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J Clin Microbiol. Jul 2012; 50(7): 2343–2352.
PMCID: PMC3405591
Biochemical, Serological, and Virulence Characterization of Clinical and Oyster Vibrio parahaemolyticus Isolates
Jessica L. Jones,corresponding authora Catharina H. M. Lüdeke,ab John C. Bowers,c Nancy Garrett,d Markus Fischer,b Michele B. Parsons,d Cheryl A. Bopp,d and Angelo DePaolaa
aFDA, Division of Seafood Science and Technology, Gulf Coast Seafood Laboratory, Dauphin Island, Alaska, USA
bUniversity of Hamburg, Department of Food Chemistry, Hamburg, Germany
cFDA, Center for Food Safety and Nutrition, Division of Mathematics, College Park, Maryland, USA
dCenters for Disease Control and Prevention, Atlanta, Georgia, USA
corresponding authorCorresponding author.
Address correspondence to Jessica L. Jones, Jessica.Jones/at/fda.hhs.gov.
Received January 18, 2012; Revisions requested March 12, 2012; Accepted April 12, 2012.
In this study, 77 clinical and 67 oyster Vibrio parahaemolyticus isolates from North America were examined for biochemical profiles, serotype, and the presence of potential virulence factors (tdh, trh, and type III secretion system [T3SS] genes). All isolates were positive for oxidase, indole, and glucose fermentation, consistent with previous reports. The isolates represented 35 different serotypes, 9 of which were shared by clinical and oyster isolates. Serotypes associated with pandemic strains (O1:KUT, O1:K25, O3:K6, and O4:K68) were observed for clinical isolates, and 7 (9%) oyster isolates belonged to serotype O1:KUT. Of the clinical isolates, 27% were negative for tdh and trh, while 45% contained both genes. Oyster isolates were preferentially selected for the presence of tdh and/or trh; 34% contained both genes, 42% had trh but not tdh, and 3% had tdh but not trh. All but 1 isolate (143/144) had at least three of the four T3SS1 genes examined. The isolates lacking both tdh and trh contained no T3SS2α or T3SS2β genes. All clinical isolates positive for tdh and negative for trh possessed all T3SS2α genes, and all isolates negative for tdh and positive for trh possessed all T3SS2β genes. The two oyster isolates containing tdh but not trh possessed all but the vopB2 gene of T3SS2α, as reported previously. In contrast to the findings of previous studies, all strains examined that were positive for both tdh and trh also carried T3SS2β genes. This report identifies the serotype as the most distinguishing feature between clinical and oyster isolates. Our findings raise concerns about the reliability of the tdh, trh, and T3SS genes as virulence markers and highlight the need for more-detailed pathogenicity investigations of V. parahaemolyticus.
Vibrio parahaemolyticus is the leading cause of seafood-associated illness in the United States and is generally associated with the consumption of raw molluscan shellfish (17). Even so, V. parahaemolyticus illnesses are believed to be highly underdiagnosed (143-fold), possibly due to the relatively low hospitalization (22.5%) and fatality (0.9%) rates (36). Another factor that may influence the underdiagnosis is the fact that traditional biochemical identification of V. parahaemolyticus has proven less reliable than molecular identification for clinical and food isolates (8, 21). When an isolate is confirmed as V. parahaemolyticus, virulence characterization is typically performed. The virulence of V. parahaemolyticus was initially attributed to the production of a thermostable direct hemolysin (TDH) (15, 26). The TDH is encoded by the tdh gene, and subsequently a related gene, termed the tdh-related hemolysin (trh), was identified (26). Together, tdh and trh are widely considered the predominant indicators of strain virulence in V. parahaemolyticus (26, 37, 38). This is substantiated by the prevalence of tdh and/or trh in clinical V. parahaemolyticus isolates and the infrequent detection of these virulence markers in food and environmental samples (9, 27, 28, 35, 40).
Investigations of the in vitro virulence effects of the TDH have demonstrated its involvement in cytotoxicity, hemolytic activity, and mouse lethality (14, 22). Although few studies have directly addressed the effects of TRH production on pathogenicity, the TRH is believed to act similarly to the TDH, based on genetic sequence similarity (2). A recent review of the pathogenicity effects of individual factors concluded that deletion of tdh does not affect cytotoxicity, indicating that the virulence of V. parahaemolyticus involves more than just the presence of tdh and/or trh (2). The type III secretion system (T3SS) of V. parahaemolyticus has been investigated as a potential indicator of strain virulence (2, 33, 37). Two nonredundant T3SSs can be found in many V. parahaemolyticus strains (2, 33). T3SS1 is involved in cytotoxicity, mouse lethality, and possibly the induction of autophagy (2, 4, 14, 33). T3SS2 appears to be involved in enterotoxicity in vitro and may play a role in the environmental fitness of strains (14, 23, 33). All V. parahaemolyticus isolates contain T3SS1 (31, 33). Two distinct lineages of T3SS2 have been described, showing correlations of tdh with T3SS2α and of trh with T3SS2β (31, 33).
