Inactivation and elimination of human enteric ... - Wiley Online Library

Journal of Applied Microbiology ISSN 1364-5072

ORIGINAL ARTICLE

Inactivation and elimination of human enteric viruses by Pacific oysters C. Mcleod1, B. Hay2, C. Grant2, G. Greening3 and D. Day1 1 School of Biological Sciences, Victoria University of Wellington, Wellington, New Zealand 2 AquaBio Consultants Ltd, Auckland, New Zealand 3 Institute of Environmental Science and Research Limited, Porirua, New Zealand

Keywords elimination, hepatitis A virus, inactivation, norovirus, Pacific oysters, poliovirus. Correspondence Darren Day, School of Biological Sciences, Victoria University of Wellington, PO Box 600, Wellington, New Zealand. E-mail: [email protected] Present address C. McLeod, South Australian Research and Development Institute, 33 Flemington Street, Adelaide, SA, Australia.

2009 ⁄ 0250: received 8 February 2009, revised 27 March 2009 and accepted 6 April 2009 doi:10.1111/j.1365-2672.2009.04373.x

Abstract Aims: To investigate the comparative elimination of three different human enterically transmitted viruses [i.e. hepatitis A virus (HAV), norovirus (NoV) and poliovirus (PV)] and inactivation of HAV and PV by Pacific oysters. Methods and Results: New Zealand grown Pacific oysters (Crassostrea gigas) were allowed to bioaccumulate HAV, NoV and PV. Samples of oyster gut, faeces and pseudofaeces were then analysed by using real-time RT-PCR to determine the amount of viral RNA and cell culture methods to identify changes in the number of plaque forming units. The results suggest that the majority of the PV present in the oyster gut and oyster faeces is noninfectious, while in contrast, most of the HAV detected in the oyster gut are infectious. Depuration experiments identified a large drop in the count of PV in the gut over a 23-h cleansing period, whereas the levels of HAV and NoV did not significantly decrease. Conclusions: Human enterically transmitted viruses are eliminated and inactivated at different rates by Pacific oysters. Significance and Impact of Study: The research presented in this article has implications for risk management techniques that are used to improve the removal of infectious human enteric viruses from bivalve molluscs.

Introduction Human enteric viruses replicate in the human alimentary canal with large quantities of virus shed with the faeces (Sobsey and Jaykus 1991; Racaniello 2001; Griffin et al. 2003; Atmar et al. 2008). Through the process of filterfeeding bivalve, molluscan shellfish concentrate these human enteric viruses, which are present in sewage that has entered the marine environment. Although there are many different types of human enterically transmitted viruses, epidemiologic studies have shown that shellfish transmit only a few of these (Gerba 1988; Sobsey and Jaykus 1991; Le Guyader et al. 2008). Outbreaks of disease caused by norovirus (NoV) and hepatitis A virus (HAV) have been frequently linked to the consumption of shellfish, whereas shellfish-related outbreaks of hepatitis E virus, astrovirus, poliovirus, adenovirus, Aichi virus and

rotavirus disease are less frequently reported, despite these viruses being commonly detected in shellfish (Chau et al. 1997; Anon 1998; Le Guyader et al. 2000, 2008; Lees 2000; Formiga-Cruz et al. 2002, 2003; Koizumi et al. 2004; Myrmel et al. 2004). There are several possible reasons why shellfish appear to preferentially transmit some viruses better than others; one of which is that shellfish may inactivate and eliminate viruses at different rates. Several risk management procedures, such as depuration and relaying, were introduced to reduce the numbers of microbial contaminants in oysters (Richards 1988). These processes rely on the elimination of contaminants from the oyster gut through normal feeding, digestion and excretion activities in contaminant-free seawater (Cook and Ellender 1986). Several studies on the depuration of viruses from Pacific oysters have been undertaken. Fletcher et al. (1991) undertook experiments in Pacific

ª 2009 The Authors Journal compilation ª 2009 The Society for Applied Microbiology, Journal of Applied Microbiology 107 (2009) 1809–1818

