APPLIED AND ENVIRONMENTAL MICROBIOLOGY, May 2004, p. 2678–2684 0099-2240/04/$08.00⫹0 DOI: 10.1128/AEM.70.5.2678–2684.2004 Copyright © 2004, American Society for Microbiology. All Rights Reserved.
Vol. 70, No. 5
Detection of Enteric Viruses in Shellfish from the Norwegian Coast M. Myrmel,1* E. M. M. Berg,2 E. Rimstad,1 and B. Grinde2 Department of Food Safety and Infection Biology, The Norwegian School of Veterinary Science, 0033 Oslo,1 and Division of Infectious Disease Control, Norwegian Institute of Public Health, 0403 Oslo,2 Norway Received 7 December 2003/Accepted 4 February 2004
Common blue mussels (Mytilus edulis), horse mussels (Modiolus modiolus), and flat oysters (Ostrea edulis) obtained from various harvesting and commercial production sites along the Norwegian coast were screened for the presence of norovirus by a real-time reverse transcription (RT)-nested PCR assay and for possible indicators of fecal contamination, i.e., for F-specific RNA bacteriophages (F-RNA phages) by plaque assay and for human adenoviruses and human circoviruses by nested PCR assay. The aims were to obtain relevant information for assessing the risk of transmission of enteric viruses by shellfish and to investigate the potential of various indicator viruses in routine screening. Noroviruses were detected in 6.8% of the samples, and the indicators were detected in 23.8% (F-RNA phages), 18.6% (adenoviruses), and 8.0% (circoviruses) of the samples. A seasonal variation was observed, with the exception of circoviruses, with more positive samples in the winter. A positive correlation was found between F-RNA phages and noroviruses. However, F-RNA phages were present in only 43% of the norovirus-positive samples. The results show that mussels from the Norwegian coast can constitute a risk of infection with enteric viruses and that routine testing of samples may be justified. Advantages and disadvantages of various options for screening are discussed. composition, and the ability to survive in the environment, but the suitability of F-RNA phages as indicators of human viruses in shellfish seems to vary with the type of virus and the geographical location (16, 19). It is therefore of interest to investigate whether other viral indicators might be more useful. In the present study, we assessed the viral contamination of shellfish from areas along the Norwegian coast with various levels of fecal pollution. NV were genotyped to map the distribution of strains (54), and F-RNA phages were genotyped in an attempt to indicate a human or animal origin (40, 45, 46). In addition to testing for the presence of NV and F-RNA phages, we evaluated two commonly occurring viruses as possible alternative indicators: human adenoviruses (hAdV) and human circoviruses (huCV). The latter involved separate analyses of TT virus (TTV) and TTV-like minivirus (TLMV). Adenoviruses are prevalent in sewage, and recent studies have indicated that they may serve as indicators of enteric viruses in shellfish (19, 41). Human circoviruses are a recently discovered group of small DNA viruses (39, 50). They replicate continuously, are shed in the feces, and are present in the majority of people worldwide (25, 34, 49). To our knowledge, they have not previously been suggested as indicators of fecal contamination, but their prevalence suggests that they could prove useful.
Disease caused by the consumption of bivalve molluscan shellfish containing pathogenic viruses of human origin is a well-known phenomenon, particularly in connection with raw oysters (29). The shellfish concentrate viral particles as a consequence of their feeding process, i.e., the filtering of large water volumes. When water is contaminated with human feces, viral pathogens may get trapped in the shellfish. Several enteric viruses may use this route of infection. Noroviruses (NV; old term, Norwalk-like viruses), gastroenteritis viruses of the family Caliciviridae, seem to be the predominant cause of disease worldwide; however, hepatitis A virus-associated outbreaks have been reported from several countries such as the United States, Italy, and China (42). Commercial use of shellfish is an expanding industry, which may increase the transmission of pathogens associated with shellfish. The presence of fecal indicator bacteria is routinely used for microbiological quality assurance of shellfish (3). However, the bacteria are not reliable indicators of the presence of enteric viruses in bivalves, as viral particles generally are more resistant to inactivation in water sources and are more slowly removed from shellfish by depuration (38, 47). In fact, outbreaks of viral gastroenteritis due to oysters complying with the relevant European fecal coliform standards have been reported (11, 29). Testing for the presence of either pathogenic viruses or indicator viruses should therefore be considered. Owing to their abundance in sewage, F-specific RNA bacteriophages (F-RNA phages) have been suggested as possible indicators of fecal contamination (22). F-RNA phages use Escherichia coli as a host but are unlikely to replicate in bacteria outside the gut, at least in temperate climates (56). The F-RNA phages are relatively similar to enteric viruses in size,
MATERIALS AND METHODS Samples. A total of 681 mussel samples, either the common blue mussel (Mytilus edulis) or the horse mussel (Modiolus modiolus), were obtained from the regional offices of the Directorate of Fisheries or from local food control authorities or were harvested directly from the Oslo Fjord. From each locality, mussels were gathered every 2 to 4 weeks over a period lasting from 3 to 36 months between June 2000 and June 2003. The localities were representative for the various parts of the Norwegian coast and included sites used for commercial production, sites used for noncommercial harvesting, and sites known to be polluted by sewage. The commercial production sites were all classified as category A areas in accordance with the European directive (3). The mussels were put on ice upon harvesting and shipped overnight to the laboratory. An additional 15 flat-oyster (Ostrea edulis) samples were obtained from a distributor.