In addition to the complications described above, there are other confounding factors in the identification of reliable V. parahaemolyticus virulence markers. For example, certain V. parahaemolyticus serotypes, including O3:K6, O4:K68, O1:KUT (untypeable), and O4:K12, are generally considered more virulent than others, even though there are 11 known O types and more than 70 known K types (2). Some of these pathogenicity traits are associated with V. parahaemolyticus strains isolated from specific geographical regions. For example, illnesses are more frequently reported in the Pacific Northwest, where O4:K12, trh-positive, urease-positive strains are commonly isolated from patients, than along the Gulf Coast (12). However, the level of pathogenic V. parahaemolyticus strains in shellfish and environmental samples is lower in the Pacific Northwest than along the Gulf Coast (9, 11, 19, 28, 40). Except for the association of urease production with potentially virulent trh-positive strains (12, 20), little focus has been placed on the biochemical activity of V. parahaemolyticus as a potential contributor to strain differentiation since the advent of genomics. However, biochemical activities may reveal insights into virulence potential and should not be overlooked.
In this study, a geographically diverse set of V. parahaemolyticus strains from North America, including clinical and retail oyster isolates, were characterized biochemically and serologically and were tested for the presence of known pathogenicity markers. These investigations were undertaken with the intent of revealing differences between the clinical and oyster isolates that may provide further insights into the pathogenic potential of this organism and inform the direction of future virulence mechanism studies.
Strains utilized in this study.
The 144 strains examined in this study are listed in Tables 1 and and22 along with their origins. V. parahaemolyticus oyster strains were isolated from oysters collected in a nationwide survey conducted in 2007 (9). Typical V. parahaemolyticus isolates were selected from thiosulfate-citrate-bile salts-sucrose (TCBS) plates after alkaline peptone water (APW) enrichment, and the presence of the tlh (species-specific marker), tdh, and/or trh gene was confirmed by DNA probe colony hybridization (24, 25, 29). Oyster isolates were preferentially selected based on the presence of the tdh and/or trh gene to facilitate the comparison of potentially pathogenic isolates. However, 14 strains lacking both tdh and trh were also selected as “nonpathogenic” representatives and to facilitate comparison with the clinical isolates that lacked these virulence markers. Clinical V. parahaemolyticus strains were obtained from the collection of the Centers for Disease Control and Prevention (CDC). All 77 V. parahaemolyticus isolates submitted to the CDC in 2007 from wound infections or food-borne illness were included in this study. The date of reporting of illness ranged from July 2006 to November 2007. The information provided to the CDC identified the strains as 1 blood, 5 wound, 12 “other,” and 59 stool isolates; no additional information on the isolates was available (Table 1).
Table 1
Table 1
Sources and reporting dates for the clinical isolates utilized in this study
Table 2
Table 2
Harvest dates and locations for the oyster strains utilized in this study
All the data presented in this report can be accessed at www.PATRN.net.
Biochemical characterization.
Partial biochemical profiles of all V. parahaemolyticus isolates were obtained using API 20E diagnostic strips (bioMérieux, Durham, NC) with 2% NaCl as the suspension solution. Oxidase testing was conducted using Dry Slides (BBL, Difco, Sparks, MD). Urease and Voges-Proskauer (VP) reactions were consistently ambiguous on the API strips, so those reactions were confirmed using urea (pH 6.7) and methyl red, VP (MR-VP) broths, respectively (1). The VP test was completed using VP reagent (API, Durham, NC) according to the recommendations in reference 1.
Serology.
For O antigen typing, a heavy colony suspension was prepared in a 3% NaCl solution and was then autoclaved at 121°C for 1 h. A 1-μl loopful of the autoclaved suspension was added to one drop of each O antiserum and was tested for agglutination. If the isolate did not agglutinate in any O antisera, a fresh cell suspension was made and was autoclaved for 2 h prior to testing. For K antigen typing, a 1-μl loopful of plate growth was mixed into one drop of each pool of K antisera and was tested for agglutination. If an isolate agglutinated in any K pool, the individual antisera of that pool were tested. O and K antiserum sets were purchased from Denka Seiken (Tokyo, Japan).
PCR template preparation.
Each isolate was inoculated into LB plus 1% NaCl and was incubated at 35°C with shaking (~150 rpm). After overnight incubation, 1 ml was removed, heated to 100°C for 10 min, and then placed in ice for 10 min. Samples were stored at −20°C in a manual defrost freezer until analysis. For testing, samples were thawed at room temperature, and appropriate amounts were added to the reaction mixture. Samples were refrozen for subsequent analyses.
Pathogenicity gene testing.
Primers and TaqMan-style probes were purchased from Integrated DNA Technologies (IDT; Coralville, IA), and TaqMan probes were purchased from Applied Biosystems (Foster City, CA). All sequences and the expected product sizes are provided in Table 3.
Table 3
Table 3
Primer and probe sequences utilized in this study
All 144 V. parahaemolyticus isolates were tested for the presence of the tlh (species-specific marker), tdh, and trh genes by real-time multiplex PCR as described previously (30). Strain F11-3A was used as a positive control (30).