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Removal of human enteric viruses by oysters

oysters that had bioaccumulated cricket paralysis virus (CrPV). Results confirmed low levels of CrPV elimination (an approximate 20-fold reduction) after 5 days of cleansing in virus-free seawater (seawater temperature 17–20C) using an infectivity assay to measure virus quantities (Scotti et al. 1983; Fletcher et al. 1991). In contrast, a separate elimination study that also utilized cell culture techniques identified that poliovirus (PV) was almost completely removed from Pacific oysters between 48 and 96 h (Hoff and Becker 1969). Similarly, Dore and Lees (1995) tested Pacific oysters for F+ bacteriophage (utilizing the host cell Salmonella typhimurium WG49) and showed that 80% of the surrogate virus was eliminated from the oysters during a 48-h depuration period (seawater temperature 12C). Together, these studies suggest that the rate of virus elimination could differ with virus type ⁄ strain. However, it has been established that factors such as temperature, salinity, or whether a flow-through or recirculating seawater system has been used can affect the elimination rates of bacteria and viruses from shellfish, necessitating experiments exploring the concurrent depuration of multiple viruses from shellfish (Richards 1988, 2001). Studies on the concurrent depuration of several different viruses from species of shellfish other than the Pacific oyster (e.g. the simultaneous depuration of HAV and PV from mussels, Mytilus chilensis) have shown that different viruses are eliminated at distinct rates (Sobsey et al. 1987; Power and Collins 1989; Enriquez et al. 1992; Schwab et al. 1998; Nappier et al. 2008). However, studies to examine the elimination of multiple enteric viruses from Pacific oysters have not been undertaken. Information is also limited regarding inactivation of viruses by oysters through the digestive process. A study that examined the uptake of radiolabelled reovirus by rock oysters (Crassostrea glomerata) showed that the virus was less infectious after ingestion and digestion by the oysters (Bedford et al. 1978). This suggests that following the concentration of virus particles from the seawater, virus inactivation in the oyster gut and virus elimination via the faecal pathway may both play a role in the reduction of infectious virus particles within oysters. The aim of this study was to determine the relative inactivation and elimination rates of three different human enterically transmitted viruses, HAV, NoV and PV by New Zealand grown Pacific oysters. Our hypothesis was that inactivation and elimination rates of these viruses by Pacific oysters would be specific to the type of virus involved. This study describes the virus uptake and elimination experiments carried out to test this hypothesis. The relative contribution of the inactivation and elimination processes could be an important consideration for future research when trying to improve the 1810

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depuration rates of viruses from commercially farmed oysters. Materials and methods Viruses and cells Buffalo green monkey kidney (BGM) cells and foetal rhesus monkey kidney (FRhK-4) cells were propagated, and PV and HAV stock inocula were prepared from the BGM and FRhK-4 cells, respectively, as previously described (Greening et al. 2001). NoV is not readily culturable so a stock inoculum was prepared as a 10% (wt ⁄ vol) suspension of NoV-positive faecal specimen derived from an outbreak as previously described (Hewitt and Greening 2004; Seamer 2007). The NoV strain was genotyped as GII.4, a Farmington Hills-like strain (Zheng et al. 2006). Bioaccumulation and cleansing All Pacific oysters were harvested from an approved commercial oyster growing area in the Mahurangi Harbour, New Zealand, and were scrubbed to remove fouling organisms prior to experimentation. To mimic the natural tidal cycle and to maintain the entrained oyster digestive rhythms throughout the experiments, once in each 12-h period, the oysters were removed from the tanks for 3 h and resubmerged in synchrony with the tidal cycle of the harbour of origin. Prior to the experiments, oysters were acclimatized to these experimental conditions for at least 48 h. The temperature of the seawater during experiments was ambient (approximately 20C) and the salinity 30 ppt. Three groups of sixty oysters were bioaccumulated separately with each HAV (1Æ7 · 106 PCR amplifiable units per oyster), NoV (4Æ2 · 105 PCR amplifiable units per oyster) or PV (2 · 108 PCR amplifiable units per oyster) for 48 h in three tank systems containing 80 l of constantly circulating, aerated, unfiltered seawater. The initial titres were different for each virus investigated, because the maximum possible amount of virus was used to seed into the bioaccumulation tanks to ensure easy detection of virus in subsequent oyster samples. The oysters were fed a quarter of the total stock inocula with each change of seawater (three changes in 48 h) to give sufficient time to feed and bioaccumulate virus. After the bioaccumulation phase, eight oysters were removed from each tank to determine the initial amount of virus present in the oysters, and the remaining oysters were depurated in new 80-l tanks of virus-free unfiltered seawater for 23 h. During the cleansing period, the seawater was replaced once after 12 h, and UV irradiation was not undertaken. These experiments were undertaken once using multiple


