Skip to main page content

• HOME
• Current Issue
• Archive
• Alerts
• About ASM
• Contact us
• Tech Support
• Journals.ASM.org

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via Google Scholar Google Scholar Articles by Datta, S. Articles by Simonson, J. G. Search for Related Content PubMed PubMed Citation Articles by Datta, S. Articles by Simonson, J. G.

 Previous Article  |  Next Article 

Clinical and Vaccine Immunology, October 2008, p. 1541-1546, Vol. 15, No. 10
1071-412X/08/$08.00+0     doi:10.1128/CVI.00141-08
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Immunomagnetic Separation and Coagglutination of Vibrio parahaemolyticus with Anti-Flagellar Protein Monoclonal Antibody

S. Datta,¹ M. E. Janes,¹ and J. G. Simonson^2*

Department of Food Science,¹ Department of Agricultural Chemistry, Louisiana State University AgCenter, Baton Rouge, Louisiana 70803²

Received 26 March 2008/ Returned for modification 13 June 2008/ Accepted 15 August 2008

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS AND DISCUSSION REFERENCES

 
Mice were immunized by injection of Vibrio parahaemolyticus ATCC 17802 polar flagellin in order to produce monoclonal antibodies (mAbs). mAbs were analyzed by anti-H enzyme-linked immunosorbent assay using V. parahaemolyticus polar flagellar cores. The mAb exhibiting the highest anti-H titer was coated onto Cowan I Staphylococcus aureus cells at a concentration of 75 µg/ml cell suspension and used for slide coagglutination. Of 41 isolates identified genetically as V. parahaemolyticus, 100% coagglutinated with the anti-H mAb within 30 s, and the mAb did not react with 30 isolates identified as Vibrio vulnificus. A strong coagglutination reaction with V. parahaemolyticus ATCC 17802 was still observed when the S. aureus cells were armed with as little as 15 µg of mAb/ml S. aureus cell suspension. At this concentration, the mAb cross-reacted with three other Vibrio species, suggesting that they share an identical H antigen or antigens. The anti-H mAb was then used to optimize an immunomagnetic separation protocol which exhibited from 35% to about 45% binding of 10² to 10³ V. parahaemolyticus cells in phosphate-buffered saline. The mAb would be useful for the rapid and selective isolation, concentration, and detection of V. parahaemolyticus cells from environmental sources.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS AND DISCUSSION REFERENCES

 
Vibrio parahaemolyticus is a naturally occurring marine bacterium responsible for the majority of seafood-associated human gastroenteritis cases in the United States and is considered an important seafood-borne pathogen throughout the world (14, 28, 43). Conventional bacteriological methods for the detection and enumeration of V. parahaemolyticus bacteria can be costly in labor, materials, and time (25), while the expeditious identification of V. parahaemolyticus in the laboratory is desirable. Ideally, a method is needed which can easily detect and enumerate V. parahaemolyticus cells by the direct examination of shellfish, seafood, or water and which does not involve lengthy enrichment steps or overnight incubation.

After several V. parahaemolyticus outbreaks in the United States (8, 9, 16), the Interstate Shellfish Sanitation Conference (ISSC) implemented a plan for monitoring the levels of V. parahaemolyticus bacteria in freshly harvested oysters. Since the standard most probable number (MPN)/biochemical method for enumerating V. parahaemolyticus bacteria (17) was so labor-intensive and time-consuming, the procedure the ISSC recommended involved plating oyster homogenates directly onto agar plates and, after an overnight incubation, transferring resultant colonies to filters that could be hybridized with DNA probes to detect total (tlh) and pathogenic (tdh) strains of V. parahaemolyticus (12). The probes were successfully used for the direct examination of total and pathogenic V. parahaemolyticus in oysters harvested from Washington, Texas, and New York (16). Gooch et al. (20) compared two direct plating methods to the MPN protocol using probes specific for tlh to confirm V. parahaemolyticus isolates in Alabama oysters. They concluded that both direct plating methods were equivalent to, yet faster and less labor-intensive than, the MPN method for the enumeration and confirmation of total V. parahaemolyticus cells in oyster homogenates. Since then, probes for tlh and tdh have been successfully and routinely employed for the detection of V. parahaemolyticus in retail oysters (11), shellfish and sediments (15), and freshly harvested oysters (15, 24). However, the direct plating/DNA probe hybridization procedure still requires an overnight incubation and exhibits, at best, a detection sensitivity of 10 CFU/g.

It has also been shown that the detection of V. parahaemolyticus by PCR is specific and less time-consuming than the conventional bacteriological method (2, 7). However, to achieve the desired detection sensitivity of 10¹ to 10² CFU/g, either enrichment cultures were employed or homogenates had to be subjected to DNA purification prior to PCR analysis. A real-time PCR (RT-PCR) method based on the amplification of the tdh gene was developed and evaluated for the detection of pathogenic V. parahaemolyticus (6). The research demonstrated that RT-PCR was a rapid and reliable technique for detecting virulent V. parahaemolyticus in pure cultures and oyster homogenate enrichment cultures. Kaufman et al. (23) used RT-PCR for the enumeration of total V. parahaemolyticus directly from oyster mantle fluid; however, a loss of efficiency was observed when the numbers of V. parahaemolyticus were low and/or PCR inhibitors were present in the mantle fluids of certain oysters.

