Abstract
Because biochemical testing and 16S rRNA sequence analysis have proven inadequate for the differentiation of Vibrio parahaemolyticus from closely related species, we employed the gyrase B gene (gyrB) as a molecular diagnostic probe. The gyrB genes of V. parahaemolyticus and closely related Vibrio alginolyticus were cloned and sequenced. Oligonucleotide PCR primers were designed for the amplification of a 285-bp fragment from within gyrB specific for V. parahaemolyticus. These primers recognized 117 of 117 reference and wild-type V. parahaemolyticus strains, whereas amplification did not occur when 90 strains of 37 other Vibrio species or 60 strains representing 34 different nonvibrio species were tested. In 100-μl PCR mixtures, the lower detection limits were 5 CFU for live cells and 4 pg for purified DNA. The possible application of gyrB primers for the routine identification of V. parahaemolyticus in food was examined. We developed and tested a procedure for the specific detection of the target organism in shrimp consisting of an 18-h preenrichment followed by PCR amplification of the 285-bp V. parahaemolyticus-specific fragment. This method enabled us to detect an initial inoculum of 1.5 CFU of V. parahaemolyticus cells per g of shrimp homogenate. By this approach, we were able to detect V. parahaemolyticus in all of 27 shrimp samples artificially inoculated with this bacterium. We present here a rapid, reliable, and sensitive protocol for the detection of V. parahaemolyticus in shrimp.
Vibrio parahaemolyticus is considered to be the causative agent in 50 to 70% of all cases of diarrhea associated with the consumption of fishery products in the summer months (9, 12, 26). The common method for the detection of V. parahaemolyticus is a culture-based procedure which employs enrichment in liquid media and the subsequent isolation of colonies on selective plating media (6). Unfortunately, a number of other Vibrio species are taxonomically similar to V. parahaemolyticus, necessitating the utilization of additional biochemical tests for reliable identification (30). Because the conventional detection method for V. parahaemolyticus requires 3 days and positive identification requires 7 days (19), a more rapid and sensitive 8-h detection assay based on the measurement of trypsin-like activity was developed by Miyamoto et al. (10, 11). Subsequently, our group demonstrated that the intracellular trypsin-like activity was not specific to V. parahaemolyticus and did not differentiate this pathogen from closely related Vibrio species, such as V. alginolyticus and V. harveyi (25). Unfortunately, 16S rRNA sequences revealed 99.7% homology between V. parahaemolyticus and V. alginolyticus, and thus 16S rRNA was unable to differentiate between V. parahaemolyticus and other vibrios (18).
To further complicate matters, there are several phenotypes (2), serotypes (25), and toxin-producing strains of V. parahaemolyticus (1, 5, 21), and specific hemolysin probes have failed to detect all of these types (13, 14). Several of the nondetected isolates were toxin producers, rendering the method less useful with regard to identifying potentially contaminated food.
Yamamoto and Harayama (32) suggested that genes that are not spread horizontally among different bacterial species may be used to trace the evolutionary record of host bacteria. Assuming that the average substitution rate for 16S rRNA is 1% per 50 million years and that the rate for synonymous sites of protein-coding DNA is 0.7 to 0.8% per million years (15), Yamamoto and Harayama (31) have proposed the use of the gyrB gene as a molecular taxonomic marker for bacterial species. The gyrB gene that encodes the B subunit protein of DNA gyrase (topoisomerase type II) is a single-copy gene and is essential for DNA replication; it also has conserved regions for the development of PCR primers. Because no universal probe was available to differentiate V. parahaemolyticus from related species, we studied the possibility of using the gyrB gene as a highly specific probe (31, 32). In this report, a 1.2-kb fragment of the gyrB gene of V. parahaemolyticus and V. alginolyticus was amplified, cloned, and sequenced. We have designed and employed suitable PCR primers that amplify only the gyrB fragment of V. parahaemolyticus to specifically identify this pathogen irrespective of its phenotypes, serotypes, and virulence status. Application of gyrB primers for the detection of V. parahaemolyticus directly from food with a PCR protocol is also described.