All strains were tested for the presence of type III secretion system (T3SS) genes specific to T3SS1, T3SS2α, and T3SS2β by using primers described previously (31, 32). Strain TX2103 was used as a positive control for T3SSα and strain AQ4037 for T3SSβ (31). Both strains served as positive controls for T3SS1 (31).
The presence of T3SS1 genes (VP1670 [vscP], VP1686 [putative], VP1689 [vscK], and VP1694 [vscF]) was tested using a conventional multiplex PCR. The 30-μl reaction mixture consisted of 1.5 U Taq DNA polymerase (Roche, Indianapolis, IN), 1× buffer with MgCl2 (final concentration, 1.5 mM; Roche), 200 μM deoxynucleoside triphosphates (dNTPs) (Roche, Indianapolis, IN), 300 nM each primer (Table 3), and 3 μl template. Thermocycling was conducted in a DNA Engine thermal cycler (PTC-200; MJ Research, Waltham, MA) with the following parameters: Taq activation at 94°C for 2 min, followed by 33 cycles of amplification consisting of a denaturation step at 94°C for 45 s, annealing at 60°C for 40 s, and extension at 72°C for 45 s, with a final extension for 7 min.
The presence of T3SS2α genes (VP1362 [vopB2], VP1339 [vscC2], VP1335 [vscS2], and VP1327 [vopT]) was tested using a conventional multiplex PCR. The 30-μl reaction mixture consisted of 1.5 U Taq DNA polymerase (Roche), 1× buffer with MgCl2 (Roche), an additional 2 mM MgCl2 (final concentration, 3.5 mM; Invitrogen), 200 μM dNTPs (Roche), 500 nM each primer (Table 3), and 3 μl template. Cycling conditions were those described above for the T3SS1 screening.
For the detection of T3SS2β genes (vscC2, vopB2, vopC, vscS2), a conventional multiplex PCR was conducted using the reaction conditions described above for T3SS2α. The cycling parameters were as follows: an activation step at 94°C for 2 min, followed by 30 cycles of denaturation at 94°C for 30 s, annealing at 55°C for 30 s, and extension at 72°C for 2 min, with a final extension at 72°C for 7 min. However, amplification of all four gene products in the control strain AQ4037 was inconsistent (see Fig. S1A and B in the supplemental material), so all isolates were tested under the reaction conditions and with the cycling parameters described above in four conventional simplex PCRs (see Fig. S1C in the supplemental material), one for each gene target in T3SS2β.
All amplified products were separated using a 2% agarose (Fisher, Suwanee, GA) gel containing 0.5% ethidium bromide. If an isolate was negative for any T3SS1 or any T3SS2α gene in the multiplex assay, that isolate was retested for that gene in a simplex PCR to confirm the absence of the gene. Only strong amplification (equivalent to that of the positive control) is reported as a positive reaction. An isolate was considered positive if amplification resulting in a product of the expected size was observed in a multiplex or simplex PCR.
Statistical analysis.
Associations between the source of the isolate (clinical versus oyster) and the prevalence of selected biochemical, serotype, and virulence characteristics were evaluated by contingency table analysis. In comparisons between clinical and oyster isolates, confidence intervals for differences in the prevalence of virulence characteristics were calculated by the asymptotic method with continuity correction (13). The statistical significance of observed associations (contingency) was determined by Fisher's exact test. All statistical analyses were performed using R (34).
Biochemical characterization.
In this study, 28 of 67 (41.8%) oyster isolates and 42 of 77 (54.6%) clinical isolates were identified by the API 20E test as Vibrio parahaemolyticus. Most commonly, the misidentified isolates gave API codes for Vibrio vulnificus or Aeromonas hydrophila; however, identifications of Vibrio fluvialis, Vibrio cholerae, and Vibrio mimicus were also made. Table 4 lists the biochemical properties of the clinical and oyster isolates. As expected, all V. parahaemolyticus isolates were positive for oxidase, indole, and glucose fermentation. Only two of the oyster isolates (3%) were sucrose positive, and one clinical isolate (1.3%) was VP positive, both unusual traits for this organism. V. parahaemolyticus is generally considered to be o-nitrophenyl-β-d-galactopyranoside (ONPG) negative (no β-galactosidase production) (3, 10), but 51% of the isolates were positive. This one test was responsible for the majority (>90%) of the misidentifications by the API 20E system, indicating that the isolates otherwise produced biochemical profiles typical of V. parahaemolyticus (3). No traits distinguishing between the biochemical profiles of the clinical and oyster isolates were observed.
Table 4
Table 4
Biochemical properties of V. parahaemolyticus clinical and oyster strains examined in this study
Serology.