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samples, each of which was analysed in triplicate. Eight oysters not exposed to virus were acclimatized for 48 h in unfiltered seawater and served as negative controls. Oyster tissue preparation At the end of the bioaccumulation period and 1Æ5, 16 and 23 h after depuration, eight oysters were randomly selected and removed for analysis. These oysters were rinsed and scrubbed under fresh running water, then the visceral mass was removed by severing the adductor muscle where it attaches to the valves. The oyster gut tissue (stomach, intestine and digestive diverticula) was then dissected out, and the surrounding gonad tissue discarded. Dissected gut tissue from two individual oysters was pooled to form one sample (to reduce the number of tests required), and four replicate samples were evaluated at each time point. All tissues were finely chopped using a scalpel then homogenized in a stomacher (Stomacher Lab-Blend 80; Fisher Scientific) for 2 min. The homogenate (60 mg) was resuspended in 120 ll phosphate-buffered saline (PBS) (pH 7Æ3, 160 mmol l)1 NaCl, 3 mmol l)1 KCl, 1 mmol l)1 KH2PO4, 8 mmol l)1 Na2HPO4) (Oxoid, Basingstoke, UK) and mixed thoroughly by vortexing for 2 min. The homogenate was sonicated (model FX10, Unisonics Pty, Ltd, Sydney, Australia) at room temperature for 3 min, then centrifuged at 2000 g for 3 min and the supernatant collected for analysis. The supernatant was then split into two equal portions (each containing extract from 30 mg of tissue), one for RNA extraction and PCR analysis and the other for infectivity testing by plaque assay. Oyster faeces and pseudofaeces preparation Oysters acquire their food by filter feeding using cilia on the gill surface to produce water currents to capture food particles in mucus on the gills. The gills transport food particles via a string of mucus forward to the labial palps, where particles are either guided to the mouth for ingestion, digestion and excretion as faeces, or dropped into the mantle cavity and rejected as pseudofaeces. Twenty hours into the bioaccumulation phase, three pooled samples (i.e. faeces from several oysters) of oyster faeces and concurrently three pooled samples of oyster pseudofaeces were separately collected using pipette from the bottom of each tank. Faeces were compact dark brown ribbons and were easily distinguishable from the lighter and looser formed pseudofaeces. Samples were washed by resuspending in 1 ml nuclease-free water and recovering by centrifugation at 2000 g for 1 min. The resulting faeces, or pseudofaeces, pellet (20–100 mg)


were weighed and resuspended in 120 ll PBS (pH 7Æ3, 160 mmol l)1 NaCl, 3 mmol l)1 KCl, 1 mmol l)1 KH2PO4, 8 mmol l)1 Na2HPO4) (Oxoid) then divided into two equal portions, one for RNA extraction and PCR analysis and the other for infectivity testing by plaque assay. Viral RNA extraction Virus extraction methods involving either dilution or concentration steps in the protocols have been frequently used for the recovery of viruses from shellfish tissues (Lewis and Metcalf 1988; Lewis et al. 1996; Lopez-Sabater et al. 1997; Green and Lewis 1999; Sunen and Sobsey 1999; Kingsley and Richards 2001; Greening and Hewitt 2008). When ‘concentration’ or ‘dilution’-based extraction methods are used, the amount of virus present in the extracts may not linearly reflect the amount of viruses originally present in the shellfish tissues. Therefore, a method that was previously shown to extract high quality RNA from rat brains was utilized for the extraction of viruses from shellfish tissues in our research (Baelde et al. 2007). Viral RNA was isolated from oyster tissue homogenate (30 mg), faeces (10–50 mg) and pseudofaeces (10–50 mg) by extraction with 750 ll of Trizol (Invitrogen, Life Technologies, Auckland, New Zealand). The aqueous phase containing RNA was recovered from the Trizol extract by the addition of chloroform (150 ll) and centrifugation at 10 000 g for 5 min. The aqueous phase was removed, 0Æ5 volumes of 100% ethanol (Sigma-Aldrich NZ Ltd., Auckland, New Zealand) was added, and the solution applied to a High Pure RNA Tissue Kit column (Roche Molecular Biochemicals Ltd, Mannheim, DEU, Germany). The bound RNA was washed according to the manufacturer’s instructions then eluted in 50 ll of nuclease-free water and stored at )70C until used. Five samples of the same PV-infected oyster digestive tract were individually extracted and assayed by real-time RT-PCR in triplicate to determine reproducibility of the extraction, and realtime RT-PCR methods and virus spiking experiments were undertaken to investigate recovery rate. Virus quantification by real-time PCR analysis Anticontamination procedures were followed for all RNA and DNA protocols. RNAse-free reagents and procedures were used to minimize contamination. Onestep RT-PCR amplification was performed using Platinum Quantitative RT-PCR Thermoscript One-Step System (Invitrogen Corporation, CA, USA) on a RotorGene 3000 real-time cycler (Corbett Life Science, Sydney, Australia). Primer sets are listed in Table 1. The PV and


