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Effects of high-pressure processing on murine norovirus-1 in oysters (Crassostrea gigas) in situ
Food Control 20 (2009) 992–996
Contents lists available at ScienceDirect
Food Control journal homepage: www.elsevier.com/locate/foodcont
Effects of high-pressure processing on murine norovirus-1 in oysters (Crassostrea gigas) in situ Dan Li1, Qingjuan Tang1, Jingfeng Wang, Yuming Wang, Qin Zhao, Changhu Xue * College of Food Science and Engineering, Ocean University of China, No. 5, Yushan Road, Qingdao, Shandong Province 266003, PR China
a r t i c l e
i n f o
Article history: Received 3 July 2008 Received in revised form 20 November 2008 Accepted 25 November 2008
Keywords: High-pressure processing Murine norovirus-1 Oyster RT-PCR
a b s t r a c t We assessed the effects of high-pressure processing (HPP) at 200–400 MPa on murine norovirus-1 (MNV1)-contaminated oysters by using a flow-through seawater system. Plaque assays demonstrated that a 5-min 400-MPa treatment at 0 °C inactivated MNV-1 within oysters to undetectable levels. No correlation was found between MNV-1 RNA detection by reverse transcription-PCR (RT-PCR) and the infectivity determined by the plaque assay before and after pressure exposure. Pretreatment with proteinase K and RNase A enabled the differentiation between infectious and HPP-inactivated MNV-1 by RT-PCR, indicating that HPP might subtly alter the viral capsid proteins but that the RNA remains protected. Ó 2008 Published by Elsevier Ltd.
1. Introduction Marine organisms consumed as seafood, particularly bivalve shellfish such as oysters, can bioaccumulate microbial pathogens at high levels from surrounding marine and estuarine waters (Murchie et al., 2005). Thorough cooking is an absolutely safe way of eliminating pathogens; however, this technique also alters the organoleptic qualities of shellfish. Hence, the consumption of raw or minimally cooked oysters is a common custom practiced worldwide. An alternate technology, high-pressure processing (HPP), has recently been developed as a potential means for reducing pathogens within raw shellfish. With the use of this technology, the appearance, flavor, texture, and nutritional qualities of unprocessed foods like oysters can be retained. In addition to enhancing the safety and extending the shelf-life of seafood, HPP treatment has the additional advantage of shucking shellfish, thus rendering this technology particularly useful to the shellfish-processing industry and consumers alike. HPP has become a commercial success in the American and European markets with the availability of several ready-to-eat meat products and seafoods, as reviewed in Grove et al. (2006) and Murchie et al. (2005). As the single most common cause of outbreaks as well as sporadic cases of acute gastroenteritis, noroviruses (NoVs) are a great threat to the safety of edible shellfish worldwide (Beuret, Baumgartner, & Schluep, 2003; Guyader, Haugarreau, Miossec, Dubois, & Pommepuy, 2000; Nishida et al., 2003; Noda, Fukuda, & Nishio, * Corresponding author. Tel.: +86 532 82032468; fax: +86 532 82032468. E-mail address: [email protected] (C. Xue). 1 These authors contributed equally to this work. 0956-7135/$ - see front matter Ó 2008 Published by Elsevier Ltd. doi:10.1016/j.foodcont.2008.11.012
2008). NoVs were first detected by immune electron microscopy during the 1968 outbreak of gastroenteritis in Norwalk, Ohio, US; the lack of suitable animal models and the inability to propagate NoVs in cell cultures have hampered further study on NoVs (Duizer et al., 2004). Straub et al. (2007) have described a highly differentiated 3D cell culture model that supports the natural growth of human NoVs. However, this system is rather complex; therefore, surrogate viruses that share common pathological and molecular features with human NoVs are employed instead. Feline calicivirus (FCV) was almost exclusively used as a surrogate virus in previous studies (Doultree, Druce, Birch, Bowden, & Marshall, 1999; Thurston-Enriquez, Haas, Jacangelo, Riley, & Gerba, 2003). Recently, a NoV infecting mice, namely, murine norovirus-1 (MNV-1), has been identified (Karst, Wobus, Lay, Davidson, & Virgin, 2003; Wobus, Thackray, &Virgin, 2006; Wobus et al., 2004). A report investigating the applicability of MNV-1 and FCV as surrogates for human NoV in studies on the stability and inactivation of NoV demonstrated that MNV-1 was more acid-tolerant than FCV, thus making it a more suitable surrogate for human NoV (Cannon et al., 2006). Currently, RT-PCR is the most commonly used detection method because of its rapidness and high sensitivity. However, since it cannot discriminate between inactivated and infectious viral particles, it may provide potentially false-positive results for foods that have actually been properly treated (Richards, 1999). The inclusion of a pretreatment with enzymes prior to RNA extraction has been investigated to solve this problem and has proved effective in the case of human hepatitis A virus, vaccine poliovirus 1, and feline calicivirus inactivated by ultraviolet light, hypochlorite, or heating (Nuanualsuwan & Cliver, 2002). Baert et al. (2008) examined MNV1 following heat exposure but observed no correlation between the