MATERIALS AND METHODS
Bacterial strains.
The microorganisms included in this study were purchased from various culture collections (Table 1). Gift strains from Sumio Shinoda, Okayama University, and wild types isolated from various aquatic environments and foods were also included. These strains include various phenotypes, serotypes, and toxin-producing isolates of V. parahaemolyticus. In addition, care was taken to include all available Vibrio and closely related species (2). All microorganisms were grown in either Marine broth (Difco Laboratories, Detroit, Mich.) or alkaline peptone water (APW [Nissui, Tokyo]) at 35°C for 24 h prior to use. For the isolation of wild-type V. parahaemolyticus, green colonies appearing on thiosulfate citrate bile salt agar (TCBS [Eiken Co., Tokyo, Japan]) were picked and biochemically characterized as described elsewhere (25).
TABLE 1.
Specificity of VP-1 and VP-2r primers in detection of V. parahaemolyticus
| Bacterial taxon | Sourcea | Strain no. | No. of strains tested | No. of strains showing gyrB ampliconsb of:
|
|
|---|---|---|---|---|---|
| 1.2 kb | 285 bp | ||||
| Aeromonas hydrophila | ATCC | 19570 | 1 | 1 | 0 |
| Alteromonas macleodii | ATCC | 27126T | 1 | 1 | 0 |
| Deleya aquamarina | JCM | 27128T | 1 | 1 | 0 |
| Deleya cupida | JCM | 27124T | 1 | 1 | 0 |
| Deleya halophilica | Unknown | 1 | 1 | 0 | |
| Deleya marina | JCM | 25374T | 1 | 1 | 0 |
| Deleya pacifica | JCM | 27122T | 1 | 1 | 0 |
| Deleya venusta | JCM | 27125T | 1 | 1 | 0 |
| Escherichia coli | ATCC | 25922T | 1 | 1 | 0 |
| Listonella anguillarum | ATCC | 19264T | 1 | 1 | 0 |
| Listonella pelagia | ATCC | 25916T | 1 | 1 | 0 |
| Marinomonas communis | ATCC | 27118T | 1 | 1 | 0 |
| Marinomonas vaga | ATCC | 27119T | 1 | 1 | 0 |
| Photobacterium damsela | ATCC | 33539T | 1 | 1 | 0 |
| Pseudoalteromonas atlantica | ATCC | 19262T | 1 | 1 | 0 |
| Pseudoalteromonas aurantia | ATCC | 33046T | 1 | 1 | 0 |
| Pseudoalteromonas carrogeenovara | ATCC | 43555T | 1 | 1 | 0 |
| Pseudoalteromonas citrea | ATCC | 29719T | 1 | 1 | 0 |
| Pseudoalteromonas espejiana | ATCC | 29659T | 1 | 1 | 0 |
| Pseudoalteromonas haloplanktis | ATCC | 14393T | 1 | 1 | 0 |
| Pseudoalteromonas luteoviolacea | ATCC | 33492T | 1 | 1 | 0 |
| Pseudoalteromonas nigrifaciens | ATCC | 19375T | 1 | 1 | 0 |
| Pseudoalteromonas rubra | ATCC | 29570T | 1 | 1 | 0 |
| Pseudoalteromonas sp. | Unknown | 7 | 7 | 0 | |
| Pseudoalteromonas tetraodonis | Unknown | 1 | 1 | 0 | |
| Pseudoalteromonas undina | ATCC | 29660T | 1 | 1 | 0 |