* Corresponding author. Mailing address: The Norwegian School of Veterinary Science, Department of Food Safety and Infection Biology, P.O. Box 8147 Dep., 0033 Oslo, Norway. Phone: 47 22964771. Fax: 47 22964818. E-mail: [email protected]
VOL. 70, 2004
DETECTION OF ENTERIC VIRUSES IN SHELLFISH
They were processed in the laboratory the same day. Each sample consisted of 10 to 25 mussels or five oysters. All of the 696 shellfish samples were tested for NV and F-RNA phages, 86 samples were also tested for hAdV, and 113 samples were also tested for huCV (TTV and TLMV). Recovery of viral particles. The method used for recovery of viral particles was based on previous studies (37, 41, 52). Briefly, the outer surface of the shellfish was rinsed in water prior to opening. The intestine and hepatopancreas were dissected and homogenized in a food processor. Homogenates (approximately 25 g) were diluted with equal volumes of glycine buffer (0.05 M glycine, 0.15 M NaCl, pH 9.0), and the solution was treated for 15 min at 4°C on a shaker in order to release virus from the tissue. The homogenates were subsequently centrifuged at 5,000 ⫻ g for 15 min, and the shellfish supernatant was collected for either phage analysis or further concentration of viral particles. In the latter case, samples (12 ml) were centrifuged at 190,000 ⫻ g for 90 min at 4°C (Beckman SW40TI). The pellets were resuspended in 250 l of phosphate-buffered saline, aliquoted, and stored at ⫺70°C prior to use (referred to as shellfish extract). After every fifth sample, a negative control was included, consisting of glycine buffer prepared together with the shellfish samples. RNA extraction and purification. Viral RNA was isolated from 50 l of shellfish extract by addition of 100 l of TRIzol (Gibco). After 5 min of incubation at room temperature, 40 l of chloroform was added and the tubes were incubated for another 2 min at room temperature. The preparations were then centrifuged at 1,200 ⫻ g for 15 min in order to separate the phases. The RNA was isolated from the water phase by addition of a suspension of silica particles (40 l) and 900 l of lysis buffer (guanidinthiocyanate in 0.1 M Tris hydrochloride, pH 6.4, supplemented with EDTA and Triton X-100) (7). After 10 min at room temperature and subsequent vortexing and centrifugation (12,000 ⫻ g for 15 s), the silica particles were washed twice with washing buffer (guanidinthiocyanate in 0.1 M Tris hydrochloride, pH 6.4), twice with 70% ethanol, and once with acetone. The silica particles were dried at 56°C for 10 min, and the RNA was eluted in 80 l of diethyl pyrocarbonate-treated water containing 160 M RNase inhibitor (ribonucleoside vanadyl complexes; Sigma). The purified RNA was stored at ⫺70°C prior to use. NV RT-nPCR. Five microliters of RNA template was used in a total reaction volume of 50 l with the Qiagen OneStep reverse transcription (RT)-PCR kit (Qiagen). RT was performed at 37°C for 30 min with 0.6 M each NV outer primer MJV12 (5⬘-TAY CAY TAT GAT GCH GAY TA-3⬘; nucleotides 4553 to 72) and RegA (5⬘-CTC RTC ATC ICC ATA RAA IGA-3⬘; nucleotides 4859 to 79) (J. Vinje´, personal communication). The positional numbers correspond to the sequence with GenBank accession number M87661. The RT enzyme was inactivated, and the polymerase was activated, by incubation at 95°C for 15 min. In order to increase assay specificity, a touchdown PCR was run, starting with annealing at 50°C and ending at 43°C after 15 cycles. An annealing temperature of 37°C was used for the last 25 cycles. Amplification cycles were 94°C for 30 s, annealing for 90 s, and 72°C for 30 s. A final elongation at 72°C for 7 min was used. For the nested PCR (nPCR), 0.5-l aliquots of the first PCR product were included in a total volume of 25 l, with the QuantiTect SYBRGreen PCR kit (Qiagen). The forward nested primers used were p290 (5⬘-GAT TAC TCC AAG TGG GAC TCC AC-3⬘; nucleotides 4568 to 90) (26) and Mp290 (5⬘-GAT TAT ACT SSM TGG GAY TCM AC-3⬘; nucleotides 4568 to 90). The reverse nested primers were modified versions of those published by Ando et al. (2), i.e., RevSR46 (5⬘-CCA GTG GGC GAT GGA ATT CCA-3⬘; nucleotides 4786 to 66) and RevSR48-52 (5⬘-CCA RTG RTT TAT RCT GTT CAC-3⬘; nucleotides 4786 to 66). Each of the nested primers was used at a final concentration of 0.32 M. Incubation at 95°C for 15 min was followed by 40 cycles of 94°C for 20 s, 49°C for 90 s, and 72°C for 30 s. The nPCR was run on a SmartCycler real-time PCR machine (Cepheid). Fluorescence was read at 78°C for 10 s, and a melt curve analysis was performed to test the specificity of the product. Each run included negative and positive controls. DNA extraction and purification. Viral DNA was isolated from 100 l of shellfish extract supplemented with 100 l of double-distilled H2O with the High Pure viral nucleic acid kit (Roche) as recommended by the manufacturer. The samples were eluted in 50 l of the elution buffer provided (double-distilled H2O) and stored at ⫺20°C prior to use. Adenovirus nPCR. The following primers were used (positional numbers correspond to the sequence with GenBank accession number M73260). The outer primers used were AdVof (5⬘-GAC ATG ACT TTT GAG GTG GAC CC-3⬘; nucleotides 21545 to 67) and AdVor (5⬘-CCG GCC GAG AAG GGC GT-3⬘; nucleotides 21684 to 68); the nested primers used were AdVif (5⬘-TTT GAG GTG GAC CCC ATG GA-3⬘; nucleotides 21554 to 73) and AdVir (5⬘-GAG AAG GGC GTG CGC AGG TA-3⬘; nucleotides 21678 to 59). The primers were designed, on the basis of available sequences, to specifically detect the human subtypes. The PCR conditions were 0.5 M each primer, 1.5 mM MgCl2, 0.25