D. Li et al. / Food Control 20 (2009) 992–996
number of infectious particles and viral genomes after heat treatment, regardless of the presence or nature of the enzyme treatment. However, whether MNV-1 inactivated by HPP can be detected by this method has not been reported yet. In this study, we aimed to examine the efficacy of HPP in inactivating MNV-1 within shellfish without altering the organoleptic qualities of oysters, and investigated the correlation between the number of infectious particles and viral genomes after HPP along with enzyme treatment. Oysters were bioaccumulated with MNV-1 in a flow-through seawater system before the inactivation by HPP; MNV-1 was detected by both plaque assay and RT-PCR. Pretreatment with proteinase K and RNase A was also performed. 2. Materials and methods 2.1. Virus and shellfish Working stocks of MNV-1 were prepared using confluent monolayers of RAW 264.7 mouse monocytes/macrophages (Shanghai Cell Bank, Shanghai, China) cultured in high-glucose Dulbecco’s modified Eagle medium (DMEM; Gibco-Invitrogen Co., Grand Island, NY, USA) supplemented with 10% fetal bovine serum (FBS; Hyclone Co., Logan, Utah, USA), 100 U/ml of penicillin (North China Pharmaceutical Group Corporation, Shijiazhuang, China), and 100 lg/ml of streptomycin sulfate (Amresco, Solon, OH, USA). After lysis, the lysate was freeze-thawed twice, centrifuged at 3500g for 10 min, and filtered through a membrane with a pore size of 0.22 lm. The MNV-1 stocks were stored at 80 °C. In order to determine the titer of the MNV-1 stock, plaque assays were performed in triplicate as described by Kingsley, Holliman, Claci, Chen, and Flick (2007) by using serially 10-fold diluted viral stocks in DMEM and assayed using confluent monolayers of RAW 264.7 mouse monocytes/macrophages in 6-well plates (Corning Inc., Corning NY, USA). Inoculation with 0.5 ml of MNV-1 was carried out for 2 h at 37 °C; this was followed by overlay with 2 ml of DMEM containing 1% low-melting-point agarose (BBI Co., Ltd., Boston, MA, USA) with 2% FBS. After 48 h of incubation, plaques were visualized by staining with 0.03% neutral red (Sigma Chemical Co., St. Louis, MO, USA) for 30 min at room temperature. The titer of the MNV-1 stock that was used to contaminate the oysters was approximately 2 1011 plaque forming units (PFUs)/ml. Pacific oysters (Crassostrea gigas) were harvested from an approved aquafarm in Jiaozhou Bay, Shandong Province, PR China. Considering the fact that the shellfish matrix composition, i.e., its fat content, water content, and salt concentration etc., can affect the pathogen inactivation rates of HPP and inhibit detection by RT-PCR (Claci, Meade, Tezloff, & Kingsley, 2005), oysters were bioaccumulated with MNV-1 in a flow-through seawater system as according to a method described by Kingsley, Hoover, Papafragkou, and Richards (2002) with a few modifications. Eight hours prior to viral accumulation, 24 commercial-size oysters were placed in an accumulation tank containing artificial seawater. The salinity of the seawater was 3.3%. An aquarium air pump was used to supply sufficient amounts of oxygen for the shellfish. MNV-1 (2 1011 PFU) was added to 4 l of double-distilled Milli-Q water (Millipore Corp., Bedford, MA, USA), which was continually mixed at 4 °C. Peristaltic pumps were adjusted to combine 4 ml of the viral suspension with 200 ml of UV-treated artificial seawater per minute in a cytostir vessel. After 16 h, the oysters were divided into 4 groups of 6 oysters each. 2.2. High-pressure treatment The oyster samples (6 oysters per treatment group) were transferred into aluminum foil pouches (30 cm 30 cm) containing an