One approach to circumvent these problems and improve the recovery and detection of V. parahaemolyticus in seafood and shellfish samples is through the use of immunomagnetic separation (IMS). IMS has been successfully used to concentrate and isolate numerous pathogens (21, 26, 30) and has often been used as a pre-PCR step to concentrate and separate the organism of interest from polymerase inhibitors in the sample matrix (13, 18, 19, 31). A successful IMS method for the concentration of V. parahaemolyticus cells utilizing species-specific antibodies coupled to PCR would improve detection sensitivity and separate V. parahaemolyticus from other bacteria, eliminating interference with DNA amplification. The employment of IMS as a pre-PCR step would also concentrate the pathogen to a suitable volume and separate V. parahaemolyticus from inhibitory factors in shellfish or seafood homogenates, thereby eliminating the need for DNA purification or enrichment cultures. The first requirement for optimizing an IMS method for the isolation of V. parahaemolyticus would be to generate antibodies which will specifically recognize the pathogen.

Species within the genus Vibrio can be identified serologically through the detection of unique H antigens expressed in the core protein of the polar flagellum (4, 5, 32, 39, 44, 45). Based on this knowledge, species-specific anti-Vibrio vulnificus H polyclonal antibodies were produced and used to construct coagglutination reagents which reacted with 99% of those bacterial isolates identified bacteriologically as V. vulnificus (40). Since little cross-reaction with other Vibrio species was observed, this offered a specific, rapid, and economical approach to the detection of the pathogen one step beyond primary isolation. Vibrio cholerae and V. parahaemolyticus, however, exhibit H determinants shared with other vibrios, which complicates the use of polyclonal anti-H antibodies to detect these pathogens serologically (5, 38, 39, 40, 45). This situation was rectified, in the case of V. cholerae, by the production of several anti-H monoclonal antibodies (mAbs) which reacted only with the flagella of V. cholerae and V. mimicus, thereby eliminating cross-reaction with V. fluvialis, V. metschnikovii, V. anguillarum, and V. ordalii (41). Likewise, the production of mAbs reactive with the polar flagellar core of V. parahaemolyticus might also eliminate the cross-reaction observed with V. alginolyticus, V. natriegens, V. harveyi, V. campbellii, and V. carchariae, thereby resulting in the desired species specificity for use in an IMS protocol. In addition, the creation and preservation of anti-H-secreting hybridomal cell lines would ensure the continuing availability of anti-V. parahaemolyticus mAbs for analytical purposes. This report describes the production and analysis of several mAbs that are reactive with the polar flagellar core of V. parahaemolyticus and the use of one of the mAbs in an optimized IMS protocol.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS AND DISCUSSION REFERENCES

 
Flagellar core purification. Flagellar cores were purified from the polar flagella of V. parahaemolyticus ATCC 17802 by methods previously described (40). Briefly, V. parahaemolyticus cells were grown overnight in alkaline peptone broth on a shaker at 35°C. The cells were harvested by centrifugation, resuspended in chilled saline, and blended, and the sheared flagella were separated from the bacterial cell bodies and sheath material by differential centrifugation and cesium chloride density gradient ultracentrifugation in the presence of Triton X-100. The purified cores were examined by transmission electron microscopy (TEM) to verify that naked cores, free of sheath material and cell debris, were present. The flagellin concentration was determined by bicinchoninic acid protein assay (Pierce, Rockford, IL).

Electron microscopy. Carbon-coated grids were floated on drops of either a suspension of a V. parahaemolyticus ATCC 17802 nutrient broth culture containing 3% NaCl (NB+) which had been incubated for 18 h at 35°C, centrifuged, and resuspended in 0.01 M phosphate-buffered saline (PBS), pH 7.0 or purified flagellar cores. The flagella were then negatively stained with 2% uranyl acetate.

PAGE. The approximate molecular weight of the purified flagellar core protein was determined by using sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) under reducing conditions using a 10% acrylamide resolving gel. Molecular-mass markers (10 to 250 kDa) were used as standards to estimate the molecular mass of the V. parahaemolyticus flagellin (Bio-Rad, Hercules, CA).

Immunization. BALB/c mice were immunized at 2-week intervals for 4 to 8 weeks by intraperitoneal injection of 50 µg purified V. parahaemolyticus polar flagellar core protein. Mice exhibiting elevated anti-H flocculation titers (45) were boosted with 50 µg of flagellin, and their spleen cells were collected after 3 days.

Hybridoma production. The method used to promote cell fusion was modified from that described previously (41). Briefly, spleen cells from immunized mice were fused at a 4:1 ratio with log-phase P3X63Ag8.653 nonsecreting myeloma cells by the slow addition of a 50% polyethylene glycol solution (Hybrimax; Sigma Chemical Co., St. Louis, MO). The cells were sedimented and resuspended in RPMI 1640 containing 15% fetal bovine serum, 2 mM glutamine, 1% nonessential amino acids, 100 units/ml penicillin-streptomycin, 100 µM hypoxanthine, 4 x 10^–7 M aminopterin, 1.6 x 10^–5 M thymidine, and 10% hybridoma cloning factor (BioVeris Corp., Gaithersburg, MD). The cells were then distributed in 96-well, flat-bottom tissue culture plates. Cell cultures were fed and maintained at 37°C in a humidified atmosphere of 5 to 7% CO₂.