| Pseudomonas aeruginosa | IFO | 12689 | 1 | 1 | 0 |
| Pseudomonas fluorescens | IFO | 14160 | 1 | 1 | 0 |
| Salmonella typhimurium | ATCC | 13311T | 1 | 1 | 0 |
| Shewanella algae | ATCC | 51192Tc | 2 | 2 | 0 |
| Shewanella benthica | ATCC | 43992T | 1 | 1 | 0 |
| Shewanella hanedai | ATCC | 33224T | 1 | 1 | 0 |
| Shewanella putrefaciens | ATCC | 8071Tc | 20 | 20 | 0 |
| Shigella dysenteriae | ATCC | 13313T | 1 | 1 | 0 |
| Shigella sonnei | ATCC | 29930T | 1 | 1 | 0 |
| Staphylococcus aureus | ATCC | 12600T | 1 | 1 | 0 |
| Vibrio aestuarianus | ATCC | 35048T | 1 | 1 | 0 |
| Vibrio albensis | ATCC | 14547T | 1 | 1 | 0 |
| Vibrio alginolyticus | ATCC | 17749Tc | 20 | 20 | 0 |
| Vibrio campbellii | ATCC | 25920T | 1 | 1 | 0 |
| Vibrio carchariae | ATCC | 35084T | 1 | 1 | 0 |
| Vibrio cholerae non-O1 | Unknown | 2 | 2 | 0 | |
| Vibrio cincinnatiensis | ATCC | 35912T | 1 | 1 | 0 |
| Vibrio costicola | ATCC | 33508T | 1 | 1 | 0 |
| Vibrio diazotrophicus | ATCC | 33466T | 1 | 1 | 0 |
| Vibrio fischeri | ATCC | 7744T | 1 | 1 | 0 |
| Vibrio fluvialis | JCM | 3752 | 1 | 1 | 0 |
| Vibrio furnissii | ATCC | 35016T | 1 | 1 | 0 |
| Vibrio gazogenes | ATCC | 29988T | 1 | 1 | 0 |
| Vibrio harveyi | ATCC | 14126Tc | 6 | 6 | 0 |
| Vibrio hollisae | CDC | 75-80 | 1 | 1 | 0 |
| Vibrio ichthyoenteri | IFO | 15847 | 1 | 1 | 0 |
| Vibrio logei | ATCC | 29985T | 1 | 1 | 0 |
| Vibrio marinus | ATCC | 15381T | 1 | 1 | 0 |
| Vibrio mediterranei | ATCC | 43341T | 1 | 1 | 0 |
| Vibrio metschnikovii | ATCC | 7708T | 1 | 1 | 0 |
| Vibrio mimicus | Unknown | 1 | 1 | 0 | |
| Vibrio mytili | NCIMB | 13275 | 1 | 1 | 0 |
| Vibrio natriegens | ATCC | 14048T | 1 | 1 | 0 |
| Vibrio navarrensis | NCIMB | 13120 | 1 | 1 | 0 |
| Vibrio nereis | ATCC | 25917T | 1 | 1 | 0 |
| Vibrio nigripulchritudo | ATCC | 27043T | 1 | 1 | 0 |
| Vibrio ordalii | ATCC | 33509T | 1 | 1 | 0 |
| Vibrio orientalis | ATCC | 33934T | 1 | 1 | 0 |
| Vibrio proteolyticus | ATCC | 15338T | 1 | 1 | 0 |
| Vibrio salmonicida | ATCC | 43839T | 1 | 1 | 0 |
| Vibrio splendidus | ATCC | 33125T | 1 | 1 | 0 |
| Vibrio spp. | Food isolatesd | 28 | 28 | 0 | |
| Vibrio tubiashii | ATCC | 19109T | 1 | 1 | 0 |
| Vibrio vulnificus | ATCC | 2046Tc | 2 | 2 | 0 |
ATCC, American Type Culture Collection; CDC, Centers for Disease Control and Prevention; IFO, Institute of Fermentation—Osaka; JCM, Japan Collection of Microorganisms; NCIMB, National Collections of Industrial and Marine Bacteria, U.K.
gyrB gene length is 1.2 kb; the V. parahaemolyticus-specific fragment is 289 bp in length.