Among the 144 isolates tested, 35 serotypes were identified (Table 5). There were representatives of all but three (O types 2, 7, and 9) of the known O types, but nearly half (71 of 144 [49.3%]) of the isolates were untypeable for the K antigen (KUT). Fifteen isolates (10.4%) had unique serotypes within this study. Only nine of the serotypes were shared by clinical and oyster isolates (O1:KUT, O1:K20, O3:KUT, O4:KUT, O4:K8, O4:K9, O5:KUT, O10:KUT, and O11:KUT). Thirteen serotypes were found only in clinical isolates (O1:K33, O1:K56, O3:K39, O3:K56, O4:K4, O4:K13, O4:K53, O4:K63, O5:K17, O5:K30, O5:K47, O6:K18, and O8:K41), and nine serotypes were found only in oyster isolates (O1:K43, O3:K5, O4:K10, O4:K34, O4:K37, O4:K42, O6:KUT, O8:KUT, and O8:K70). O1:KUT was the dominant serotype overall, with 18 isolates (12.5%) from clinical and oyster sources, geographically distributed. O1:KUT is one of the serogroups associated with pandemic V. parahaemolyticus strains (6, 7); of the remaining pandemic serotypes, four O3:K6, one O4:K68, and one O1:K25 isolate were identified among the clinical isolates but none of the oyster isolates. The prevalence of pandemic serotypes was higher among clinical isolates (22.1%) than among oyster isolates (10.4%), but the difference was only marginally significant (P, <0.10; 95% confidence interval, −1.6%, 25%).
Table 5
Table 5
Distribution of serotypes for all 144 V. parahaemolyticus isolates based on isolation source and location
Hemolysin gene testing.
As expected, all 144 isolates tested were positive for the presence of the tlh gene, confirming their identity as V. parahaemolyticus (data not shown). For this study, oyster isolates were preferentially selected for the presence of tdh and/or trh; however, all clinical isolates submitted to the CDC in 2007 were included. Twenty-one of 77 (27%) clinical isolates and 14 of 67 (21%) oyster isolates were negative for both tdh and trh (Table 6). Thirty-five (45.5%) clinical and 23 (34.3%) oyster isolates contained both the tdh and trh genes. Nine (11.7%) clinical and two (3.0%) oyster isolates were tdh positive and trh negative; 12 (15.6%) clinical and 28 (41.8%) oyster isolates were tdh negative and trh positive.
Table 6
Table 6
Distribution of hemolysin genotypes for all 144 V. parahaemolyticus isolates based on isolation source and location
The most common virulence genotype (45%) among clinical V. parahaemolyticus isolates was positivity for both tdh and trh; all but one of these strains was isolated from stool specimens. All nine of the tdh-positive, trh-negative clinical isolates were also isolated from stool specimens. Of the 12 tdh-negative, trh-positive isolates, 8 (66.7%) were isolated from stool specimens. The 25 (37.3%) oyster isolates that were positive for tdh (whether positive or negative for trh) were from market oysters harvested between March and November from Gulf and Mid-Atlantic states. In contrast, the majority (22 of 28 [78.6%]) of tdh-negative, trh-positive isolates came from oysters harvested from the North Atlantic or Pacific Northwest during July.
T3SS gene testing.
In the testing of isolates for T3SS1 genes by multiplex PCR, 17 isolates were missing at least one of the four genes (representative strains are shown in Fig. S2 in the supplemental material). These 17 isolates were retested for amplification of each of the missing genes by simplex PCR, and 7 were found to contain the genes not amplified by the multiplex PCR (see Fig. S3 in the supplemental material). The remaining 10 isolates demonstrated weak amplification by simplex PCR, sometimes with a stronger product at a size other than that expected (see Fig. S3). Similar problems were observed in the application of the T3SS2α multiplex PCR, including weak amplification in many isolates (see Fig. S4 in the supplemental material). In the two tdh-positive, trh-negative oyster isolates, only three of the four genes were strongly amplified, even by simplex PCR (see Fig. S5 in the supplemental material). Additionally, three oyster isolates showed strong amplification of VPA1335 (vscS2) when tested by simplex PCR (see Fig. S5).
All but three of the clinical isolates (96%) contained all four T3SS1 genes (Table 7). Two (5.7%) clinical isolates positive for both tdh and trh did not amplify with primers for VP1694 (vscF), and one (8.3%) tdh-negative, trh-positive isolate was negative for VP1686. Only 60 of 67 (90%) oyster isolates amplified all four T3SS1 genes. Two isolates positive for both tdh and trh, one tdh-negative, trh-positive isolate, and two isolates negative for both tdh and trh did not amplify VP1686; one tdh-negative, trh-positive isolate and one isolate negative for both tdh and trh did not amplify VP1694 (vscF).
Table 7
Table 7
Distribution of T3SS genes for all 144 V. parahaemolyticus isolates based on isolation source and hemolysin genotypea
All nine of the tdh-positive, trh-negative clinical isolates contained all four of the T3SS2α genes. The two tdh-positive, trh-negative oyster strains did not amplify VPA1362 (vopB2) but were positive for the other three genes. The observed difference in the prevalence of VPA1362 (vopB2) among clinical versus oyster tdh-positive, trh-negative isolates was statistically significant (P, 0.018). Additionally, one oyster isolate positive for both tdh and trh, one negative for tdh and positive for trh, and one negative for both tdh and trh amplified VAP1335 (vscC2) of the T3SS2α system. Overall, the observed association between the tdh-positive, trh-negative genotype and the presence of T3SS2α system genes was statistically significant (P, <0.0001).