mM deoxynucleoside triphosphate, and 1 U of AmpliTaq Gold DNA polymerase per reaction in GeneAmp PCR Gold buffer (Applied Biosystems). The final volumes were 50 l (outer PCR), including 3 l of template, and 25 l (nPCR), including 1 l of template. The polymerase was activated for 10 min at 95°C, followed by 28 (outer PCR) or 30 (nPCR) amplification cycles of 95°C for 15 s, 60°C for 15 s, and 72°C for 15 s and a final elongation step of 72°C for 2 min. The PCR products from the nPCR were analyzed by agarose gel electrophoresis (2% agarose with ethidium bromide). Each run included negative and positive controls. Circovirus (TTV and TLMV) nPCRs. The outer primers were designed to capture both TTV and TLMV, while the nested primers were specific for either virus (36). The outer primers were TTV/TLMVf (5⬘-TCC GAA TGG CTG AGT TT-3⬘; nucleotides 102 to 118) and TTV/TLMVr (5⬘-CGA ATT GCC CCT TGA CT-3⬘; nucleotides 219 to 203). The nested primers for TTV (2:1 mixture of a and b) were primers TTVfa (5⬘-GTT TTC TAC GCC CGT CC-3⬘; nucleotides 115 to 131) and TTVfb (5⬘-GTT TTC YAC GCC CGT CC-3⬘; nucleotides 115 to 131) and primers TTVra (5⬘-CCT TGA CTC CGG TGT GTA A-3⬘; nucleotides 210 to 192) and TTVrb (5⬘-CCT TGA CTB CGG TGT GTA A-3⬘; nucleotides 210 to 192). The nested primers for TLMV were TLMVf (5⬘-AGT TTA TGC CGC CAG ACG-3⬘; nucleotides 193 to 210) and TLMVr (5⬘-CCC TAG ACT TCG GTG GTT TC-3⬘; nucleotides 287 to 268). The positional numbers are those of the TTV strain TA278 genome (GenBank accession no. AB008394) and TLMV reference strain CBD231 (GenBank accession no. AB026930). The PCR conditions were the same as described above for the hAdV PCR, except for the 2.5 mM MgCl2 final concentration in both huCV PCRs. Genotyping of NV. In order to verify the NV RT-nPCR results and to genotype the detected strains, reverse line blot (RLB) hybridization was performed with a protocol slightly modified from that of Vinje´ and Koopmans (54). The products from the real-time nPCR were too short to include all of the binding sites for the hybridization probes. Therefore, the outer RT-PCR products from samples that were positive in the real-time nPCR were used in a semi-nPCR to produce DNA fragments of sufficient length for RLB hybridization. The primers (0.3 M) used were p290, Mp290, and biotinylated RegA (defined above). The HotStarTaq Master Mix kit (Qiagen) was used, with cycling conditions as described for the NV real-time nPCR. The nylon membrane (Roche) blots included 3 genogroupspecific probes (GIa, GIb, and GII) and 15 genotype-specific probes (sequences are listed in reference 54) immobilized in parallel rows. The biotinylated PCR products were applied in columns at a 90° angle to the probes, and hybridization was performed at 50°C for 60 min. The membranes were subsequently washed, incubated with streptavidin-peroxidase conjugate (Roche) for 45 min at 42°C, and washed again. The PCR products were detected by chemiluminescence (ECL; Amersham Pharmacia) and visualized by exposure of an X-ray film (Roche) for 15 min. Sequencing. The nPCR products from nine samples of NV which could not be genotyped by RLB hybridization, as well as eight samples of hAdV, were sequenced in order to examine the authenticity of the PCR products. The products were sequenced in both directions on an ABI PRISM 310 (Applied Biosystems) automatic sequencing machine with an ABI BigDye Terminator Cycle Sequencing kit (Applied Biosystems) and the inner primers referred to above. The sequences were analyzed with Sequencer 3.1 (Gene Codes) and compared to sequences in the GenBank database. Plaque assay for detection of F-RNA phages. The ISO 10705-1 method (4) for enumeration of F-RNA phages was used during the first 12 months of the study. Owing to sudden and repeated loss of the host strain’s (Salmonella enterica serovar Typhimurium WG49) sensitivity to phages, E. coli HS(pFamp)R (14) was used for the last 24 months of the study. E. coli HS(pFamp)R was kindly provided by M. D. Sobsey, University of North Carolina, Chapel Hill. A previous study showed that S. enterica serovar Typhimurium WG49 detected slightly higher numbers of F-RNA phages than did E. coli HS(pFamp)R (45). There were no differences between the two host strains with regard to detection of the four genotypes of F-RNA phages. A mixture of 1 ml of shellfish supernatant (based on 0.5 g of intestinal tract), 80 l of bacterium culture [E. coli HS(pFamp)R] in the exponential growth phase, and 5 ml of tryptic soy broth semisolid agar (0.7%) with ampicillin and streptomycin at 0.015% each was poured on top of tryptic soy broth solid agar (1.5%). Samples were initially run in two parallel experiments, with and without RNase, in order to estimate the relative amount of f-RNA phages to that of DNA phages. However, this protocol was eventually found to be redundant because of the relative scarcity of RNase-insensitive phages. Plaques were counted after approximately 18 h of incubation at 37°C. Plaque hybridization for genotyping of F-RNA phages. Plaques from selected plates were transferred to four positively charged nylon membranes (Roche) by repeated plaque lifts. The virus particles and nucleic acids were denatured with 7.5⫻ SSC (1⫻ SSC is 0.15 M NaCl plus 0.015 M sodium citrate) and 4.6 M
MYRMEL ET AL.
APPL. ENVIRON. MICROBIOL.
TABLE 1. Monthly detection of NV and F-RNA phages in mussels collected between June 2000 and June 2003a Mo
January February March April May June July August September October November December Total
No. (%) of samples positive
No. of samples tested
74 62 78 52 50 64 25 45 55 47 64 65
7 (9.5) 5 (8.1) 6 (7.7) 5 (9.6) 0 (0.0) 3 (4.7) 0 (0.0) 2 (4.4) 4 (7.3) 6 (12.8) 4 (6.3) 4 (6.2)
F-RNA phage levelb
Presence of NV
F-RNA phages 1–9 10–99 100–999 ⱖ1,000
681 46 (6.8)
31 (41.9) 22 (35.5) 23 (29.5) 8 (15.4) 3 (6.0) 4 (6.3) 1 (4.0) 6 (13.3) 6 (10.9) 7 (14.9) 27 (42.2) 24 (36.9)
21 11 13 7 1 4 1 6 6 2 22 13
7 6 8 1 1
1 5 1
5 3 10
9/113 (8.0) 0.132
The samples were split between those positive and those negative for NV, and the number of indicator virus-positive samples in each group is indicated. b The significance of the correlations was tested with the Pearson chi-square (two-sided) test.