Isolates, including the type strain of the species from various sources, were tested.
Isolated from various seafoods (25).
DNA isolation.
Chromosomal DNA from overnight cultures was purified by phenol-chloroform extraction and ethanol precipitation (20). Dried DNA pellets were dissolved in Tris-EDTA (TE) buffer (pH 7.5) and used as DNA templates for PCR when applicable. DNA purity was checked by agarose gel electrophoresis, and the DNA concentration was measured with a spectrophotometer (20).
Cloning and sequencing of the gyrB gene.
Primers (UP-1 and UP-2r) within the known DNA sequence (31) were added to the PCR mixture at a concentration of 1 μM, and the solution was subjected to 30 cycles of PCR (denaturing, 1 min at 94°C; annealing, 1 min at 60°C; extension, 2 min at 72°C). The amplified gyrB fragments from V. parahaemolyticus ATCC 17802 and V. alginolyticus ATCC 17749 were cloned in pGEM ZF+ (Promega, Madison, Wis.) by conventional recombinant methods (20). Expansion of the probes was carried out as documented previously (20). After ligation of the PCR fragments into the vector, E. coli cells were transformed with the ligation mixture by calcium chloride-mediated transformation. After transformation, the transformants were cultured under conditions which promote growth. Plasmids were recovered from a transformant by lysis and purification by the alkaline method. The purified intact plasmids were then utilized as probes. The identity of the fragment was verified by sequencing from both ends by the dye deoxy chain termination method with a Sequenase DNA sequencing kit (U.S. Biochemical Corporation, Cleveland, Ohio) and with an ABI 373A automatic sequencer as described by the manufacturer (Perkin-Elmer-Applied Biosystems, Foster City, Calif.). DNA sequences were determined from both strands by extension from vector-specific (T7 and SP6 primers from pGEM ZF+) priming sites and by primer walking.
V. parahaemolyticus-specific primers.
Oligonucleotide primers (Table 2) based on the nucleotide sequence data of the V. parahaemolyticus gyrB gene (Fig. 1) were synthesized (Beckman, Fullerton, Calif.) according to the manufacturer’s instructions.
TABLE 2.
Primers specific for the amplification of V. parahaemolyticus
| Primer | Oligonucleotide sequence | Length (bases) | Positiona | Orientation |
|---|---|---|---|---|
| VP-1 | CGG CGT GGG TGT TTC GGT AGT | 21 | 75–95 | Sense |
| VP-2r | TCC GCT TCG CGC TCA TCA ATA | 21 | 339–359 | Antisense |
Positions correspond to the gyrB nucleotide sequence of E. coli (ECGYRBF in the EMBL database).
FIG. 1.
Nucleotide sequence of gyrB of V. parahaemolyticus ATCC 17802 and V. alginolyticus ATCC 17749. Nucleotides identical to those of V. parahaemolyticus are indicated with dots.
PCR assay.
When whole bacterial cells were used as templates for PCR in the absence of a DNA extraction step, freshly grown cells from agar plates or centrifuged and washed (phosphate-buffered saline [PBS; 0.1 M, pH 7.5] containing 2% [wt/vol] NaCl) cells from a liquid culture were used.
PCR assays were performed with a DNA Thermal Cycler (Perkin-Elmer Corp., Foster City, Calif.). The 1.2-kb gyrB gene was amplified as described elsewhere (31). The amplification of a V. parahaemolyticus-specific 285-bp fragment was performed by using PCR for 30 cycles, each consisting of 1 min at 94°C, 1.5 min at 58°C, 2.5 min at 72°C, and a final extension step at 72°C for 7 min. After DNA amplification, the 285-bp amplicon was analyzed by submarine gel electrophoresis, stained, and visualized under UV illumination (31). Suitable molecular size markers were included in each gel.
PCR assay sensitivity for the detection of artificially contaminated V. parahaemolyticus in food.