Anti-H ELISA. Supernatant fluid from each well exhibiting hybridomal growth was tested by enzyme-linked immunosorbent assay (ELISA) for the presence of anti-V. parahaemolyticus H antibody by using V. parahaemolyticus polar flagellar cores (2 µg protein/well) bound to Costar high-binding microtiter plates (Corning Inc., Corning, NY) and alkaline phosphatase-labeled rabbit anti-mouse immunoglobulin G (IgG; Sigma Chemical Co., St. Louis, MO) diluted 1:500 as described previously (41). Culture supernatant fluids which produced an absorbance of at least 0.5 were initially considered positive. Anti-H-secreting hybridomas were cloned at least two times by limiting dilution and stored in liquid nitrogen.

mAb purification. Three hybridomas secreting V. parahaemolyticus anti-H mAbs which exhibited ELISA absorbance of >1.0 were each expanded to 1 liter of cell culture medium not containing aminopterin, thymidine, or hypoxanthine and were incubated for 2 weeks. Culture fluids were clarified by centrifugation, and IgG was precipitated by the addition of (NH₄)₂SO₄ to 50% saturation. The precipitates were collected by centrifugation, dissolved in 50 to 100 ml 0.067 M PBS, pH 7.2 to 7.4, and dialyzed against several changes of buffer. After clarification through a 0.45-µm filter, IgG was purified by affinity chromatography on protein A-Sepharose (Sigma Chemical Co., St. Louis, MO), following the manufacturer's instructions. The mAbs were isotyped by ELISA (Southern Biotech, Birmingham, AL), and the IgG concentration was determined by bicinchoninic acid protein assay.

Coagglutination. Formalin-fixed Staphylococcus aureus Cowan 1 ATCC 12598 cells were prepared by methods reported previously (40). Anti-H coagglutination reagents were prepared by mixing 75 µg of each mAb with 1 ml of S. aureus cells. Each of the three anti-H coagglutination reagents was tested against the following Vibrio species: V. parahaemolyticus ATCC 17802, V. parahaemolyticus ATCC 33847, V. vulnificus ATCC 27562, V. cholerae ATCC 14035, V. mimicus ATCC 33653, V. fluvialis ATCC 33809, V. natriegens ATCC 14048, V. alginolyticus ATCC 33787, V. harveyi ATCC 14126, V. harveyi ATCC 35084, and V. campbellii ATCC 25920. The anti-H mAb, designated 3-F-3, which demonstrated the strongest slide coagglutination reaction against the vaccine V. parahaemolyticus strain was then tested against 41 additional V. parahaemolyticus and 30 V. vulnificus clinical and environmental strains. Each Vibrio isolate tested by coagglutination was grown on peptone agar slants (1% peptone, 2% NaCl, 0.2% yeast extract, 1.5% agar) for 24 h at 35°C. The bacterial cells were harvested in 1 ml of 0.1 M Tris buffer (pH 7.8) containing 0.1 mM EDTA, 1% Triton X-100, and 0.001% thimerosal (TET buffer). One drop (30 to 50 µl) of the cell suspension was placed on a clean glass plate, and 30 to 50 µl of coagglutination reagent was placed adjacent to it. The two drops were mixed and observed for 3 min over a light box for evidence of agglutination.

IMS. Vibrio parahaemolyticus ATCC 17802 was grown in NB+ at 35°C for 18 h. Serial 10-fold dilutions of the culture were made in 0.1 M PBS, pH 7.2. In separate experiments, 0.1 ml of cell suspension was removed from selected dilutions and placed in sterile Eppendorf tubes containing 0.9 ml PBS (nine tubes/dilution). Either 20 µl (about 30 µg) of mAb 3-F-3 or 20 µl of PBS was added to each test or control tube (three test and six control tubes/dilution). The tubes were then placed on a rocker and gently agitated for 30 min at 25°C. Twenty microliters (10⁷) of paramagnetic beads coated with sheep anti-mouse IgG (Dynabeads M-280; Dynal Biotech, Oslo, Norway) which were washed once with PBS and suspended in PBS containing 0.1% BSA and 0.02% NaN₃ were added to each test tube, and one set of three control tubes and the test tubes were agitated for an additional 5 min. The beads were then isolated by placement of the tubes in a magnetic concentrator for 5 min, and the supernatant fluid was carefully removed. Aliquots of the aspirated fluid were spread on NB+ agar plates, and the plates were incubated overnight at 35°C. Vibrio parahaemolyticus colonies were counted to determine the number of bacteria which did not bind to the immunomagnetic beads (IMB) and the number of cells present in control tubes containing no IMB (PBS control tubes).

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS AND DISCUSSION REFERENCES

 
Production and characterization of mAbs. Prior to the immunization of the mice, the purified V. parahaemolyticus polar flagellar cores were examined by TEM to ensure that flagellar cores, free from cell debris and flagellar sheath, were obtained (Fig. 1A). When subjected to SDS-PAGE, the purified flagellar cores exhibited a major protein band corresponding to approximately 45 kDa (Fig. 2). Two fusion experiments yielded about 20 hybridomas producing mAbs that resulted in an absorbance reading higher than 0.5 by anti-H ELISA. Only hybridomas secreting IgG were selected, by intentionally using a second antibody-enzyme conjugate specific for mouse IgG. After the positive hybridomas were cloned two or three times, three stable and rapidly growing hybridomas secreting mAbs specific for V. parahaemolyticus polar flagellar cores when tested by anti-H ELISA were selected and expanded in approximately 1 liter of cell culture fluid. After antibody purification and subclass determination for each mAb, the anti-H ELISA titers were determined and defined as the reciprocal of the highest dilution of mAb, starting with 250 µg of IgG in well 1 of a microtiter plate, that gave an absorbance reading of 0.2 or higher in three separate experiments (Table 1). The mAb produced by hybridoma 3-F-3 exhibited a much higher anti-H titer than the other two mAbs, indicating higher affinity for the V. parahaemolyticus flagellar core protein.