Prevalences. NV was detected in 46 (6.8%) of the 681 mussel samples, while F-RNA phages were detected in 162 (23.8%) (Table 1). A higher prevalence was found for both NV and F-RNA phage-positive samples in the winter (October through March), but with a significant correlation (P ⬍ 0.01) only for the F-RNA phage-positive samples (Table 2). Of the 46 NV-positive samples, only 20 (43.5%) were also positive for F-RNA phages and 19 of these 20 samples were found during October through March, 1 was found in April, and none were found in May through September. NV was detected in samples from all of the participating counties except for two counties in northern Norway, and there was a tendency for the presence of viruses to correlate with the population density of the area. Repeated positive NV results were obtained from 8 of the 40 TABLE 2. Prevalence of viruses in mussels in summer and wintera No. of samples positive/no. tested (% positive) NV
14/291 (4.8) 32/390 (8.2)
28/291 (9.6) 134/390 (34.4)
2/44 (4.5) 14/42 (33.3)
5/52 (9.6) 4/61 (6.6)
4/27 (14.8) 5/86 (5.8)
8/27 (29.6) 8/59 (13.6)
20/46 (43.5) 142/635 (22.4)
formaldehyde at 65°C for 15 min, and RNA was fixed by baking at 80°C for 90 min (24). The four membranes were hybridized with four different digoxigeninmarked probes (numbered 1 to 4) at 37°C overnight (6). Anti-digoxigenin antibody conjugated with alkaline phosphatase (Roche) and CSPD (Roche) was used to produce chemiluminescence detectable on X-ray films (Kodak).
Each sample was based on an intestine- hepatopancreas extract from 10 to 25 mussels and analyzed by either RT-nPCR (NV) or plaque assay (F-RNA phages). b The number of PFU per 0.5 g of intestine-hepatopancreas has been grouped into four intervals, and the number of samples within each interval is indicated.
No. of samples positive/no. tested (% positive) F-RNA phages
Season or parameter
TABLE 3. Correlation between the presence of NV and that of candidate indicator virusesa
F-RNA phages were detected by plaque assay; the other viruses were detected by PCR methods designed to specifically detect human variants. b April to September. c October to March. d Determined with the Pearson chi-square (two-sided) test.
commercial harvesting areas. The two areas with the most positive samples had, respectively, 22% (4 of 18) and 9% (4 of 44) of the samples positive for NV and 33 and 29% of the samples positive for phages. After analyzing 390 shellfish samples by plaque assay, a total of approximately 9,200 PFU of F-specific phages, but only 58 PFU of DNA phages, had been detected. Consequently, as DNA phages did not seem to interfere appreciably with the shellfish RNA phage assay, a parallel sample with RNase treatment was no longer included. The number of phages observed varied considerably between samples. Of the positive samples, 66% had less than 10 PFU/0.5 g of intestine-hepatopancreas tissue, 25% had 10 to 99 PFU, 7% had 100 to 999 PFU, and 2% had ⱖ1,000 PFU (Table 1). Of the 55 samples with 10 or more PFU/0.5 g of intestine-hepatopancreas, 52 (94.5%) were found in the winter. Assuming that the intestine-hepatopancreas represents approximately 10% of the weight of the shellfish, and that the phages are present solely in this tissue, then the phage-positive mussels contained from 20 to 40,000 PFU/100 g of edible shellfish. The average level of F-RNA phages found in the samples positive for NV was 2,040 (range, 20 to 15,200) PFU/ 100 g. Of the samples tested for alternative indicator viruses, hAdV was detected in 18.6% (16 of 86) and huCV was detected in 8.0% (9 of 113) (TTV in 6 samples and TLMV in 5 samples, including 2 samples with both viruses). hAdV was significantly (P ⬍ 0.01) more prevalent in the winter; the occurrence of huCV, however, did not show a seasonal variation (Table 2). There was a significant correlation between the presence of NV and the presence of F-RNA phages (P ⬍ 0.01), but not for hAdV and huCV (P ⬎ 0.05) (Table 3). Of the oyster samples gathered during the same 3-year period, 3 out of 15 were positive for F-RNA phages while none were positive for NV. The three positive samples were among the eight samples obtained during the winter. Genotyping of NV. PCR products from the 46 NV-positive samples were subtyped by RLB hybridization (semi-nPCR products) or sequencing (nPCR products). Of the 46 PCR products, 30 hybridized to the GII probe, 10 hybridized to the GIb probe, 3 hybridized to the GIa probe, and 9 did not react with any genogroup-specific probe. Three products reacted with all of the GII, GIb, and GIa probes, and six hybridized to both GII and GIb. Two products reacted only with the GIb
VOL. 70, 2004
DETECTION OF ENTERIC VIRUSES IN SHELLFISH
probe, and 23 reacted solely with the GII probe. Of the 46 PCR products, 10 hybridized with one or several of the genotypespecific probes. Three of these 10 products did not react with any group-specific probes but were typed as Leeds strains on the basis of the genotype probes. Three other products came up as single strains: Desert Shields, Lordsdale, and Hawaii. Four products hybridized with more than one probe: one hybridized with Sindlesham and Leeds, one hybridized with Lordsdale and Wortley, one hybridized with Lordsdale and Hawaii, and one hybridized with Winchester, Sindlesham, Lordsdale, Leeds, and Wortley. Three of these four products originated from shellfish collected in areas most vulnerable to sewage contamination, in February 2001 (one sample) and February 2003 (three samples). The PCR product that hybridized with five genotype-specific probes represented shellfish from the Oslo city marine area. Nine of the nPCR products reacted with neither genogroupnor genotype-specific probes. Three of these products and six products that only hybridized with the GII probe were sequenced. Compared with sequences in the GenBank database, five samples (including the three RLB hybridization-negative strains) were related to Lordsdale. Four samples were related to a recent variant designated GGIIb that circulated throughout several European countries during 2000 and 2001 (10). Sequencing of adenovirus. Of the 16 samples that were positive in the hAdV nPCR, 8 were sequenced. When compared to GenBank sequences, they all matched hAdV most closely. Moreover, they matched the two subtypes, 40 and 41, normally associated with gastroenteritis in humans. Genotyping of F-RNA phages. Genotyping of 1,565 F-RNA phage plaques originating from 52 shellfish samples was performed in order to distinguish between phages associated with animals (G1 and G4) and those associated particularly with humans (G2 and G3) (24). Of the 52 samples, 22 contained solely phages belonging to G1 and 16 contained only G2 phages. Of the remaining 14 samples, 11 contained a mixture of G1 and G2; 1 contained G1, G2, and G3; and 2 contained G1, G2, and G4 phages. A predominance of G1 and G2 phages was found for the two sites most vulnerable to contamination with sewage.