A 25-g sample of shrimp (Indonesia-White) in triplicate was homogenized for 1 min with a homogenizer (model SH-001; Elmex, Tokyo, Japan) in 225 ml of APW to produce a uniform food homogenate for all experiments. V. parahaemolyticus ATCC 17802 and V. alginolyticus ATCC 17749 were grown in APW overnight at 37°C, serially diluted with the food homogenate as the diluent to final concentrations ranging from 0 to 109 CFU per g. These artificially contaminated food homogenate microcosms (10 ml each) were incubated at 37°C with shaking at 140 rpm. Subsampling (1 ml) was carried out after 0-, 6-, and 18-h (overnight) incubations; cells were centrifuged (4°C, 10,000 × g for 10 min) and resuspended in 1 ml of sterile PBS. A 10-μl sample suspension was used as a template for the PCR assay without extraction of DNA.
Suitable controls such as buffer, media, PCR mixtures, and V. parahaemolyticus DNA were employed to check any false-positive or false-negative reactions. Appropriate dilutions of seafood homogenates prepared at various intervals in APW were spread plated onto Marine agar (Difco) for total viable counts and onto TCBS agar for the enumeration of total vibrios and the V. parahaemolyticus population.
The experiment was repeated by inoculation of 1.5 CFU of V. parahaemolyticus in 1 g of shrimp homogenate as described above. Three samples of the following nine varieties of shrimps were tested: Indonesia-White (King), Indonesia-White (Bintni King), Australia-Banana, Indonesia-Black Tiger, Thai-Black Tiger, Indonesia-Indiva Pink Tail, Indonesia-Indiva Blue Tail, Mexico-Brown, and Surinam-Pink. Shrimp samples were received as frozen from our food processing centers.
Nucleotide sequence accession number.
The nucleotide sequence data reported in this paper will appear in the GenBank, EMBL, and DDBJ nucleotide sequence databases under the following accession numbers: V. parahaemolyticus, AF007287; V. alginolyticus, AF007288; V. harveyi, AF007289; Vibrio mytili, AF007290; Vibrio natriegens, AF007291; Listonella pelagia, AF007292; and Pseudoalteromonas undina, AF007293.
RESULTS
gyrB sequence of V. parahaemolyticus.
Complete sequences of the 1,258-bp gyrB fragments of V. parahaemolyticus ATCC 17802 and V. alginolyticus ATCC 17749 were determined and aligned (Fig. 1). The frequency of base substitutions in the published sequence of the 16S rRNA was lower than that in gyrB. For example, between the sequences of V. parahaemolyticus and V. alginolyticus, 166 base substitutions among 1,258 bp were observed in gyrB, while only 5 base substitutions among 1,451 bp were observed in 16S rRNA. The homology of the gyrB sequences between V. parahaemolyticus and V. alginolyticus was 86.8%, versus 99.7% homology for the 16S rRNA sequence. Figure 2 shows the alignment of amino acid sequences for the gyrB proteins translated from the nucleotide sequences. For V. parahaemolyticus and V. alginolyticus, only 17 of the 166 substitutions caused amino acid substitutions. Amino acid sequence homology between the gyrase subunit B proteins of V. parahaemolyticus and V. alginolyticus was 92.8%.
FIG. 2.
Amino acid sequence alignment of the gyrB products from V. parahaemolyticus ATCC 17802 and V. alginolyticus ATCC 17749. Amino acids identical to those of V. parahaemolyticus are indicated with dots.
Designing V. parahaemolyticus-specific PCR primers.