View larger version (155K): [in this window] [in a new window]

FIG. 1. Electron micrograph of V. parahaemolyticus ATCC 17802 purified flagellar cores (A) and sheathed and unsheathed flagella (B).

View larger version (45K): [in this window] [in a new window]

FIG. 2. Results of SDS-PAGE of purified V. parahaemolyticus flagellar core protein. Lanes: 1, standard protein markers; 2, 1.5 µg purified flagellin.

View this table: [in this window] [in a new window]

TABLE 1. Analysis of Abs purified from cell culture fluid

Coagglutination reactions. The serological specificities of the three selected mAbs were assessed by slide coagglutination. Each of the three selected mAbs was coated onto S. aureus cells, and the reagents were tested for positive slide coagglutination reactions with several Vibrio species which had been harvested and held in TET buffer for 24 h (Table 2). Each of the three mAbs exhibited positive coagglutination reactions with homologous strains of V. parahaemolyticus and heterologous V. alginolyticus, V. harveyi, and V. campbellii strains. Even when S. aureus cells were armed with as little as 15 µg of mAb 3-F-3, positive agglutination reactions were still observed with these Vibrio species. Further dilution of mAb 3-F-3 resulted in the elimination of a positive agglutination reaction within a minute when tested against the V. parahaemolyticus vaccine strain. As expected, no reaction was observed between the anti-V. parahaemolyticus coagglutination reagents and V. vulnificus, V. cholerae, V. mimicus, and V. fluvialis. These results substantiate those of earlier studies with polyclonal antibodies in which V. parahaemolyticus anti-H sera cross-reacted with V. alginolyticus, V. campbellii, V. harveyi, V. carchariae, and V. natriegens (38, 39, 40, 45), but not other vibrios tested, including V. cholerae, V. mimicus, V. fluvialis, V. metschnikovii, V. anguillarum, and V. vulnificus (32, 38, 39, 46). Only the cross-reaction observed between V. natriegens and polyclonal anti-V. parahaemolyticus H serum (40) was eliminated by the use of the mAbs selected. It is still possible that a V. parahaemolyticus species-specific polar H antigen exists since the coagglutination observed with V. alginolyticus, V. harveyi, V. carchariae, V. natriegens, and V. campbellii was eliminated by the absorption of polyclonal anti-V. parahaemolyticus H serum with formalin-killed V. campbellii cells (40). In addition, Hongprayoon (22) reported the isolation of a hybridoma that produced anti-H mAb specific for V. parahaemolyticus that only cross-reacted with V. alginolyticus, and not with 28 other Vibrio species tested. Unfortunately, the hybridoma and the mAb it produced were lost. The V. parahaemolyticus species-specific H antigen may represent a minor flagellar antigen compared to the H antigen shared with the four heterologous Vibrio species. Thus, many more hybridomas would have to be created and screened before one producing species-specific mAb could be detected.

View this table: [in this window] [in a new window]

TABLE 2. H antigen relationships among Vibrio species examined serologically by coagglutination with anti-V. parahaemolyticus H mAbsa

Shinoda and colleagues (36, 37) reported that the sheathed polar and lateral, nonsheathed flagella produced by V. parahaemolyticus differed morphologically and immunologically. To further complicate matters, V. parahaemolyticus polar and lateral flagella were shown to possess a surface antigen which was present on intact flagellar cores and an internal antigen which was revealed when the flagellar cores were denatured into flagellin monomers (33, 34, 35). It was concluded that the surface antigen-antibody reaction was responsible for H agglutination, whereas the internal antigen-antibody reaction produced the precipitin lines observed in gel diffusion tests. In 1992, Chen et al. (10) also found that many mAbs raised against V. alginolyticus recognized a polar flagellar antigen of 52 kDa common to V. alginolyticus and V. parahaemolyticus. This common flagellar antigenic determinant represented an internal antigen since it was exhibited by denatured flagella and detected by gel immunoblot. The mAbs that were produced in this study should have been generated in response to immunization with V. parahaemolyticus polar flagellar cores only since V. parahaemolyticus does not produce lateral flagella when grown in liquid medium (1, 3, 27, 35, 36). In addition, when analyzed by SDS-PAGE, the lateral flagellar core is composed of a flagellin with a molecular mass of 27 kDa (29), whereas the polar flagellins of V. parahaemolyticus and V. cholerae exhibit a molecular mass of about 45 kDa (29, 48).