All four of the T3SS2β genes were detected in all 12 clinical and 28 oyster strains that were trh positive and tdh negative. Additionally all 35 clinical isolates and 22 of 23 (95.7%) oyster isolates positive for both tdh and trh amplified all four T3SS2β genes tested. The one remaining oyster isolate positive for both tdh and trh amplified all but vopC. Overall, the observed association between the tdh-positive, trh-positive and tdh-negative, trh-positive genotypes and the presence of T3SS2β system genes was statistically significant (P, <0.0001).
This study was undertaken to examine the diversity in biochemical activity, serotype, and the presence of known virulence markers (the tdh, trh, and T3SS genes) among 144 V. parahaemolyticus isolates. To our knowledge, no other study has characterized such a diverse panel of V. parahaemolyticus isolates at this level of detail. The isolates examined in this study included clinical strains from across North America isolated from July 2006 to November 2007 and oyster isolates obtained from market oysters collected across the United States in 2007. While the focus of this study was to identify differences between clinical and oyster isolates in order to provide insights into strain virulence, it should be noted that there are likely pathogenic strains among the group of oyster isolates. All potentially pathogenic (based on the presence of tdh and/or trh) oyster isolates were included in the study, as well as a proportion of oyster isolates negative for both tdh and trh similar to the proportion of clinical isolates available with the same hemolysin genotype. Our sample selection criteria, including preferential selection of tdh-negative, trh-negative strains from oysters, introduces a possibility of sample selection bias in the statistical comparisons, but the relative proportions of strains negative for both tdh and trh in the two groups of isolates, clinical and oyster, were similar.
As in previous reports, the API 20E test was unreliable for the identification of the V. parahaemolyticus isolates (8, 21), but all strains were identified by the presence of the V. parahaemolyticus species-specific tlh gene. The only novel trait observed in this set of isolates was the ability to produce β-galactosidase (ONPG test), but this was not a differential feature between clinical and oyster isolates. Historically, the majority (>90%) of V. parahaemolyticus strains have been reported to be negative for ONPG (3, 10), but 46% and 57% of the clinical and oyster isolates, respectively, were positive. This may be an evolution of V. parahaemolyticus that was previously undocumented due to the decrease in the use of biochemical examination of isolates because of its lack of reliability for species identification. Previously, urease production has been linked to the presence of the trh gene (16, 39). In this study, 95 of 96 (99%) isolates that produced urease also harbored the trh gene. However, three trh-positive strains were negative for urease production. Since this study did not test for the presence of the ure gene, it is possible that these strains simply did not express the gene for urease production under the experimental conditions used.
The most prevalent serotype seen in this study was O1:KUT, one of the serogroups associated with pandemic V. parahaemolyticus strains (6, 7). Eleven clinical and 7 oyster isolates belonged to serotype O1:KUT. However, it has been reported that serotype O1:KUT alone is not a reliable indicator of a pandemic lineage isolate (7). The remaining pandemic serotypes, O3:K6, O4:K68, and O1:K25, were infrequently identified among the clinical isolates but were not identified in any of the oyster isolates. Taken together, these data indicate a likely low prevalence of these pandemic strains in U.S. market oysters. The second most prevalent serotype was O4:K12, the most common serotype in human illnesses from Washington State (12, 20). All 11 of the O4:K12 isolates were from patient stool specimens. Although only three were from the state of Washington, the majority of others were from states that may have received seafood products from Washington. Thirteen additional serotypes were found only in clinical isolates, and nine serotypes were found only in oyster isolates. As such, serotype was the trait least shared by clinical and oyster isolates. Future work on virulence models is planned to determine whether certain serotypes, or groups of serotypes, can be predictive of pathogenic potential.
The hemolysin genes, tdh and trh, have generally been considered reliable indicators of strain virulence (2, 38). Additionally, it has been suggested that strains containing T3SS2 should also be considered more virulent than others (5). More than 90% of clinical V. parahaemolyticus strains with tdh and/or trh and T3SS2 were isolated from stool specimens, supporting the hypothesis that these hemolysin genes and the presence of T3SS2 are predictive of food-borne illness risk. In contrast, 27% of clinical V. parahaemolyticus strains in this study were negative for tdh, trh, and T3SS2, so they would, by convention, be considered avirulent. This group of strains was diverse with respect to the isolation source, with one (5%) from blood, five (24%) from wounds, seven (33%) from stool specimens, and eight (38%) reported as “other.” This could indicate that these strains are more opportunistic or have decreased virulence potential, since “other” may refer to infections resulting from recreational water activities as opposed to seafood consumption. This information does raise some concerns about the reliability of the tdh, trh, and T3SS2 genes as predictors of overall strain virulence. However, a definitive association of each isolate with a patient with food-borne illness is lacking, making it difficult to determine whether different sets of virulence factors may play a role in wound infections caused by V. parahaemolyticus versus food-borne illnesses.