A species-specific primer set of 21 bp each was designed to specifically detect and differentiate V. parahaemolyticus from other bacteria. A forward primer with nucleotide positions 75 to 95 (VP-1) and an antisense primer with positions 321 to 341 were synthesized. When these primers were used to generate 267-bp PCR products, V. parahaemolyticus could be differentiated from V. alginolyticus. However, V. harveyi NCIMB 1896, V. natriegens ATCC 14048, V. mytili NCIMB 13275, Listonella pelagia ATCC 25916, and Pseudoalteromonas undina ATCC 29660 also showed positive amplification of the same 267-bp fragment. The gyrB gene of these five strains was partially sequenced with the UP-1s primer (31). By comparing the nucleotide sequences of these five strains to those of V. parahaemolyticus and V. alginolyticus, a 21-bp variable region located between nucleotide positions 339 and 359 of the gyrB fragment of V. parahaemolyticus was synthesized as an antisense primer (VP-2r). The nucleotide sequence of each primer is presented in Table 2. The primer pair VP-1 and VP-2r were predicted to prime amplification products of 285 bp when the gyrB sequence was utilized as a target fragment.
Specificity of PCR primers in the detection of V. parahaemolyticus.
A total of 267 strains comprising 72 different species were screened for both the 1.2-kb gyrB gene and the V. parahaemolyticus-specific 285-bp fragments. The results are presented in Table 1. A specific band of 285 bp was noticed for all V. parahaemolyticus strains but no other bacterial species. However, PCR amplification with primer set UP-1 and UP-2r revealed a 1.2-kb gyrB fragment in all strains examined, thus ascertaining the presence of the DNA gyrase B subunit. Thus, the primers developed and described here (VP-1 and VP-2r) are specific to V. parahaemolyticus and could be applied to the molecular diagnosis of this bacterium. It should be noted that 20 V. alginolyticus strains tested failed to show any V. parahaemolyticus-specific fragment. V. parahaemolyticus strains isolated from various environments, food, and clinical sources comprising various phenotypes, serotypes, and toxigenic properties were tested with the PCR assay, and all 117 strains exhibited the 285-bp fragment when tested under the PCR conditions described here.
Sensitivity of the VP-1 and VP-2r PCR primers in the detection of V. parahaemolyticus.
To evaluate the sensitivity of our PCR assay, a dilution series of genomic DNA from V. parahaemolyticus ATCC 17802 was prepared with TE buffer and used as the templates for PCR amplification. Samples (100 μl) containing 4 pg of genomic DNA were successfully detected after amplification with primer pair VP-1 and VP-2r (Fig. 3). A dilution series of freshly cultured V. parahaemolyticus ATCC 17802 cells showed that the primer set employed in this study amplified the V. parahaemolyticus-specific 285-bp fragment when 5 CFU of bacterial cells per reaction tube (100 μl) was used (Fig. 4).
FIG. 3.
Sensitivity of VP-1 and VP-2r PCR primers for the amplification of the V. parahaemolyticus-specific amplicon at various DNA concentrations. DNAs were extracted from overnight cultures in APW by phenol-chloroform extraction and ethanol precipitation. DNA concentrations were measured with a spectrophotometer, and DNA was serially diluted in TE buffer to obtain the appropriate concentrations. Lanes contained V. parahaemolyticus DNA (unless otherwise noted): M, 100-bp DNA ladder; 1, 4.3 ng; 2, 430 pg; 3, 43 pg; 4, 4 pg; 5, 430 fg; 6, V. alginolyticus DNA (10 ng).
FIG. 4.
Sensitivity of VP-1 and VP-2r PCR primers for the amplification of the V. parahaemolyticus-specific amplicon at various bacterial concentrations. Bacterial cells were grown in APW for 18 h at 37°C and serially diluted in PBS containing 2% NaCl. Appropriate dilutions were spread plated on marine agar, and bacterial counts were enumerated after 18 h of incubation at 37°C. Lanes contained V. parahaemolyticus (unless otherwise noted): M, 100-bp DNA ladder; 1, 5.2 × 103 CFU per reaction tube; 2, 5.2 × 102 CFU per reaction tube; 3, 5.2 × 101 CFU per reaction tube; 4, 5.2 CFU per reaction tube; 5, PCR mixture control without any added bacterial cells; 6, V. alginolyticus bacterial cells (7.9 × 103 CFU per reaction tube).