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Table 1 HAV, NoV and PV primer and probe sequences

Primer or probe

Sequence (5¢–3¢)

PV-for PV-rev PV-probe HAV-for HAV-rev HAV-probe NoV-for NoV-rev NoV-probe

GGC CCC TGA ATG CGG CTA AT ACC GGA TGG CCA ATC 6-FAM d(CGG ACA CCC AAA GTA GTC GGT TCC G) BHQ-1 GGG TAA CAG CGG CGG ATA T TTA AGC CTA AAG ACA GCC CCG 6-FAM d(TCA ACG CCG GAG GAC TGA CTC TCA TC) BHQ-1 AGT TGA TGT CCT TAC TGG GAG AGG TGA CTA ACT TGC TGA TTT TGC TGT AGA 6-FAM d(CGC ACT CCA CGG CCC AGC A) BHQ-1

PCR product size (bp)

Genome region amplified

190

5¢ noncoding region (bases 449–639)

120

5¢ noncoding region (bases 451–571)

72

Nonstructural polyprotein (polymerase gene) (bases 4886–4958)

BHQ-1, Black hole quencher; 6-FAM, 5¢-Fluorescein-CE Phosphoramidite; for, forward primer; rev, reverse primer; PV, poliovirus; HAV, hepatitis A virus; NoV, norovirus.

HAV primers and probes have been previously described (Donaldson et al. 2002; Hewitt and Greening 2004). GII.4 NoV real-time RT-PCR primers were designed in the RNA pol region using Primer Express v 1Æ5 (Applied Biosystems Ltd). The simplex reactions (25 ll) contained 2Æ5 ll of RNA template, 20 U RNase inhibitor (RNaseOUT; Invitrogen Corporation), 0Æ5 ll reverse transcriptase-Taq polymerase enzyme mix (Invitrogen Corporation), forward and reverse primers (0Æ6 lmol l)1 PV, 0Æ4 lmol l)1 NoV and HAV), and Taqman probe (0Æ25 lmol l)1 PV and 0Æ2 lmol l)1 NoV and HAV) diluted in the supplied amplification buffer (Invitrogen Corporation). Following an initial 30-min reverse transcription step at 60C, and a denaturation step at 95C for 5 min, amplification was achieved by a two-step cycling protocol of denaturation at 95C for 20 s and annealing ⁄ extension at 60C (PV) or 59C (NoV and HAV) for 60 s over 40 cycles. Data were analysed using the RotorGeneTM software to calculate cycle threshold (Ct) values. Data were transformed to real-time PCR amplifiable units per milligram of oyster tissue (PAU mg)1) using external standard curves generated from log dilutions (100–108) of HAV, NoV and PV stock preparations of purified RNA (Wong and Medrano 2005). The limit of detection for the real-time PCR assays in shellfish was 0Æ67 PAU mg)1 oyster tissue. All samples were assayed in triplicate, and the mean number of PAU mg)1 oyster tissue, or faeces and pseudofaeces material, determined. To determine whether components present in the sample extracts contained inhibitory material that would interfere with estimation of viral titres, blank sample extracts of oyster gut, faeces, and pseudofaeces were spiked with known counts of PV, NoV and HAV standards and analysed by real-time RT-PCR in parallel with the analysis of the virus standards resuspended in nuclease-free water. 1812