The flagellar core preparation used in this study was composed of a single flagellin with an approximate molecular mass of 45 kDa on SDS-PAGE (Fig. 2). Thus, the cross-reactions observed must have occurred from a shared H antigen displayed on the polar flagellum. In addition, the three mAbs selected also must have recognized a surface antigen since intact flagellar cores were used as the murine immunogen and also as the antigen in all anti-H ELISA determinations. Most importantly, mAb specificity was determined by anti-H slide coagglutination which could only occur by a surface antigen-antibody reaction. One might also argue that the mAbs analyzed could be reactive to V. parahaemolyticus polar flagellar sheath. This, however, is unlikely since the flagellar sheath was stripped from the core by suspension of the sheared flagella in TET buffer and several rounds of differential centrifugation (40). Thomashow and Rittenberg (47) and Yang et al. (48) demonstrated that the flagellar sheaths of Bdellovibrio bacteriovorus and V. cholerae were readily solubilized by Triton X-100. Also, examination of the V. parahaemolyticus purified flagellar cores by TEM prior to immunization revealed no contaminating sheath material or cellular debris (Fig. 1A). In addition, the morphology of the purified flagellar cores and intact flagella obtained from an overnight V. parahaemolyticus 17802 broth culture were studied by electron microscopy. Both unsheathed and sheathed flagella were evident in the broth culture (Fig. 1B). The diameter of the sheathed flagellum was measured and found to be about 30 nm. The diameters of the unsheathed flagellar cores from the broth culture (Fig. 1B) and the purified flagellar cores (Fig. 1A) were about 15 and 20 nm, respectively, indicating that the purified flagellar core preparation that was used as the mouse immunogen was free of sheath material. These results demonstrate that V. parahaemolyticus, V. alginolyticus, V. harveyi, and V. campbellii must share at least one H antigen displayed on the surface of their polar flagellar cores.

One of the goals of this investigation was to assess the efficiency of an anti-H mAb for the isolation and concentration of V. parahaemolyticus cells in an IMS protocol. Skjerve et al. (42) demonstrated that a slide coagglutination reaction was a simple and reliable tool to assess the ability of antibody-coated IMB to bind bacterial strains of interest in an IMS protocol. Since the mAb produced by hybridoma 3-F-3 exhibited the highest affinity for purified V. parahaemolyticus flagellar cores and the strongest coagglutination reaction against the two V. parahaemolyticus strains originally tested, it was selected for use in the development of a V. parahaemolyticus IMS method. The serological specificity and potential reactivity of mAb 3-F-3 in an IMS protocol was further examined by testing additional V. parahaemolyticus and V. vulnificus clinical and environmental strains by S. aureus slide coagglutination. Of 41 isolates identified genetically as V. parahaemolyticus, 41 (100%) were coagglutinated with the anti-H mAb within 30 s, and the mAb did not react with 30 isolates identified as V. vulnificus (data not shown).

IMS of V. parahaemolyticus. Several guidelines were followed for the optimization of the IMS method. First of all, there are two ways that antibodies can be used in an IMS protocol. In the direct method, the antigen-specific antibody is first coated onto IMB, which are then mixed with the test sample. The indirect method differs from the direct approach in that the organisms of interest are first incubated with an excess of antigen-specific antibody and then the IMB are added to the test sample. Roberts and Hirst (31) found that indirect IMS produced consistently better results for the detection of Mycobacterium ulcerans from heterogenous samples, increasing the sensitivity up to 10-fold. Corona-Barrera et al. (13) also found that the indirect IMS method was 10- to 100-fold more sensitive for the detection of Brachyspira species in porcine feces. Based on this information, we used mAb 3-F-3 in an indirect IMS protocol by incubating an excess of the mAb (30 µg/test sample) with target V. parahaemolyticus cells prior to the addition of the IMB. Secondly, the concentration of IMB employed per test sample was predetermined at 10⁷ since other successful IMS methods recommended bead concentrations in this range (18, 26, 31, 42). Skjerve et al. (42) determined that at this concentration of IMB, 5 µg of mAb exhibited near maximum Listeria monocytogenes binding, while Roberts and Hirst (31) found that 15 µg IgG was required for optimal coating of the IMB. We used 30 µg 3-F-3 IgG/test sample for indirect V. parahaemolyticus IMS since that concentration of mAb was slightly in excess of the optimal range determined by other researchers. Finally, it has been shown that not just a single bacterial cell but often clumps of cells are bound to each bead during IMS (21, 30, 31). Since more than one cell can bind to a single bead, determining the number of bacteria bound to the IMB by plating the IMB and counting the resulting CFU would not be accurate. Therefore, the numbers of cells bound by the IMB in this study was determined indirectly by counting the number of unbound V. parahaemolyticus cells in the aspirated supernatant fluid.

