., and C. D. Upper. 2000. Bacteria in the leaf ecosystem with emphasis on Pseudomonas syringae: a pathogen, ice nucleus, and epiphyte. Microbiol. Mol. Biol. Rev.64:624-653.
Hoitink, H. A. J., and S. L. Sinden. 1970. Partial purification and properties of chlorosis inducing toxins of Pseudomonas phaseolicola and Pseudomonas glycinea. Phytopathology60:1236-1238.
Horst, R. K. 1990. Westcott's plant disease handbook, 5th ed. Chapman & Hall, New York, N.Y.
Jackson, R. W., E. Athanassopoulos, G. Tsiamis, J. W. Mansfield, A. Sesma, D. L. Arnold, M. J. Gibbon, J. Murillo, J. D. Taylor, and A. Vivian. 1999. Identification of a pathogenicity island, which contains genes for virulence and avirulence, on a large native plasmid in the bean pathogen Pseudomonas syringae pathovar phaseolicola. Proc. Natl. Acad. Sci. USA96:10875-10880.
King, E. O., M. K. Ward, and D. E. Raney. 1954. Two simple media for the demonstration of phycocyanin and fluorescin. J. Lab. Clin. Med.44:301-307.
Kinscherf, T. G., R. H. Coleman, T. M. Barta, and D. K. Willis. 1991. Cloning and expression of the tabtoxin biosynthetic region from Pseudomonas syringae. J. Bacteriol.173:4124-4132.
Kumar, S., K. Tamura, and M. Nei. 1994. MEGA: molecular evolutionary genetics analysis software for microcomputers. Comput. Appl. Biosci.10:189-191.
Leach, J. E., and F. F. White. 1996. Bacterial avirulence genes. Annu. Rev. Phytopathol.34:153-179.
Lim, M. T., and B. N. Kunkel. 2004. Mutations in the Pseudomonas syringae avrRpt2 gene that dissociate its virulence and avirulence activities lead to decreased efficiency in AvrRpt2-induced disappearance of RIN4. Mol. Plant-Microbe Interact.17:313-321.
Lindow, S. E. 1987. Competitive-exclusion of epiphytic bacteria by icePseudomonas syringae mutants. Appl. Environ. Microbiol.53:2520-2527.
Lydon, J., and C. D. Patterson. 2001. Detection of tabtoxin-producing strains of Pseudomonas syringae by PCR. Lett. Appl. Microbiol.32:166-170.
Mahapatra, N. R., and P. C. Banerjee. 1996. Extreme tolerance to cadmium and high resistance to copper, nickel and zinc in different Acidiphilium strains. Lett. Appl. Microbiol.23:393-397.
McDermott, P. F., R. D. Walker, and D. G. White. 2003. Antimicrobials: modes of action and mechanisms of resistance. Int. J. Toxicol.22:135-143.
Miller, J. H. 1972. Experiments in molecular genetics. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y.
Neu, H. C. 1992. The crisis in antibiotic resistance. Science257:1064-1073.
Ritter, C., and J. L. Dangl. 1996. Interference between two specific pathogen recognition events mediated by distinct plant disease resistance genes. Plant Cell8:251-257.
Rossier, O., K. Wengelnik, K. Hahn, and U. Bonas. 1999. The Xanthomonas Hrp type III system secretes proteins from plant and mammalian bacterial pathogens. Proc. Natl. Acad. Sci. USA96:9368-9373.
Sarkar, S. F., and D. S. Guttman. 2004. The evolution of the core genome of Pseudomonas syringae, a highly clonal, endemic plant pathogen. Appl. Environ. Microbiol.70:1999-2012.
Schneider, S., D. Roessli, and L. Excoffier. 2000. Arlequin: a software for population genetics data analysis, 2.0 ed. Genetics and Biometry Lab, Department of Anthropology, University of Geneva, Geneva, Switzerland.
Staskawicz, B. J., and N. J. Panopoulos. 1979. Rapid and sensitive microbiological assay for phaseolotoxin. Phytopathology69:663-666.
Swofford, D. L. 1993. PAUP: phylogenetic analysis using parsimony, 3.1 ed. Illinois Natural History Survey, Champaign, Ill.
Thompson, J. D., T. J. Gibson, F. Plewniak, F. Jeanmougin, and D. G. Higgins. 1997. The CLUSTAL_X windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res.25:4876-4882.
Tsiamis, G., J. W. Mansfield, R. Hockenhull, R. W. Jackson, A. Sesma, E. Athanassopoulos, M. A. Bennett, C. Stevens, A. Vivian, J. D. Taylor, and J. Murillo. 2000. Cultivar-specific avirulence and virulence functions assigned to avrPphF in Pseudomonas syringae pv. phaseolicola, the cause of bean halo-blight disease. EMBO J.19:3204-3214.
Turner, J. G., and R. R. Taha. 1984. Contribution of tabtoxin to the pathogenicity of Pseudomonas syringae pv. tabaci. Physiol. Plant Pathol.25:55-69.
Volksch, B., F. Bublitz, and W. Fritsche. 1989. Coronatine production by Pseudomonas syringae pathovars: screening method and capacity of product formation. J. Basic Microbiol.29:463-468.
Volksch, B., and H. Weingart. 1998. Toxin production by pathovars of Pseudomonas syringae and their antagonistic activities against epiphytic microorganisms. J. Basic Microbiol.38:135-145.
Zhao, Y., R. Thilmony, C. L. Bender, A. Schaller, S. Y. He, and G. A. Howe. 2003. Virulence systems of Pseudomonas syringae pv. tomato promote bacterial speck disease in tomato by targeting the jasmonate signaling pathway. Plant J.36:485-499.

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Published In

cover image Applied and Environmental Microbiology
Applied and Environmental Microbiology
Volume 71Number 9September 2005
Pages: 5182 - 5191
PubMed: 16151103


Received: 11 February 2005
Accepted: 29 March 2005
Published online: 1 September 2005


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Michael S. H. Hwang
Department of Botany, University of Toronto, Toronto, Ontario M5S 3B2, Canada
Robyn L. Morgan
Department of Botany, University of Toronto, Toronto, Ontario M5S 3B2, Canada
Sara F. Sarkar
Department of Botany, University of Toronto, Toronto, Ontario M5S 3B2, Canada
Pauline W. Wang
Department of Botany, University of Toronto, Toronto, Ontario M5S 3B2, Canada
David S. Guttman [email protected]
Department of Botany, University of Toronto, Toronto, Ontario M5S 3B2, Canada

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