enteritis outbreaks associated with mussels: first, in Norway, mussels are often collected by the consumers themselves, which decreases the chance of having outbreaks of sufficient magnitude to be registered by the health authorities; second, shells are typically harvested during the summer months, a season when shellfish are less likely to contain NV (16, 20, 30). The present study was initiated to obtain knowledge of the presence of fecal viral contamination in Norwegian shellfish. Mussels were the primary target, as they are consumed in larger quantities than oysters, but the results presumably reflect the risk of viral contamination of any shellfish. A real-time RT-nPCR was developed for the detection of NV. Owing to the genomic heterogeneity among the NV, SYBRGreen staining of RT-nPCR products was preferred for verification rather than fluorescing probes, as the latter may not hybridize as a consequence of nucleotide variation. Moreover, the specificity of the assay was confirmed by hybridization, as well as by sequencing of some of the amplification products. Of the 681 samples tested, based on approximately 9,000 mussels, 6.8% contained NV. Two of the sites used for mussel collection were in the marine areas outside the cities of Oslo and Trondheim and are considered representative of the polluted part of the coast. Excluding these samples, the prevalence of NV would be 5.6%. The presence of NV in mussels was anticipated, as raw oysters have been the suspected source for outbreaks of NV infections in Norway, according to the Norwegian Food Control Authority. The prevalence obtained is most likely an underestimate due to the limits of sensitivity inherent in any test method. On the other hand, molecular detection does not discriminate between infective and noninfective viral particles (15). Although a contaminated environment is indicated, the potential health impact is uncertain. A system for NV cultivation does not exist (5). RLB hybridization was useful for verification of NV PCR products, and for grouping of strains into genogroup GI or GII. Most strains belonged to GII, which is in accordance with the worldwide dominance of GII strains in fecal samples (18, 27, 32, 48). However, hybridization to strain-specific probes failed in 36 of 46 samples, presumably because of variations in the part of the genome used for these probes (55). Of the 10 shellfish samples that hybridized to strain-specific probes, 4 reacted to more than one probe. This result might be anticipated, as NV in shellfish presumably reflects all of the NV strains that circulate in the population. Three of the four shellfish samples were collected in densely populated areas during the winter, i.e., when most cases of NV gastroenteritis are reported (53). Most of the Norwegian coastline is reasonably sheltered from fecal contamination because the land is sparsely populated and the sewage is generally treated in decontamination plants. On the other hand, the treatment types differ and there is limited information about the effects of various sewage treatments on infectious viruses. Moreover, fecal waste from persons recreating in or on the water may be a source of sporadic contamination. NV was detected in samples from all but two of the counties studied, indicating that even in apparently unpolluted seawater there is a potential risk involved when consuming shellfish. As expected, there was a tendency for the pres-
DISCUSSION NV are regarded as one of the most common causes of foodborne infections (17, 31, 51). Foodborne transmission of NV is typically due to contamination caused by people handling the food rather than by contaminated raw materials; however, in the case of shellfish, particularly oysters, this is not the case. Several outbreaks of gastroenteritis due to NV have been associated with consumption of raw oysters grown in fecally contaminated waters (29). In Norway, mussels are the shellfish most commonly used for human consumption. Presumably, mussels are as likely to be contaminated with human enteric viruses as are oysters, yet they are rarely recognized as sources of viral infection. This observation may be partly explained by the custom of steaming or boiling mussels prior to consumption. On the other hand, the heating required to open the shells is not necessarily sufficient to inactivate viruses (1, 28, 33). Thus, there may be other factors that help explain the scarcity of reported gastro-
MYRMEL ET AL.
ence of viruses to correspond to the population density of the area. The percentages of NV- or hAdV-positive shellfish in this study were equal to or lower than those reported for related studies in category A (depuration or relaying not required) or nonclassified areas in other European countries (20). The lack of standardized methods may contribute to the variability of the results; however, the present data can also be interpreted to support the assumption that Norwegian coastal waters are relatively unpolluted. The latter interpretation is supported by the fact that only two of the present harvesting areas in Norway are classified as type B (depuration or relaying required) according to the European directive (3). Although NV may be detected in feces for up to 3 weeks after infection (21, 43), one would not expect NV particles to be continuously present in the sewage of small Norwegian coastal settlements. Therefore, the absence of NV does not necessarily imply that the shellfish are free of fecal contamination. In order to monitor the risk of fecal contamination, it would be preferable to have indicators that reflect the possible presence of any human virus. The F-RNA phages have been extensively investigated for this purpose. According to some reports, these phages correlate well with the presence of human enteric viruses (12, 16), while in other reports the correlation is less obvious (23). In the present study, the occurrence of F-RNA phages correlated significantly with that of NV (Table 3). However, more than half of the NV-positive samples were negative for F-RNA phages and a positive F-RNA phage result was less than twice as common in samples with NV than in those without NV. Thus, it is questionable whether the results can be taken to support the use of F-RNA phages as indicators of NV contamination. The F-RNA phages do, however, offer the advantages of ease of methodology and of screening approximately seven times as much bivalve tissue as the PCR-based methods. The F-RNA phages appear to be consistently present in sewage (22, 45, 46), yet they are not always present in the feces of individual humans (22, 40, 46). Sporadic fecal contamination may therefore explain the observation that several samples were NV positive and F-RNA phage negative. Differences in viral stability may also help explain this observation. Chung et al. (13) found that F-RNA phages were inactivated faster than hepatitis A virus, poliovirus, and rotavirus at 25°C, while at 5°C the inactivation of phages and poliovirus was comparable. The present results support this observation in that 19 of the 20 F-RNA phage- and NV-positive samples were found in the winter. Moreover, the majority of samples with high levels of F-RNA phages were found in the cold season. This seasonal distribution can hardly be attributed to variations in viral production but more likely reflects viral stability. The winter corresponds to both a decrease in sea temperature and a decrease in the biological activity in freshwater and seawater; both factors may help conserve viruses (8). It should also be pointed out that the detection of phages is based on replication-competent phage particles, while PCR does not discriminate between viable and nonviable virus particles. In a study of oysters collected in connection with outbreaks of gastroenteritis in Great Britain, all of the samples tested positive for F-RNA phages (29). The average concentration of F-RNA phages (2,500 [range, 60 to 17,500] PFU/100 g) was
APPL. ENVIRON. MICROBIOL.