One objective in developing an IMS method for the isolation of V. parahaemolyticus was to increase the detection sensitivity. Thus, the detection sensitivity, and not the maximum binding capacity of the anti-H-coated IMB, was explored in this study. Skjerve et al. (42) demonstrated a sensitivity of 10 to 100 L. monocytogenes CFU/ml PBS. They found that an increase in the number of bacteria in suspension resulted in an increase in the number of bacteria bound to the IMB from <10% (100 L. monocytogenes CFU/ml) to 50% (1.5 x 10⁴ L. monocytogenes CFU/ml). Likewise, when Roberts and Hirst (31) mixed 10-fold dilutions of M. ulcerans cells with nontarget bacteria, the IMB were unable to capture less than about 100 M. ulcerans CFU. The maximum IMB binding capacity for L. monocytogenes and Mycobacterium paratuberculosis was about 10⁴ to 10⁵ CFU/ml when 10⁶ to 10⁷ antibody-coated IMB were employed for each test sample (21, 42). When mAb 3-F-3 was employed in the IMS protocol, an average binding of about 35% was exhibited at 10² V. parahaemolyticus CFU/ml and 45% at 10³ CFU/ml (Table 3). The binding of V. parahaemolyticus at lower concentrations (<100 CFU/ml) could not be determined accurately by plating methods alone since 0.1-ml aliquots were spread on the agar plates and further dilution resulted in less than 10 CFU/plate. About 15% nonspecific binding to the IMB was observed at 10³ V. parahaemolyticus CFU/ml. Fluit et al. (19) developed an IMS coupled to PCR for the detection of low numbers of L. monocytogenes cells in food samples. The procedure involved the isolation of Listeria cells by IMS using genus-specific mAbs followed by PCR amplification of an L. monocytogenes-specific DNA sequence. The method served to separate Listeria cells from PCR-inhibitory factors present in selective food enrichment broths, allowing a detection sensitivity of 1 CFU/g. A similar method could be employed using 3-F-3 mAb to capture V. parahaemolyticus cells. Even if cross-reacting Vibrio species were bound by the mAb to the IMB or attached to the IMB by nonspecific binding, V. parahaemolyticus cells could be specifically detected by coupling the IMS to tdh or tlh PCR amplification.

View this table: [in this window] [in a new window]

TABLE 3. Results of IMS of V. parahaemolyticus using anti-H mAb

In future research, we will concentrate on the detection of V. parahaemolyticus by anti-H IMS in mixed bacterial cultures and oyster homogenates. It is also important to couple the IMS to RT-PCR to determine if low levels of V. parahaemolyticus captured by IMS can be quantified and to explore the limits of V. parahaemolyticus detection sensitivity.

 
We thank Cindy Henk of the LSU Socolofsky Microscopy Center for her assistance in the preparation of the electron micrographs. We also thank the laboratory of Lee-Ann Jaykus for providing the environmental V. parahaemolyticus and V. vulnificus cultures used in this investigation.

 
* Corresponding author. Mailing address: 102 Agricultural Chemistry Building, Highland Road, LSU, Baton Rouge, LA 70803. Phone: (225) 342-5812. Fax: (225) 342-0027. E-mail: jsimonson{at}agctr.lsu.edu

Published ahead of print on 27 August 2008.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS AND DISCUSSION REFERENCES

 