Market oyster isolates in this study were preferentially selected for harboring the tdh and/or trh gene. This precludes any conclusions about the distribution of strains negative for both tdh and trh, but all strains of other genotypes (positive for both tdh and trh, tdh positive and trh negative, tdh negative and trh positive) isolated during the previous study (9) were included. All of the isolates positive for tdh (whether positive or negative for trh) were from oysters harvested from Gulf and Mid-Atlantic state estuaries from late spring to early winter. This is in agreement with a previous study of Gulf oysters and water, where pathogenic V. parahaemolyticus strains were isolated more frequently when water temperatures were elevated (18); however, there is a report describing an inverse correlation between water temperature and the isolation of tdh-positive V. parahaemolyticus strains from oysters (11).
A majority of tdh-negative, trh-positive isolates were obtained from Canada, Maine, and Washington during the early summer, indicating a potential preferential distribution of these strains in northern areas. This is reflected in the previous report that trh-positive V. parahaemolyticus strains constitute a higher proportion of the total V. parahaemolyticus population in the Mid-Atlantic than in other areas during the summer (9). It is also interesting that the remaining tdh-negative, trh-positive V. parahaemolyticus strains were isolated from Texas oysters harvested from January through May, since this is one of the few areas of the Gulf Coast where the prevalence of trh-positive strains had not been well documented.
In this study, 10 of the V. parahaemolyticus isolates were missing at least one of the four T3SS1 genes tested (even when amplified by simplex PCR), in contrast to previous reports that T3SS1 is present in all strains of V. parahaemolyticus (31, 33). VP1686 and/or VP1694 (vscF) were the genes missing from these isolates. The functions of these genes are not yet defined, and VP1686 is a “putative” open reading frame (ORF), indicating that the loss or absence of these genes may not impair the functionality of T3SS1. Because some weak amplification was observed by simplex PCR, it is possible that these strains do possess the VP1686 and VP1694 genes but have a divergent sequence that cannot be amplified efficiently with the current primer set.
Noriea et al. suggested that VPA1362 (vopB2) of T3SS2α may be a more reliable predictor of virulence than tdh, based on the absence of vopB2 in environmental isolates that contained other T3SS2α genes (31). As in that study, the vopB2 gene specific to T3SS2α was amplified in all nine of the tdh-positive, trh-negative clinical isolates. Additionally, neither of the two oyster tdh-positive, trh-negative isolates amplified VPA1362 (vopB2) by multiplex PCR. Weak amplification was observed from the two oyster isolates by simplex PCR, suggesting a potential sequence variation in the vopB2 gene that might be utilized as a more specific indicator of strain virulence than tdh.
As reported previously (31), all clinical and oyster isolates that were tdh negative and trh positive amplified all four genes specific for T3SS2β. Whereas a previous report demonstrated the absence of T3SS2α and T3SS2β in isolates positive for both tdh and trh (31), all clinical isolates and all but one oyster isolate with tdh and trh amplified all four T3SS2β genes. The one remaining oyster isolate positive for both tdh and trh amplified three of the four genes, with no amplification of the vopC effector protein. This is the first report of the presence of T3SS2β in isolates positive for both tdh and trh. This difference may be attributed to diversity in the strain panel and the use of simplex PCR to examine all isolates, whereas only the multiplex PCR was utilized in previous reports.
This study demonstrated more-reliable amplification of the T3SS genes by simplex PCR than by the previously published multiplex PCR assays. The anomalies observed in the PCR amplifications of these genes bring to light many questions about the sequence diversity of these systems. The primer sets utilized in the current study were designed based on the limited number of available sequences, which were predominately from clinical isolates. It is likely that a higher diversity of these systems will be revealed as the number of isolates examined increases. The sequence divergence of the T3SSs from the strains studied, particularly the T3SSα vopB2 gene, is a subject of investigations currently under way in our laboratory.
In summary, this study characterized a relatively diverse group of V. parahaemolyticus clinical and oyster isolates for biochemical differences and the distribution of pathogenicity factors. Surprisingly, more than one-quarter of the clinical isolates were negative for both tdh and trh and did not possess T3SS2. These results indicate that the virulence of V. parahaemolyticus is more complex than historically believed: the tdh and/or tdh and T3SS2 genes are not necessarily predictive of pathogenic potential. Serotype was a distinguishing feature of the clinical isolates; 17 of the serotypes were found only in clinical isolates. However, the variety of serotypes may be too wide for use as a predictor of virulence. Overall, this study exposes a higher level of complexity in the virulence potential of V. parahaemolyticus and brings to light concerns about the reliability of the long-standing virulence markers of the species. More-discriminatory analyses of these and additional isolates are under way in our laboratory with the goal of elucidating more-reliable predictors of V. parahaemolyticus virulence.
Supplementary Material
Supplemental material
ACKNOWLEDGMENT
The findings and conclusions in this report are those of the authors and do not necessarily represent the views of the Centers for Disease Control and Prevention.