comparable to what was found in the samples positive for both NV and F-RNA phages in the present study (2,040 [range, 20 to 15,200] PFU/100 g). The variability of the number of phages present, in both the British study and the present study, supports the suggestion that samples may contain pathogenic viruses even if they lack F-RNA phages. Genotyping of F-RNA phages showed a predominance of types G1 and G2, indicating fecal contaminants from animal (G1) and human (G2) sources. The same distribution was found in shellfish from the two sites most vulnerable to contamination with sewage, but several of these samples contained only G1 phages. Phages belonging to G3, a genotype that is consistently related to human fecal contamination (9), were only found in one sample (obtained from a commercial harvesting area). Differences in inactivation rates might help explain the scarcity of G3 phages, and different rates have been found by Schaper: G3 and G4 phages were more sensitive to environmental stresses than were G1 phages, while G2 phages showed intermediate resistance (44). In fact, G3 phages have been proposed as an indicator of recent human fecal pollution (9). The results of the present study support the impression that this system for genotyping of F-RNA phages has limited relevance to the analysis of sources of fecal contamination (9, 44), at least regarding shellfish. The limited reliability of phages as predictors of human pathogenic viruses prompted the examination of alternative indicators. hAdV constitute a possible alternative, as they appear to be common in sewage (41). In fact, they were found in 18.6% of the shellfish samples tested in our study. The presence of hAdV did not correlate significantly with the detection of NV (Table 3); however, this may have been due to the relatively low number of samples tested. Other interesting viruses in this respect are the two newly discovered human circoviruses (TTV and TLMV). Both of these viruses seem to be persistently present in the majority of people, and they are shed in feces (25, 34, 49). Moreover, their lack of envelope and their compact size and small, circular DNA genomes indicate stability of the viral particles. Despite these properties, they were only found in 8% of the samples tested and their presence did not correlate significantly with that of NV (Table 3). Their low prevalence in bivalves probably reflects a combination of low titers of shed virus (35) and that they are not particularly stable in sewage or marine environments. It is possible that low recovery of circovirus particles from shellfish tissue also contributed to the findings. Interestingly, while the presence of F-RNA phages and hAdV significantly peaked during the winter, that of NV and huCV did not (Table 2). NV showed a higher prevalence in shellfish collected during the winter; however, the correlation was not significant. NV was expected to be found more frequently in the winter, as the infection is often referred to as “winter vomiting disease,” while huCV probably are equally common throughout the year. The prevalence of NV in Swedish shellfish was also found to be relatively independent of season; however, in contrast to our results, mussels were found to contain more hAdV in the summer (23). In another study, including shellfish from Greece, Spain, Sweden, and the United Kingdom, no seasonal variation was found in the prevalence of hAdV (20). Besides the seasonal variations in human infections, the main factor that is assumed to have an impact
VOL. 70, 2004
DETECTION OF ENTERIC VIRUSES IN SHELLFISH
on the presence of viruses in shellfish is temperature, as discussed above. Local climatic variations in rainfall and water currents may also explain the apparent incompatibility of results coming from various geographical regions. There are no regulations or standards in Norway regarding the testing of commercial shellfish for viral contamination. Although the data reported here demonstrate that there is a potential risk involved in eating Norwegian shellfish, it is not obvious that routine screening for viruses is justified. There is limited commercial exploitation of shellfish. However, there is an increasing interest in mussel and oyster farming, and with increased production, the potential hazards of eating shellfish may become more obvious. Moreover, although the environmental load of viruses may be relatively small in Norway because of the low population density, the cold climate may enhance the survival of viruses. In order to improve the safety of mussel consumption, an important measure would be to ensure that mussels are grown in areas with limited fecal contamination. F-RNA phages may be used as a supplement to E. coli as indicators of fecal contamination in the risk assessments of possible locations. The present results suggest that if viral testing of shellfish is introduced on a routine basis, the more meaningful parameters to choose may be NV or a combination of F-RNA phages and hAdV.
13. Chung, H., and M. D. Sobsey. 1993. Comparative survival of indicator viruses and enteric viruses in seawater and sediment. Water Sci. Technol. 27:425– 428. 14. Debartolomeis, J., and V. J. Cabelli. 1991. Evaluation of an Escherichia coli host strain for enumeration of F male-specific bacteriophages. Appl. Environ. Microbiol. 57:1301–1305. 15. De Medici, D., L. Croci, S. Di Pasquale, A. Fiore, and L. Toti. 2001. Detecting the presence of infectious hepatitis A virus in molluscs positive to RTnested-PCR. Lett. Appl. Microbiol. 33:362–366. 16. Dore´, W. J., K. Henshilwood, and D. N. Lees. 2000. Evaluation of F-specific RNA bacteriophage as a candidate human enteric virus indicator for bivalve molluscan shellfish. Appl. Environ. Microbiol. 66:1280–1285. 17. Fankhauser, R. L., S. S. Monroe, J. S. Noel, C. D. Humphrey, J. S. Bresee, U. D. Parashar, T. Ando, and R. I. Glass. 2002. Epidemiologic and molecular trends of “Norwalk-like viruses” associated with outbreaks of gastroenteritis in the United States. J. Infect. Dis. 186:1–7. 18. Fankhauser, R. L., J. S. Noel, S. S. Monroe, T. Ando, and R. I. Glass. 1998. Molecular epidemiology of “Norwalk-like viruses” in outbreaks of gastroenteritis in the United States. J. Infect. Dis. 178:1571–1578. 19. Formiga-Cruz, M., A. K. Allard, A. C. Conden-Hansson, K. Henshilwood, B. E. Hernroth, J. Jofre, D. N. Lees, F. Lucena, M. Papapetropoulou, R. E. Rangdale, A. Tsibouxi, A. Vantarakis, and R. Girones. 