1
1. Allen, R. D., and P. Baumann. 1971. Structure and arrangement of flagella in species of the genus Beneckea and Photobacterium fischeri. J. Bacteriol. 107:295-302.[Abstract/Free Full Text]
2
2. Bej, A. K., D. P. Patterson, C. W. Brasher, M. C. Vickery, D. D. Jones, and C. A. Kaysner. 1999. Detection of total and hemolysin-producing Vibrio parahaemolyticus in shellfish using multiplex PCR amplification of tlh, tdh and trh. J. Microbiol. Methods 36:215-225.[CrossRef][Medline]
3
3. Belas, M. R., and R. R. Colwell. 1982. Scanning electron microscope observation of the swarming phenomenon of Vibrio parahaemolyticus. J. Bacteriol. 150:956-959.[Abstract/Free Full Text]
4
4. Bhattacharyya, F. K. 1975. Vibrio cholerae flagellar antigens: a serodiagnostic test, functional implications of H-reactivity and taxonomic importance of cross-reactions within the Vibrio genus. Med. Microbiol. Immunol. 162:29-41.[CrossRef][Medline]
5
5. Bhattacharyya, F. K., and S. Mukerjee. 1974. Serological analysis of the flagellar antigen or H agglutinating antigens of cholera and NAG vibrios. Ann. Microbiol. (Paris) 125:167-181.[Medline]
6
6. Blackstone, G. M., J. L. Nordstrom, M. C. Vickery, M. D. Bowen, R. F. Meyer, and A. DePaola. 2003. Detection of pathogenic Vibrio parahaemolyticus in oyster enrichments by real time PCR. J. Microbiol. Methods 53:149-155.[CrossRef][Medline]
7
7. Brasher, C. W., A. DePaola, D. D. Jones, and A. K. Bej. 1998. Detection of microbial pathogens in shellfish with multiplex PCR. Curr. Microbiol. 37:101-107.[CrossRef][Medline]
8
8. Centers for Disease Control and Prevention. 1998. Outbreak of Vibrio parahaemolyticus infections associated with eating raw oysters-Pacific Northwest, 1997. MMWR Morb. Mortal. Wkly. Rep. 47:457-462.[Medline]
9
9. Centers for Disease Control and Prevention. 1999. Outbreak of Vibrio parahaemolyticus infection associated with eating raw oysters and clams harvested from Long Island Sound—Connecticut, New Jersey, and New York, 1998. Morb. Mortal. Wkly. Rep. 48:48-51.[Medline]
10
10. Chen, D., P. J. Hanna, K. Altman, A. Smith, P. Moon, and L. S. Hammond. 1992. Development of monoclonal antibodies that identify Vibrio species commonly isolated from infections of humans, fish, and shellfish. Appl. Environ. Microbiol. 58:3694-3700.[Abstract/Free Full Text]
11
11. Cook, D. W., P. O'Leary, J. C. Hunsucker, E. M. Sloan, J. C. Bowers, R. J. Blodgett, and A. DePaola. 2002. Vibrio vulnificus and Vibrio parahaemolyticus in U.S. retail shell oysters: a national survey from June 1998 to July 1999. J. Food Prot. 65:79-87.[Medline]
12
12. Cook, D. W., A. DePaola, and S. A. McCarthy. 2000. Direct plating procedure for the enumeration of total and pathogenic Vibrio parahaemolyticus in oyster meats. Office of Seafood, Gulf Coast Laboratory, U.S. Food and Drug Administration, Dauphin Island, AL.
13
13. Corona-Barrera, E., D. G. E. Smith, T. La, D. J. Hampson, and J. R. Thomson. 2004. Immunomagnetic separation of the intestinal spirochaetes Brachyspira pilosicoli and Brachyspira hyodysenteriae from porcine faeces. J. Med. Microbiol. 53:301-307.[Abstract/Free Full Text]
14
14. Daniels, N. A., L. MacKinnon, R. Bishop, S. Altekruse, B. Ray, R. M. Hammond, S. Thompson, S. Wilson, N. H. Bean, P. M. Griffin, and L. Slutsker. 2000. Vibrio parahaemolyticus infections in the United States, 1973-1998. J. Infect. Dis. 181:1661-1666.[CrossRef][Medline]
15
15. DePaola, A., J. L. Nordstrom, J. C. Bowers, J. G. Wells, and D. W. Cook. 2003. Seasonal abundance of total and pathogenic Vibrio parahaemolyticus in Alabama oysters. Appl. Environ. Microbiol. 69:1521-1526.[Abstract/Free Full Text]
16
16. DePaola, A., C. A. Kaysner, J. Bowers, and D. W. Cook. 2000. Environmental investigations of Vibrio parahaemolyticus in oysters after outbreaks in Washington, Texas, and New York (1997 and 1998). Appl. Environ. Microbiol. 66:4649-4654.[Abstract/Free Full Text]
17
17. Elliot, E. L., C. A. Kaysner, and M. L. Tamplin. 1992. Vibrio cholerae, V. parahaemolyticus, V. vulnificus, and other Vibrio spp., p. 111-140. In Bacteriological analytical manual, 7th ed. Association of Official Analytical Chemists, Arlington, VA.
18
18. Enroth, H., and L. Engstrand. 1995. Immunomagnetic separation and PCR for detection of Helicobacter pylori in water and stool specimens. J. Clin. Microbiol. 33:2162-2165.[Abstract]
19
19. Fluit, A. C., R. Torensma, M. J. C. Visser, C. J. M. Aarsman, M. J. J. P. Poppelier, B. H. I. Keller, P. Klapwijk, and J. Verhoef. 1993. Detection of Listeria monocytogenes in cheese with the magnetic immuno-polymerase chain reaction assay. Appl. Environ. Microbiol. 59:1289-1293.[Abstract/Free Full Text]
20
20. Gooch, J. A., A. DePaola, C. A. Kaysner, and D. L. Marshall. 2001. Evaluation of two direct plating methods using nonradioactive probes for enumeration of Vibrio parahaemolyticus in oysters. Appl. Environ. Microbiol. 67:721-724.[Abstract/Free Full Text]
21
21. Grant, I. R., H. J. Ball, and M. T. Rowe. 1998. Isolation of Mycobacterium paratuberculosis from milk by immunomagnetic separation. Appl. Environ. Microbiol. 64:3153-3158.[Abstract/Free Full Text]
22
22. Hongprayoon, R. 1993. The role of flagellar proteins in adhesion of Vibrio parahaemolyticus, and isolation of the corresponding gene(s). Ph.D. thesis. Louisiana State University, Baton Rouge.
23
23. Kaufman, G. E., G. M. Blackstone, M. C. Vickery, A. K. Bej, J. Bowers, M. D. Bowen, R. F. Meyer, and A. DePaola. 2004. Real-time PCR quantification of Vibrio parahaemolyticus in oysters using an alternative matrix. J. Food Prot. 67:2424-2429.[Medline]