Footnotes
Published ahead of print 25 April 2012
Supplemental material for this article may be found at http://jcm.asm.org/.
1. Becton Dickinson and Company 2003. Difco & BBL manual: manual of microbiological culture media. Becton, Dickinson and Company, Sparks, MD.
2. Broberg CA, Calder TJ, Orth K. 2011. Vibrio parahaemolyticus cell biology and pathogenicity determinants. Microbes Infect. 13:992–1001 doi:10.1016/j.micinf. 2011.06.013. [PMC free article] [PubMed]
3. Bryant TN, Lee JV, West PA, Colwell RR. 1986. A probability matrix for the identification of species of Vibrio and related genera. J. Appl. Bacteriol. 61:469–480. [PubMed]
4. Burdette DL, Yarbrough ML, Orth K. 2009. Not without cause: Vibrio parahaemolyticus induces acute autophagy and cell death. Autophagy 5:100–102. [PMC free article] [PubMed]
5. Caburlotto G, et al. 2010. Effect on human cells of environmental Vibrio parahaemolyticus strains carrying type III secretion system 2. Infect. Immun. 78:3280–3287. [PMC free article] [PubMed]
6. Chowdhury NR, et al. 2000. Molecular evidence of clonal Vibrio parahaemolyticus pandemic strains. Emerg. Infect. Dis. 6:631–636. [PMC free article] [PubMed]
7. Chowdhury NR, Stine OC, Morris JG, Nair GB. 2004. Assessment of evolution of pandemic Vibrio parahaemolyticus by multilocus sequence typing. J. Clin. Microbiol. 42:1280–1282. [PMC free article] [PubMed]
8. Croci L, et al. 2007. Comparison of different biochemical and molecular methods for the identification of Vibrio parahaemolyticus. J. Appl. Microbiol. 102:229–237 doi:10.1111/j.1365-2672.2006.03046.x. [PubMed]
9. DePaola A, et al. 2010. Bacterial and viral pathogens in live oysters: 2007 United States market survey. Appl. Environ. Microbiol. 76:2754–2768. [PMC free article] [PubMed]
10. DePaola A, Kaysner CA. 2004. Vibrio. In Merker RI, editor. (ed), Bacteriological analytical manual, 8th ed, revision A, chapter 9. U.S. Food and Drug Administration, Silver Spring, MD: http://www.fda.gov/Food/ScienceResearch/LaboratoryMethods/BacteriologicalAnalyticalManualBAM/default.htm.
11. DePaola A, Nordstrom JL, Bowers JC, Wells JG, Cook DW. 2003. Seasonal abundance of total and pathogenic Vibrio parahaemolyticus in Alabama oysters. Appl. Environ. Microbiol. 69:1521–1526. [PMC free article] [PubMed]
12. DePaola A, et al. 2003. Molecular, serological, and virulence characteristics of Vibrio parahaemolyticus isolated from environmental, food, and clinical sources in North America and Asia. Appl. Environ. Microbiol. 69:3999–4005. [PMC free article] [PubMed]
13. Fleiss JL. 1981. Statistical methods for rates and proportions. John Wiley & Sons, Inc., New York, NY.
14. Hiyoshi H, Kodama T, Iida T, Honda T. 2010. Contribution of Vibrio parahaemolyticus virulence factors to cytotoxicity, enterotoxicity, and lethality in mice. Infect. Immun. 78:1772–1780. [PMC free article] [PubMed]
15. Honda T, Ni YX, Miwatani T. 1988. Purification and characterization of a hemolysin produced by a clinical isolate of Kanagawa phenomenon-negative Vibrio parahaemolyticus and related to the thermostable direct hemolysin. Infect. Immun. 56:961–965. [PMC free article] [PubMed]
16. Iida T, et al. 1997. Evidence for genetic linkage between the ure and trh genes in Vibrio parahaemolyticus. J. Med. Microbiol. 46:639–645. [PubMed]
17. Iwamoto M, Ayers T, Mahon BE, Swerdlow DL. 2010. Epidemiology of seafood-associated infections in the United States. Clin. Microbiol. Rev. 23:399–411. [PMC free article] [PubMed]
18. Johnson CN, et al. 2009. Genetic relatedness among tdh+ and trh+ Vibrio parahaemolyticus cultured from Gulf of Mexico oysters (Crassostrea virginica) and surrounding water and sediment. Microb. Ecol. 57:437–443. [PubMed]
19. Kaysner CA, Abeyta C, Jr, Stott RF, Lilja JL, Wekell MM. 1990. Incidence of urea-hydrolyzing Vibrio parahaemolyticus in Willapa Bay, Washington. Appl. Environ. Microbiol. 56:904–907. [PMC free article] [PubMed]
20. Kaysner CA, et al. 1994. Urea hydrolysis can predict the potential pathogenicity of Vibrio parahaemolyticus strains isolated in the Pacific Northwest. Appl. Environ. Microbiol. 60:3020–3022. [PMC free article] [PubMed]
21. Martinez-Urtaza J, Lozano-Leon A, Vina-Feas A, de Novoa J, Garcia-Martin O. 2006. Differences in the API 20E biochemical patterns of clinical and environmental Vibrio parahaemolyticus isolates. FEMS Microbiol. Lett. 255:75–81. [PubMed]