Virus quantification by plaque assay For the PV cell culture infectivity assay, suspensions of BGM cells in agar were prepared, and samples (oyster tissue, faeces and pseudofaeces samples) assayed as previously described (Simmonds et al. 1982). For the infectious HAV assay confluent layers of foetal rhesus monkey kidney (FRhK-4), cells were grown, and oyster tissue samples assayed as previously described (Cromeans et al. 1989). All samples were tested in triplicate, and the mean number of plaque forming units per milligram (PFU mg)1) of oyster tissue or faecal and pseudofaecal material determined. Blank samples of oyster gut, faeces and pseudofaeces were spiked with known counts of either PV or HAV to determine whether components present in the sample extracts contained inhibitory material that would interfere with the estimation of infectious viral titres (as determined by plaque assay). Data analysis Descriptive data analyses (e.g. generation of mean values and graphs) were performed using Microsoft Excel (Microsoft Excel 2004 for Mac ver. 11.1). In order to satisfy normality and constant variance assumptions, the PCR and plaque assay data were transformed by the log 10(x + 1) function prior to statistical analyses being performed. Statistical analyses were performed using simple linear regression and one-way anova on spss ver. 11.5.0 for Windows (SPSS Inc., Chicago, IL). Standard errors for ratios such as the infectivity quotient were calculated using a formula derived from a delta theorem via a Taylor series expansion about the mean (kindly derived by Dr Shirley Pledger, School of Biological Sciences, Victoria University of Wellington) (Seber 1982). These values take into account the number of measurements made and the error associated with each measurement. Results with P values of 0Æ98, and efficiencies >0Æ83 being consistently achieved in each experiment. PV samples that had Ct values below 34 and HAV and NoV samples with Ct values below 35 were considered positive. PCR product was never observed in control amplifications lacking template. RNA extraction and real-time RT-PCR analysis of the control oysters confirmed that none of the oysters were initially contaminated with HAV, PV or NoV. Parallel PCR analysis of blank oyster gut, faeces and pseudofaeces RNA extracts were spiked with known counts of PV, NoV and HAV standards, and the virus standards resuspended in nuclease-free water demonstrated that PCR inhibition by oyster matrix was not significant in these experiments.


contained inhibitory material that would interfere with estimation of infectious viral titres. For PV, the expected titres were observed indicating that no inhibitory or toxic substances were present in the dilutions. However, for HAV, the oyster gut matrix contained substances that resulted in a lower number of plaques forming than would have been expected possibly because of the sequestration of virus by oyster gut matrix, resulting in reduced susceptibility of the FRhK-4 cells to infection or to decreased infectivity of the virus. Dilution of samples obtained from the gut alleviated the problem; nonetheless, our reported values probably underestimated the number of HAV plaque-forming units (PFU) present in a sample, as complete elimination of ‘plaque inhibitory’ material by dilution could not be achieved. Faeces and pseudofaeces extracts were found to completely inhibit the formation of plaques in FRhK-4 cell monolayers, and so culture analysis of HAV in these samples was not possible. BGM and FRhK-4 control plates were inoculated with culture medium, negative control oyster gut tissue, or blank faeces or pseudofaeces samples, never contained PFU. Elimination of HAV, NoV and PV from the oyster gut Oysters took up HAV, NoV and PV so that by the end of the bioaccumulation period (i.e. cleansing period 0 h) viruses were readily detectable by real-time RT-PCR and by plaque assay (except NoV) in oyster gut tissue (Table 2). Cleansing for 23 h resulted in the elimination of 93% of PV detected by RT-PCR and 71% by plaque assay from the oyster gut (Table 2). Regression analysis showed that the loss of PV PAU and PFU from the gut over the 23-h period was significant (P = 0Æ001 and P = 0Æ009 respectively). In contrast to PV, no significant reduction in the counts of HAV PAU or PFU (P > 0Æ153) (Table 2) was observed in oyster gut tissue after 23h of cleansing. Oneway anova showed that the observed increase in HAV PAU in oyster gut samples following cleansing for 1Æ5 h (Table 2) was insignificant (P = 0Æ110). Lower levels of NoV (when compared with PV and HAV) were found in oyster gut samples after bioaccumulation, which is consistent with the initial levels of virus titre introduced (Table 2). Cleansing for 23 h did not result in a significant reduction of NoV PAU from the oyster gut tissue (P = 0Æ241) (Table 2).