24
24. Kaufman, G. E., A. K. Bej, J. Bowers, and A. DePaola. 2003. Oyster-to-oyster variability in levels of Vibrio parahaemolyticus. J. Food Prot. 66:125-129.[Medline]
25
25. Kaysner, C. A., and A. DePaola. 1998. Vibrio cholerae, V. parahaemolyticus, V. vulnificus, and other Vibrio spp., chapter 9. In Bacteriological analytical manual, 8th ed., revision A. Association of Official Analytical Chemists, Arlington, VA. http://www.cfsan.fda.gov/ebam/bam-9.html.
26
26. Khare, S., T. A. Ficht, R. L. Santos, J. Romano, A. R. Ficht, S. Zhang, I. R. Grant, M. Libal, D. Hunter, and L. G. Adams. 2004. Rapid and sensitive detection of Mycobacterium avium subsp. paratuberculosis in bovine milk and feces by a combination of immunomagnetic bead separation-conventional PCR and real-time PCR. J. Clin. Microbiol. 42:1075-1081.[Abstract/Free Full Text]
27
27. Kim, Y., and L. L. McCarter. 2000. Analysis of the polar flagellar gene system of Vibrio parahaemolyticus. J. Bacteriol. 182:3693-3704.[Abstract/Free Full Text]
28
28. Levine, W. C., P. M. Griffin, et al. 1993. Vibrio infections on the Gulf Coast: results of first year of regional surveillance. J. Infect. Dis. 167:470-483.
29
29. McCarter, L. L. 1995. Genetic and molecular characterization of the polar flagellum of Vibrio parahaemolyticus. J. Bacteriol. 177:1595-1609.[Abstract/Free Full Text]
30
30. Olsvik, O., T. Popovic, E. Skjerve, K. S. Cudjoe, E. Hornes, J. Ugelstad, and M. Uhlen. 1994. Magnetic separation techniques in diagnostic microbiology. Clin. Microbiol. Rev. 7:43-54.[Abstract/Free Full Text]
31
31. Roberts, B., and R. Hirst. 1997. Immunomagnetic separation and PCR for detection of Mycobacterium ulcerans. J. Clin. Microbiol. 35:2709-2711.[Abstract]
32
32. Sakazaki, R., R. Iwanami, and K. Tamura. 1968. Studies on the enteropathogenic facultatively halophilic bacteria Vibrio parahaemolyticus. II. Serological characteristics. Jpn. J. Med. Sci. Biol. 21:313-324.[Medline]
33
33. Shinoda, S., T. Senoh, K. Asano, N. Nakahara, and B. Ono. 1980. Differences between surface antigenic determinants of polar monotrichous flagella of Vibrio parahaemolyticus and of related species. Microbiol. Immunol. 24:409-418.[Medline]
34
34. Shinoda, S., N. Nakahara, and B. Ono. 1979. Behavior of a surface antigenic determinant of lateral flagella of Vibrio parahaemolyticus. Infect. Immun. 26:322-327.[Abstract/Free Full Text]
35
35. Shinoda, S., and N. Nakahara. 1977. Antigenicity of lateral flagella of Vibrio parahaemolyticus: existence of two distinct antigenic determinants in a flagellum. Infect. Immun. 17:235-237.[Abstract/Free Full Text]
36
36. Shinoda, S., and K. Okamoto. 1977. Formation and function of Vibrio parahaemolyticus lateral flagella. J. Bacteriol. 129:1266-1271.[Abstract/Free Full Text]
37
37. Shinoda, S., T. Honda, Y. Takeda, and T. Miwatani. 1974. Antigenic difference between polar monotrichous and peritrichous flagella of Vibrio parahaemolyticus. J. Bacteriol. 120:923-928.[Abstract/Free Full Text]
38
38. Shinoda, S., T. Miwatani, and T. Fujino. 1970. Antigens of Vibrio parahaemolyticus. II. Existence of two different subunits in the flagella of Vibrio parahaemolyticus and their characterization. Biken J. 13:241-247.[Medline]
39
39. Shinoda, S., T. Miwatani, and T. Fujino. 1970. Antigens of Vibrio parahaemolyticus. II. Some physiochemical properties. Biken J. 14:75-76.
40
40. Simonson, J., and R. J. Siebeling. 1986. Rapid serological identification of Vibrio vulnificus by anti-H coagglutination. Appl. Environ. Microbiol. 52:1299-1304.[Abstract/Free Full Text]
41
41. Simonson, J. G., and R. J. Siebeling. 1988. Coagglutination of Vibrio cholerae, Vibrio mimicus, and Vibrio vulnificus with anti-flagellar monoclonal antibody. J. Clin. Microbiol. 26:1962-1966.[Abstract/Free Full Text]
42
42. Skjerve, E., L. M. Rorvik, and O. Olsvik. 1990. Detection of Listeria monocytogenes in foods by immunomagnetic separation. Appl. Environ. Microbiol. 56:3478-3481.[Abstract/Free Full Text]
43
43. Su, Y. C., and C. Liu. 2007. Vibrio parahaemolyticus: a concern of seafood safety. Food Microbiol. 24:549-558.[CrossRef][Medline]
44
44. Tassin, M. G., R. J. Siebeling, and A. D. Larson. 1984. H-antigen relationships among several Vibrio species, p. 73-82. In R. R. Colwell (ed.), Vibrios in the environment. John Wiley & Sons, Inc., New York, NY.
45
45. Tassin, M. G., R. J. Siebeling, N. C. Roberts, and A. D. Larson. 1983. Presumptive identification of Vibrio species with H antiserum. J. Clin. Microbiol. 18:400-407.[Abstract/Free Full Text]
46
46. Terada, Y. 1968. Serological studies of Vibrio parahaemolyticus. II. Flagellar antigens. Jpn. J. Bacteriol. 23:767-771.
47
47. Thomashow, L. S., and S. C. Rittenberg. 1985. Isolation and composition of sheathed flagella from Bdellovibrio bacteriovorus 109J. J. Bacteriol. 163:1047-1054.[Abstract/Free Full Text]
48
48. Yang, G. C. H., G. D. Schrank, and B. A. Freeman. 1977. Purification of flagellar cores of Vibrio cholerae. J. Bacteriol. 129:1121-1128.[Abstract/Free Full Text]

---

Clinical and Vaccine Immunology, October 2008, p. 1541-1546, Vol. 15, No. 10
1071-412X/08/$08.00+0     doi:10.1128/CVI.00141-08
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via Google Scholar Google Scholar Articles by Datta, S. Articles by Simonson, J. G. Search for Related Content PubMed PubMed Citation Articles by Datta, S. Articles by Simonson, J. G.