2003. Evaluation of potential indicators of viral contamination in shellfish and their applicability to diverse geographical areas. Appl. Environ. Microbiol. 69:1556–1563. 20. Formiga-Cruz, M., G. Tofino-Quesada, S. Bofill-Mas, D. N. Lees, K. Henshilwood, A. K. Allard, A. C. Conden-Hansson, B. E. Hernroth, A. Vantarakis, A. Tsibouxi, M. Papapetropoulou, M. D. Furones, and R. Girones. 2002. Distribution of human virus contamination in shellfish from different growing areas in Greece, Spain, Sweden, and the United Kingdom. Appl. Environ. Microbiol. 68:5990–5998. 21. Graham, D. Y., X. Jiang, T. Tanaka, A. R. Opekun, H. P. Madore, and M. K. Estes. 1994. Norwalk virus infection of volunteers: new insights based on improved assays. J. Infect. Dis. 170:34–43. 22. Havelaar, A. H., W. M. Pot-Hogeboom, K. Furuse, R. Pot, and M. P. Hormann. 1990. F-specific RNA bacteriophages and sensitive host strains in faeces and wastewater of human and animal origin. J. Appl. Bacteriol. 69:30–37. 23. Hernroth, B. E., A.-C. Conden-Hansson, A.-S. Rehnstam-Holm, R. Girones, and A. K. Allard. 2002. Environmental factors influencing human viral pathogens and their potential indicator organisms in the blue mussel, Mytilus edulis: the first Scandinavian report. Appl. Environ. Microbiol. 68:4523– 4533. 24. Hsu, F. C., Y. S. Shieh, J. van Duin, M. J. Beekwilder, and M. D. Sobsey. 1995. Genotyping male-specific RNA coliphages by hybridization with oligonucleotide probes. Appl. Environ. Microbiol. 61:3960–3966. 25. Huang, L. Y., J. T. Oystein, O. Hungnes, and B. Grinde. 2001. High prevalence of TT virus-related DNA (90%) and diverse viral genotypes in Norwegian blood donors. J. Med. Virol. 64:381–386. 26. Jiang, X., P. W. Huang, W. M. Zhong, T. Farkas, D. W. Cubitt, and D. O. Matson. 1999. Design and evaluation of a primer pair that detects both Norwalk- and Sapporo-like caliciviruses by RT-PCR. J. Virol. Methods 83: 145–154. 27. Kawamoto, H., K. Yamazaki, E. Utagawa, and T. Ohyama. 2001. Nucleotide sequence analysis and development of consensus primers of RT-PCR for detection of Norwalk-like viruses prevailing in Japan. J. Med. Virol. 64:569– 576. 28. Kirkland, K. B., R. A. Meriwether, J. K. Leiss, and W. R. Mac Kenzie. 1996. Steaming oysters does not prevent Norwalk-like gastroenteritis. Public Health Rep. 111:527–530. 29. Lees, D. 2000. Viruses and bivalve shellfish. Int. J. Food Microbiol. 59:81– 116. 30. Le Guyader, F., L. Haugarreau, L. Miossec, E. Dubois, and M. Pommepuy. 2000. Three-year study to assess human enteric viruses in shellfish. Appl. Environ. Microbiol. 66:3241–3248. 31. Lindqvist, R., Y. Andersson, B. de Jong, and P. Norberg. 2000. A summary of reported foodborne disease incidents in Sweden, 1992 to 1997. J. Food Prot. 63:1315–1320. 32. Lopman, B. A., D. W. Brown, and M. Koopmans. 2002. Human caliciviruses in Europe. J. Clin. Virol. 24:137–160. 33. McDonnell, S., K. B. Kirkland, W. G. Hlady, C. Aristeguieta, R. S. Hopkins, S. S. Monroe, and R. I. Glass. 1997. Failure of cooking to prevent shellfishassociated viral gastroenteritis. Arch. Intern. Med. 157:111–116. 34. Moen, E. M., L. Huang, and B. Grinde. 2002. Molecular epidemiology of TTV-like mini virus in Norway. Arch. Virol. 147:181–185. 35. Moen, E. M., S. Sagedal, K. Bjoro, M. Degre, P. K. Opstad, and B. Grinde. 2003. Effect of immune modulation on TT virus (TTV) and TTV-like-minivirus (TLMV) viremia. J. Med. Virol. 70:177–182. 36. Moen, E. M., J. Sleboda, and B. Grinde. 2002. Real-time PCR methods for independent quantitation of TTV and TLMV. J. Virol. Methods 104:59–67. 37. Muniain-Mujika, I., R. Girones, and F. Lucena. 2000. Viral contamination of shellfish: evaluation of methods and analysis of bacteriophages and human viruses. J. Virol. Methods 89:109–118.
ACKNOWLEDGMENTS The Norwegian Food Control Authority, the Directorate of Fisheries, the Norwegian Seafood Federation, and the Norwegian Research Council funded this work. We thank Tom Øystein Jonassen for help with the design of adenovirus primers and Inger Lill Anthonisen and Hilde Reiersen for technical assistance. REFERENCES 1. Abad, F. X., R. M. Pinto, R. Gajardo, and A. Bosch. 1997. Viruses in mussels: public health implications and depuration. J. Food Prot. 60:677–681. 2. Ando, T., S. S. Monroe, J. R. Gentsch, Q. Jin, D. C. Lewis, and R. I. Glass. 1995. Detection and differentiation of antigenically distinct small roundstructured viruses (Norwalk-like viruses) by reverse transcription-PCR and Southern hybridization. J. Clin. Microbiol. 33:64–71. 3. Anonymous. 1991. Council Directive of 15th July laying down the health conditions for the production and placing on the market of live bivalve molluscs (91/492/EEC). Off. J. Eur. Commun. 268:1–13. 4. Anonymous. 1995. ISO 10705–1. Water quality. Detection and enumeration of bacteriophages. Part 1: enumeration of F-specific RNA bacteriophages. International Organization for Standardization, Geneva, Switzerland. 5. Atmar, R. L., and M. K. Estes. 2001. Diagnosis of noncultivatable gastroenteritis viruses, the human caliciviruses. Clin. Microbiol. Rev. 14:15–37. 6. Beekwilder, J., R. Nieuwenhuizen, A. H. Havelaar, and J. van Duin. 1996. An oligonucleotide hybridization assay for the identification and enumeration of F-specific RNA phages in surface water. J. Appl. Bacteriol. 80:179–186. 7. Boom, R., C. J. A. Sol, M. M. m. Salimans, C. L. Jansen, P. M. E. Wertheimvan Dillen, and J. van der Noordaa. 1990. Rapid and simple method for purification of nucleic acids. J. Clin. Microbiol. 28:495–503. 8. Bosch, A. 1995. The survival of enteric viruses in the water environment. Microbiologia 11:393–396. 9. Brion, G. M., J. S. Meschke, and M. D. Sobsey. 2002. F-specific RNA coliphages: occurrence, types, and survival in natural waters. Water Res. 36:2419–2425. 10. Buesa, J., B. Collado, P. Lopez-Andujar, R. Abu-Mallouh, D. J. Rodriguez, D. A. Garcia, J. Prat, S. Guix, T. Llovet, G. Prats, and A. Bosch. 2002. Molecular epidemiology of caliciviruses causing outbreaks and sporadic cases of acute gastroenteritis in Spain. J. Clin. Microbiol. 40:2854–2859. 11. Christensen, B. F., D. N. Lees, K. Henshilwood, T. Bjergskov, and J. Green. 1998. Human enteric viruses in oysters causing a large outbreak of human food borne infection in 1996/97. J. Shellfish Res. 17:1633–1635. 12. Chung, H., L. A. Jaykus, G. Lovelace, and M. D. Sobsey. 1998. Bacteriophages and bacteria as indicators of enteric viruses in oysters and their harvest waters. Water Sci. Technol. 38:37–44.