22. Matsuda S, et al. 2010. Association of Vibrio parahaemolyticus thermostable direct hemolysin with lipid rafts is essential for cytotoxicity but not hemolytic activity. Infect. Immun. 78:603–610. [PMC free article] [PubMed]
23. Matz C, Nouri B, McCarter L, Martinez-Urtaza J. 2011. Acquired type III secretion system determines environmental fitness of epidemic Vibrio parahaemolyticus in the interaction with bacterivorous protists. PLoS One 6:e20275 doi:10.1371/journal.pone.0020275. [PMC free article] [PubMed]
24. McCarthy SA, DePaola A, Cook DW, Kaysner CA, Hill WE. 1999. Evaluation of alkaline phosphatase- and digoxigenin-labelled probes for detection of the thermolabile hemolysin (tlh) gene of Vibrio parahaemolyticus. Lett. Appl. Microbiol. 28:66–70. [PubMed]
25. McCarthy SA, DePaola A, Kaysner CA, Hill WE, Cook DW. 2000. Evaluation of nonisotopic DNA hybridization methods for detection of the tdh gene of Vibrio parahaemolyticus. J. Food Prot. 63:1660–1664. [PubMed]
26. Nishibuchi M, Ishibashi M, Takeda Y, Kaper JB. 1985. Detection of the thermostable direct hemolysin gene and related DNA sequences in Vibrio parahaemolyticus and other Vibrio species by the DNA colony hybridization test. Infect. Immun. 49:481–486. [PMC free article] [PubMed]
27. Nordstrom JL, DePaola A. 2003. Improved recovery of pathogenic Vibrio parahaemolyticus from oysters using colony hybridization following enrichment. J. Microbiol. Methods 52:273–277. [PubMed]
28. Nordstrom JL, et al. 2004. Effect of intertidal exposure on Vibrio parahaemolyticus levels in Pacific Northwest oysters. J. Food Prot. 67:2178–2182. [PubMed]
29. Nordstrom JL, et al. 2006. Evaluation of an alkaline phosphatase-labeled oligonucleotide probe for the detection and enumeration of the thermostable-related hemolysin (trh) gene of Vibrio parahaemolyticus. J. Food Prot. 69:2770–2772. [PubMed]
30. Nordstrom JL, Vickery MC, Blackstone GM, Murray SL, DePaola A. 2007. Development of a multiplex real-time PCR assay with an internal amplification control for the detection of total and pathogenic Vibrio parahaemolyticus bacteria in oysters. Appl. Environ. Microbiol. 73:5840–5847. [PMC free article] [PubMed]
31. Noriea NF, III, Johnson CN, Griffitt KJ, Grimes DJ. 2010. Distribution of type III secretion systems in Vibrio parahaemolyticus from the northern Gulf of Mexico. J. Appl. Microbiol. 109:953–962. [PubMed]
32. Okada N, et al. 2009. Identification and characterization of a novel type III secretion system in trh-positive Vibrio parahaemolyticus strain TH3996 reveal genetic lineage and diversity of pathogenic machinery beyond the species level. Infect. Immun. 77:904–913. [PMC free article] [PubMed]
33. Park KS, et al. 2004. Functional characterization of two type III secretion systems of Vibrio parahaemolyticus. Infect. Immun. 72:6659–6665. [PMC free article] [PubMed]
34. R Development Core Team 7 March 2011, accession date R: a language and environment for statistical computing. R Foundation for Statistical Computing, Vienna, Austria: http://www.R-project.org.
35. Roque A, et al. 2009. Detection and identification of tdh-negative and trh-positive Vibrio parahaemolyticus strains from four species of cultured bivalve molluscs on the Spanish Mediterranean Coast. Appl. Environ. Microbiol. 75:7574–7577 doi:10.1128/AEM.00772-09. [PMC free article] [PubMed]
36. Scallan E, et al. 2011. Foodborne illness acquired in the United States—major pathogens. Emerg. Infect. Dis. 17:7–15. [PMC free article] [PubMed]
37. Shimohata T, Takahashi A. 2010. Diarrhea induced by infection of Vibrio parahaemolyticus. J. Med. Invest. 57:179–182. [PubMed]
38. Su YC, Liu C. 2007. Vibrio parahaemolyticus: a concern of seafood safety. Food Microbiol. 24:549–558. [PubMed]
39. Suthienkul O, et al. 1995. Urease production correlates with possession of the trh gene in Vibrio parahaemolyticus strains isolated in Thailand. J. Infect. Dis. 172:1405–1408. [PubMed]
40. Zimmerman AM, et al. 2007. Variability of total and pathogenic Vibrio parahaemolyticus densities in northern Gulf of Mexico water and oysters. Appl. Environ. Microbiol. 73:7589–7596. [PMC free article] [PubMed]
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