Plaque assay matrix inhibition studies

Inactivation of viruses in the oyster gut, faeces and pseudofaeces

Blank samples of oyster gut, faeces and pseudofaeces were spiked with known counts of either PV or HAV to determine whether components present in the sample extracts

To assess the level of virus inactivation, the ratio of infectious viral particles (PFU mg)1 tissue) to the number of PCR amplifiable units (PAU mg)1 tissue) was calculated.


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Virus

Cleansing Period (h)

PAU mg)1 gut tissue ± SEM

PFU mg)1 gut tissue ± SEM

*IQ ± SEM

PV PV PV PV HAV HAV HAV HAV NoV NoV NoV NoV

0 1Æ5 16 23 0 1Æ5 16 23 0 1Æ5 16 23

117Æ52 74Æ50 14Æ67 8Æ50 121Æ30 220Æ47 42Æ22 110Æ80 27Æ46 36Æ46 34Æ85 39Æ36

5Æ75 3Æ50 1Æ07 1Æ65 >83Æ90 >36Æ64 >41Æ44 >47Æ12 NA NA NA NA

0Æ08 0Æ10 0Æ21 0Æ14 0Æ66 0Æ16 1Æ29 0Æ44 NA NA NA NA

± ± ± ± ± ± ± ± ± ± ± ±

59Æ27 21Æ84 14Æ16 4Æ75 33Æ02 42Æ30 20Æ08 11Æ40 5Æ81 6Æ04 4Æ32 5Æ64

± ± ± ± ± ± ± ±

1Æ57 1Æ13 0Æ71 0Æ92 23Æ70 12Æ42 10Æ99 12Æ01

± ± ± ± ± ± ± ±

0Æ01 0Æ01 0Æ19 0Æ02 0Æ06 0Æ01 0Æ34 0Æ03

Table 2 Poliovirus, hepatitis A virus and norovirus levels in the oyster gut following uptake (48 h) and cleansing (23 h) by Pacific oysters

Hepatitis A virus, norovirus and poliovirus levels following uptake and cleansing by Pacific oysters. PV, poliovirus; HAV, hepatitis A virus; NoV, norovirus; PAU, PCR amplifiable units; PFU, plaque forming units; SEM, standard error of the mean; NA, Not applicable because currently there is no culture method for NoV available. *Infectivity quotient (IQ) is the mean ratio of infectious virus (PFU mg)1 tissue) and PCR amplifiable units (PAU mg)1 tissue).

The calculated infectivity quotient (IQ) is thus a measure of the extent of viral inactivation, such that a low value for IQ indicates that the virus is of low quality relative to the initial inoculum, and an IQ of 1Æ0 that the virus is as infectious as the initial inoculum. Bioaccumulation of PV by oysters resulted in a large drop in the IQ from 1Æ0 to 0Æ08 ± 0Æ01 (Table 2), indicating that most of the virus present in the oyster gut was not infectious. While cleansing eliminated the majority of PV from the oyster gut, no significant (P = 0Æ273) further loss of virus infectivity was detected over the 23-h cleansing period (Table 2). In comparison with PV, bioaccumulation of HAV by oysters resulted in a smaller drop in the IQ from 1Æ0 to 0Æ66 ± 0Æ06 (Table 2), indicating that most of the HAV detected in the oyster gut after bioaccumulation was infectious. At the end of the cleansing period (23 h), a large proportion of HAV in the oyster gut was still infectious as indicated by the IQ (Table 2). It was not possible to calculate an IQ for NoV, as currently there is no culture method for testing NoV in oysters. Separate oyster faeces and pseudofaeces samples were taken after 20 h of PV bioaccumulation by oysters and