Detection of V. parahaemolyticus in artificially contaminated shrimp.
The sensitivity of the PCR assay for detecting V. parahaemolyticus in artificially contaminated shrimp is presented in Fig. 5 and Table 3. The absence of a V. parahaemolyticus-like organism in the test sample was confirmed by both the conventional APW enrichment method (30) and by PCR assay (Fig. 5, lane 1). When the food homogenate was incubated for 18 h in APW at 37°C, an initial inoculum of 1.5 CFU of V. parahaemolyticus per g of food homogenate amplified the desired PCR product (Fig. 5, lane 2). In a sample drawn at time zero, 1.5 × 105 CFU of V. parahaemolyticus per g of shrimp homogenate failed to yield any PCR product (Fig. 5, lane 3). When the experiment was repeated by inoculation of 1.5 CFU of V. parahaemolyticus in 1 g of shrimp homogenate, after an overnight incubation, all 27 samples of nine varieties of shrimps showed a V. parahaemolyticus-specific amplicon.
FIG. 5.
Detection of V. parahaemolyticus in artificially contaminated shrimp by the VP-1 and VP-2r PCR primers. V. parahaemolyticus cells grown overnight in APW were serially diluted in shrimp homogenate (see details in Materials and Methods) to obtain the appropriate dilutions. Total microflora of shrimp were counted in marine agar (48 h), and the V. parahaemolyticus population was counted in TCBS agar (18 h) after incubation at 37°C. Lanes: M, 100-bp DNA ladder; 1, shrimp homogenate not spiked with V. parahaemolyticus cells; 2, initial inoculum of 1.5 CFU of V. parahaemolyticus per g added to shrimp homogenate and incubated for 18 h at 37°C; 3, initial inoculum of 1.5 × 105 CFU of V. parahaemolyticus per g added to shrimp homogenate sampled at zero hour; 4, 1.5 CFU of V. parahaemolyticus cells prepared in PCR mixture.
TABLE 3.
Influence of competitive microflora and preenrichment incubationa time on amplification of PCR product specific to V. parahaemolyticus
| Initial inoculum (CFU g of food homogenate−1) | PCR band amplification after preenrichment incubation for (h)b:
|
V. parahaemolyticus/food microflora population (CFU/g) after preenrichment incubation for (h):
|
||||
|---|---|---|---|---|---|---|
| 0 | 6 | 18 | 0 | 6 | 18 | |
| 1.5 × 105 | − | + | + | 105/102 | 107/103 | 108/104 |
| 1.5 × 104 | − | + | + | 104/102 | 105/103 | 108/107 |
| 1.5 × 103 | − | − | + | 103/102 | 104/102 | 108/108 |
| 1.5 × 102 | − | − | + | 102/102 | 103/102 | 107/108 |
| 1.5 × 101 | − | − | + | 101/102 | 102/103 | 107/108 |
| 1.5 | − | − | + | 1/102 | 10/103 | 106/108 |
| 0 | − | − | − | 0/102 | 0/105 | 0/109 |
Incubation in APW.
−, 258-bp amplified product not seen; +, 285-bp amplified product seen.
Influence of competitive food microflora on the sensitivity of PCR.
A range of 10 to 90 CFU of total vibrio population per g was recorded in the shrimp samples tested. The distribution of various species of vibrios varied between sample. Ten strains from each sample, totalling 280 strains, were isolated, purified, and identified with standard biochemical tests (25, 30). V. parahaemolyticus was not present in any of the shrimp tested. V. harveyi was predominant irrespective of the sample. V. alginolyticus was isolated frequently, along with Vibrio splendidus biovar I. In addition to these species, Vibrio campbelli, Vibrio metschnikovii, Vibrio vulnificus, Vibrio marinus, Vibrio pelagia, Vibrio cholerae non-O1, and Vibrio fischeri were isolated. The majority of these species mimic V. parahaemolyticus and were found to be very difficult to differentiate in agar plates.