MYRMEL ET AL.
38. Nasser, A. M., and S. D. Oman. 1999. Quantitative assessment of the inactivation of pathogenic and indicator viruses in natural water sources. Water Res. 33:1748–1752. 39. Nishizawa, T., H. Okamoto, K. Konishi, H. Yoshizawa, Y. Miyakawa, and M. Mayumi. 1997. A novel DNA virus (TTV) associated with elevated transaminase levels in posttransfusion hepatitis of unknown etiology. Biochem. Biophys. Res. Commun. 241:92–97. 40. Osawa, S., K. Furuse, and I. Watanabe. 1981. Distribution of ribonucleic acid coliphages in animals. Appl. Environ. Microbiol. 41:164–168. 41. Pina, S., M. Puig, F. Lucena, J. Jofre, and R. Girones. 1998. Viral pollution in the environment and in shellfish: human adenovirus detection by PCR as an index of human viruses. Appl. Environ. Microbiol. 64:3376–3382. 42. Potasman, I., A. Paz, and M. Odeh. 2002. Infectious outbreaks associated with bivalve shellfish consumption: a worldwide perspective. Clin. Infect. Dis. 35:921–928. 43. Rockx, B., M. De Wit, H. Vennema, J. Vinje´, E. de Bruin, Y. Van Duynhoven, and M. Koopmans. 2002. Natural history of human calicivirus infection: a prospective cohort study. Clin. Infect. Dis. 35:246–253. 44. Schaper, M., A. E. Dura ´n, and J. Jofre. 2002. Comparative resistance of phage isolates of four genotypes of F-specific RNA bacteriophages to various inactivation processes. Appl. Environ. Microbiol. 68:3702–3707. 45. Schaper, M., and J. Jofre. 2000. Comparison of methods for detecting genotypes of F-specific RNA bacteriophages and fingerprinting the origin of faecal pollution in water samples. J. Virol. Methods 89:1–10. 46. Schaper, M., J. Jofre, M. Uys, and W. O. Grabow. 2002. Distribution of genotypes of F-specific RNA bacteriophages in human and non-human sources of faecal pollution in South Africa and Spain. J. Appl. Microbiol. 92:657–667. 47. Schwab, K. J., F. H. Neill, M. K. Estes, T. G. Metcalf, and R. L. Atmar. 1998. Distribution of Norwalk virus within shellfish following bioaccumulation and subsequent depuration by detection using RT-PCR. J. Food Prot. 61:1674– 1680.
APPL. ENVIRON. MICROBIOL. 48. Smit, T. K., P. Bos, I. Peenze, X. Jiang, M. K. Estes, and A. D. Steele. 1999. Seroepidemiological study of genogroup I and II calicivirus infections in South and southern Africa. J. Med. Virol. 59:227–231. 49. Takahashi, K., H. Hoshino, Y. Ohta, N. Yoshida, and S. Mishiro. 1998. Very high prevalence of TT virus (TTV) infection in general population of Japan revealed by a new set of PCR primers. Hepatol Res. 12:233–239. 50. Takahashi, K., Y. Iwasa, M. Hijikata, and S. Mishiro. 2000. Identification of a new human DNA virus (TTV-like mini virus, TLMV) intermediately related to TT virus and chicken anemia virus. Arch. Virol. 145:979–993. 51. Tauxe, R. V. 2002. Emerging foodborne pathogens. Int. J. Food Microbiol. 78:31–41. 52. Traore, O., C. Arnal, B. Mignotte, A. Maul, H. Laveran, S. Billaudel, and L. Schwartzbrod. 1998. Reverse transcriptase PCR detection of astrovirus, hepatitis A virus, and poliovirus in experimentally contaminated mussels: comparison of several extraction and concentration methods. Appl. Environ. Microbiol. 64:3118–3122. 53. Vainio, K., K. Stene-Johansen, J. T. Oystein, A. L. Bruu, and B. Grinde. 2001. Molecular epidemiology of calicivirus infections in Norway. J. Med. Virol. 65:309–314. 54. Vinje´, J., and M. P. Koopmans. 2000. Simultaneous detection and genotyping of “Norwalk-like viruses” by oligonucleotide array in a reverse line blot hybridization format. J. Clin. Microbiol. 38:2595–2601. 55. Vinje´, J., H. Vennema, L. Maunula, C.-H. von Bonsdorff, M. Hoehne, E. Schreier, A. Richards, J. Green, D. Brown, S. S. Beard, S. S. Monroe, E. de Bruin, L. Svensson, and M. P. G. Koopmans. 2003. International collaborative study to compare reverse transcriptase PCR assays for detection and genotyping of noroviruses. J. Clin. Microbiol. 41:1423–1433. 56. Woody, M. A., and D. O. Cliver. 1995. Effects of temperature and host cell growth phase on replication of F-specific RNA coliphage Q beta. Appl. Environ. Microbiol. 61:1520–1526.