tested for both PV PAU and PFU. The majority of the PV in the faeces was found to be noninfectious with an IQ of 0Æ15 ± 0Æ03 (Table 3). In contrast to the faeces, the pseudofaeces had a higher IQ of ‡ 1Æ0, indicating that most of the PV present was infectious, and they contained approximately 3-fold more PV PFU than the faeces (P = 0Æ009). No information was obtained on the infectivity of HAV in faeces and pseudofaeces, because the faecal samples were inhibitory to plaque formation in FRhK-4 cells, or NoV infectivity as no culture method is currently available. Discussion Elimination of HAV, NoV and PV from the oyster gut We describe the relative elimination and inactivation of three different human enterically transmitted viruses (HAV, NoV and PV) by Pacific oysters. Following a 23-h cleansing period, oysters that had bioaccumulated PV showed a significant decrease in the count of PV in the gut, but oysters that bioaccumulated NoV and HAV

Virus

Sample type

PAU mg)1 ± SEM

PFU mg)1 ± SEM

*IQ ± SEM

PV PV

Faeces Pseudofaeces

3391Æ02 ± 1192Æ18 1036Æ45 ± 703Æ19

368Æ13 ± 6Æ42 1120Æ23 ± 161Æ49

0Æ15 ± 0Æ03 ‡1Æ0

Table 3 Poliovirus levels in Pacific oyster faeces and pseudofaeces after 20-h bioaccumulation

PV, poliovirus; PAU, PCR amplifiable units; PFU, plaque forming units; SEM, standard error of the mean. *Infectivity quotient (IQ) is the mean ratio of infectious virus (PFU mg)1 tissue) and PCR amplifiable units (PAU mg)1 tissue).

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showed no significant loss of virus. While there was no significant loss of HAV over 23 h, the PCR results were quite variable, likewise the PCR results for PV following 16-h depuration were also variable. Other studies undertaken in our laboratory and by other researchers have shown that there is large variability in virus uptake between individual shellfish (e.g. up to 100-fold difference) (Seraichekas et al. 1968; Seamer 2007). This, in combination with the sample size used in these experiments (e.g. n = 4), may have contributed to the variability in the results. Other published work on virus depuration in shellfish also indicates that a large percentage of PV ingested by oysters is eliminated rapidly (Mitchell et al. 1965; Hoff and Becker 1969; Di Girolamo et al. 1975). Also in accordance with our results, Schwab et al. (1998) showed that NoV depurated slowly from Crassostrea virginica (eastern oysters), with no significant decrease in titres following 48-h depuration. Other researchers also demonstrated relatively slower depuration rates for HAV than for PV in M. chilensis (mussels) and C. virginica (Sobsey et al. 1987; Enriquez et al. 1992), which is comparable to our results that show no decrease in HAV or NoV titre in Pacific oysters but significant loss of PV. Data from a variety of shellfish growing waters over a 7-year period in the USA indicated average level of 0Æ2 viruses ml)1 seawater (Metcalf 1982). Levels in New Zealand waters are probably lower than this, in the range of 1 · 10)5 to 1 · 10)3 viruses ml)1 seawater (Personal communication, G. Lewis, University of Auckland, New Zealand). In order to facilitate the detection of viruses, much higher levels of virus than that encountered in shellfish growing waters were used in these bioaccumulation experiments (e.g. 80 NoV PAU ml)1, 319 HAV PAU ml)1 and 37 500 PV PAU ml)1). It has been suggested that the ability of shellfish to depurate a large percentage of virus in a short period (e.g. PV) may be an artefact of the high concentrations used in such studies (Metcalf et al. 1979), and it would be informative to undertake further research using various virus titres in the overlaying waters. HAV, NoV and PV have similar physical properties e.g. size and shape, but they differ in composition and structure at a molecular and biochemical level (e.g. differing capsid protein conformation and cell surface receptors) (Kapikian et al. 1996; Green et al. 2001; Hollinger and Emerson 2001; Racaniello 2001, 2006). If virus elimination from oysters was a function of particle size alone, similar patterns of HAV, NoV and PV removal would be expected, but the results of our investigation suggest that different viruses of the same size can have varying patterns of elimination. Recent studies have concluded that NoV binds to specific carbohydrates (similar to human