Changes in the population densities of V. parahaemolyticus and other food microflora are depicted in Table 3. The results presented here indicate that V. parahaemolyticus detection was enhanced when the food homogenate was subjected to overnight incubation. When the 6-h APW-enriched food homogenate was assayed, a minimum of 1.5 × 104 CFU of V. parahaemolyticus cells per g was required to amplify the specific amplicon (faint bands), and the ratio of V. parahaemolyticus to other competitive microflora in the sample was in the range of 102:1 (Table 3). However, an 18-h enrichment in APW generated the PCR band clearly, even when only 1.5 CFU of V. parahaemolyticus was used as the initial inoculum, and its population and other food microflora were present at a ratio of 1:102 (Table 3). Successful PCR amplification after an overnight incubation was due not only to the proliferation of the target organism, but also a change in food composition, and this needs further study.
DISCUSSION
The PCR assay is a useful detection method because of the demonstrated combination of speed and sensitivity, both of which are critical to any assay for the detection of bacteria. In addition to enhanced sensitivity, the use of unique oligonucleotide primers based on the sequence of the target DNA also results in absolute specificity. The primer set designed for this study was used to generate a 285-bp diagnostic PCR product. PCR detected five viable V. parahaemolyticus cells or correspondingly low levels (4 pg) of extracted DNA in a 100-μl PCR mixture. This sensitivity is consistent with that described for PCR for other bacteria of between 1 and 20 CFU (8, 16, 23, 28) or between 1 and 100 pg for DNA extracted from bacterial populations (8). A further increase in sensitivity from the picogram to femtogram level was achieved by Southern blot analyses (data not shown), as reported earlier (3). Here we report that the gyrB primer set recognized all vibrios identified as V. parahaemolyticus by conventional methods and had no false positives among the other bacteria tested.
Two major limitations to using PCR as a diagnostic tool are that false-positive reactions can occur from DNA contamination and that false-negative reactions can be derived from a number of substances found in samples that inhibit PCR (17, 22). We have included suitable negative and positive controls to overcome these limitations, when and where necessary. Sensitivity of detection in food samples has, however, been low, because only small samples (10 to 100 μl) can be analyzed and many kinds of food contain substances that are inhibitory to PCR. Such PCR-inhibitory substances were reported in many clinical samples, such as urine, blood, sputum, fecal specimens, food, and environmental specimens (7, 27, 29). With bacteria in food, the sensitivity of PCR was far lower than that with bacteria in saline. Also, the lower detection limit was higher than the number of CFU per unit volume usually found in processed food. Previous studies have suggested the extraction of DNA (8) or application of chemicals (4) and physical procedures (24) to remove PCR-inhibitory products. However, here we show that a simple preenrichment in a nonspecific medium can be applied to successfully remove any possible PCR-inhibitory substances and allow the proliferation of target bacteria. The detection of V. parahaemolyticus directly from food samples was possible by the combination of an 18-h enrichment in APW and the PCR assay, even when V. parahaemolyticus and other competitive food microflora were present at a ratio of 1:102. Because there is no DNA extraction step involved, with simple training to handle PCR machines, the procedure we present here should be used effectively by food monitors in industry for the detection of the bacterium.
The findings reported here describe a rapid, sensitive, specific, and reliable method for the detection of V. parahaemolyticus in shrimp. The fact that this technique allows the detection of genetic potential and differentiates between related species may make it useful as both a screening and a confirmatory test. The data obtained in this manner could yield additional information useful to epidemiological studies.
ACKNOWLEDGMENTS
We are grateful to Sumio Shinoda, Okayama University, and Mikio Satake for the supply of strains and useful discussions and D. Moser and K. Nealson for critically reading the manuscript. We are also thankful to Takashi Kurusu, Katsuyuki Hanai, Akiko Murakoshi, and Yuri Kamijoh for technical assistance.
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