blood group antigens) within oyster digestive tract tissues (Tian et al. 2006, 2008). It is possible that the different surface properties of PV, HAV and NoV may lead to distinct interactions with these carbohydrates, possibly accounting for the different elimination patterns observed in this study. Inactivation of viruses In this study, plaque assay experiments indicated that most PV present in the oyster gut and oyster faeces was no longer able to replicate in cultured cells immediately after uptake by the oyster. In contrast, most HAV (>69%) within the oyster gut immediately after uptake was able to infect and replicate in cultured cells, raising the possibility that HAV may be more resistant than PV to inactivation via ingestion and digestive processes. The decrease in infectivity of PV and to a lesser extent HAV in our study, after ingestion by oysters suggests that inactivation, as well as elimination, plays a role in the removal of infectious viruses from oysters. Unwanted particles are physically rejected prior to ingestion by the oyster from the gills and labial palps as pseudofaeces (Bernard 1974). The significantly higher concentrations of infectious PV found in pseudofaeces compared with the gut and faeces samples in this study suggest that particle sorting along with the gill surface does not inactivate PV while passage and inactivation through the oyster gut does. It is possible that transit through the oyster gut results in modification of the virion such that productive association of host cell receptor is blocked in cell culture, preventing plaque formation. Such modified virions potentially may be infectious to humans when ingested during consumption of shellfish. Practical implications Relaying of potentially contaminated oysters into clean growing areas is currently undertaken in order to minimize the risk of viruses being present in the oysters. The practice of relaying creates the potential for infective virus associated with the contaminated oysters to be transferred to uncontaminated oysters growing nearby in the relay area. Viruses can be potentially associated with the faeces or pseudofaeces excreted from the contaminated oysters or freely associated with the oyster valves and body. The findings in this study demonstrate that oyster faeces contain more PV than the pseudofaeces, but the oyster pseudofaeces contain a significantly higher amount of infectious PV than the faeces. This suggests that the oyster pseudofaeces may present a higher source of risk than the oyster faeces as a vehicle for the transfer of virus


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from contaminated to uncontaminated oysters in relaying situations. Our results indicate that cleansing of heavily contaminated oysters over short time frames is unlikely to lead to a significant reduction of HAV or NoV. This is corroborated by the occurrence of NoV-related gastroenteritis in people consuming depurated oysters (Grohmann et al. 1981; Gill et al. 1983; Conaty et al. 2000). Results from other research have shown that the levels of NoV and HAV in oysters do slowly decrease when oysters are placed in uncontaminated seawater over much longer periods e.g. up to 8 weeks (Greening et al. 2003; Kingsley and Richards 2003). While direct elimination of NoV and HAV from oysters via defecation is not effective over short time frames, our study demonstrates that inactivation of viruses may also play a role in the reduction of infectious virus particles. Future studies focused on increasing the inactivation rate of viruses following ingestion by oysters may lead to the possibility of depuration being a viable option for the removal of viruses from commercially farmed oysters. Acknowledgements This work was funded by the Public Good Science Fund of New Zealand Ministry of Science, Research and Technology, Contracts C03X0301 and C03X0204. We especially thank Jim Dollimore of Biomarine Limited for provision of oysters and the New Zealand Food Safety Authority for additional funding. HAV (cytopathic strain HM-175) and foetal rhesus monkey kidney (FRhK-4) cells were kindly provided by Dr M.D. Sobsey (University of North Carolina, Chapel Hill). ESR Postgraduate Scholarship supported CM. References Anon (1998) Outbreaks of gastroenteritis in England and Wales associated with shellfish: 1996 and 1997. Commun Dis Rep CDR Wkly 8, 21, 24. Atmar, R.L., Opekun, A.R., Gilger, M.A., Estes, M.K., Crawford, S.E., Neill, F.H. and Graham, D.Y. (2008) Norwalk virus shedding after experimental human infection. Emerg Infect Dis 14, 1553–1557. Baelde, H., Cleton-Jansen, A., van Beerendonk, H., Namba, M., Bovee, J. and Hogendoorn, P. (2007) High quality RNA isolation from tumours with low cellularity and high extracellular matrix component for cDNA microarrays: application to chondrosarcoma. J Clin Path 54, 778–782. Bedford, A.J., Williams, G. and Bellamy, A.R. (1978) Virus accumulation by the rock oyster Crassostrea glomerata. Appl Environ Microbiol 35, 1012–1018.

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