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ice–water mixture and sealed using a DZQ-600 vacuum packaging machine (Zhangjiagang Deshun Machinery Co., China). An overpack pouch was sealed over the inner pouch. Pressurization of the oyster samples was carried out using a high-pressure food processor (Ningbo Branch, Institute of China Enginery, Ningbo, China). The range of the HPP treatment condition was chosen on the basis of the conditions generally maintained in commercial processing (Murchie et al., 2005). All samples were pressurized at 200, 300, and 400 MPa for 5 min using di-n-octyl sebacate (DOS) medium. The pressure vessel had a capacity of 5 l and an internal diameter of 120 mm. The rate at which the final pressure was attained was approximately 100 MPa/min, and pressure release was almost instantaneous. Ice bags were used during the shipment to ensure that the samples were maintained at a low temperature. Negative (uncontaminated) and positive (contaminated and nonpressurized) controls were processed in a manner similar to the HPP-treated oysters, except that they did not undergo the HPP treatment. 2.3. Viral extraction and plaque assays The stomachs and digestive diverticula of the oysters (61 g from each of the 6 oysters per treatment group) were removed by dissection, placed in 50-ml conical tubes that contained 6 ml of glycine buffer (0.1 M glycine, 0.3 M NaCl; pH 9.5, 4 °C), and homogenized by a high-performance disperser (IKA T18 basic ULTRA-TURRAXÒ, German) at 18,000 rpm for approximately 3 min. An ice bath was used to maintain the dispersion temperature below 10 °C. The homogenized extracts were centrifuged at 12,000g for 15 min at 4 °C in a Jouan MR23i centrifuge (Thermo Scientific). The resultant supernatant was neutralized with 2 N HCl and mixed with penicillin (1000 U/ml) and streptomycin sulfate (1000 lg/ ml); this was followed by incubation at 4 °C for 1 h. Tenfold serial dilutions were made in DMEM, and plaque assays were performed in triplicate using the same method as that used for determining the titer of the MNV-1 stock. 2.4. Residual infectivity No interference was observed in the cell culture and MNV-1 plaque assay until the viral extraction was diluted at least 100 times in DMEM. The detection limit of the MNV-1 assay was consequently raised to 200 PFU MNV-1/oyster. Thus, a residual infectivity test was carried out to verify whether all viral particles in the suspension were inactivated. The viral extraction (0.1 ml) was inoculated into confluent monolayers of RAW 264.7 mouse monocytes/macrophages in 100-mm tissue culture dishes and incubated at 37 °C for 2 h. After 48 h, the viral extraction was observed in order to determine whether it could induce a cytopathic effect in the mouse monocytes/macrophages. After 48 h, the lysate was freeze-thawed twice and diluted tenfold. Plaque assays were performed using the same method as that stated before. 2.5. Composite enzymatic pretreatment RNase A (BBI Co., Ltd., Boston, MA, USA) was diluted in TrisEDTA buffer (1.0 M Tris–HCl, 0.1 M EDTA; pH 8.0) and maintained at 20 °C. Proteinase K (BBI Co., Ltd., Boston, MA, USA) was dissolved in 0.01 M PBS and prepared freshly for each experiment. Both 40 U of proteinase K and 200 ng of RNase A were added to 100 ll of the viral extraction and incubated at 47 °C for 30 min. To 10 ll of RNA isolated from MNV-1, 10 ng of RNase A was added, and this solution served as a positive control for RNase A activity. In order to test the enzyme activity of proteinase K, pretreatments were performed with both RNase A and RNase A combined with proteinase K.
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2.6. RNA extraction and RT-PCR Regardless of enzyme pretreatment, 100 ll of viral extraction and 500 ll of TRIzol Reagent (Invitrogen, Carlsbad, CA, USA) were vigorously mixed with each sample by repipetting. After incubation for 5 min at room temperature, 100 ll of chloroform was added to each sample. The samples were then vortexed vigorously for 30 s, incubated at room temperature for 5 min, and centrifuged at 12,000g for 5 min. The upper aqueous layer, which contained the RNA, was precipitated by the addition of 0.5 volume of isopropanol for 5 min at room temperature; this was followed by centrifugation at 12,000g for 5 min. The resulting white pellets were washed with cold 75% ethanol, and each pellet was then resuspended in 50 ll of RNase-free water. RT-PCR amplification of the MNV-1 sequences was performed in a two-step format. The primer 3311 50 -GATTGGTAGT GCCGTTGTG-30 and the antisense primer 3732 50 -TGACCTGCTGAAGGGAAG-30 were used. Reverse transcription of all viral RNA was performed at 37 °C for 60 min using MMLV reverse transcriptase (Promega, Madison, WI, USA). We performed 40-cycle PCR reactions that involved annealing at 55 °C for 1 min, extension at 72 °C for 1 min, and denaturation at 95 °C for 30 s. For the final cycle, the annealing time was extended to 2 min, and the extension was performed for 10 min. The final concentration of each of the primers used was approximately 0.25 lM. The PCR products were separated by 1% agarose gel electrophoresis, stained with ethidium bromide, and visualized by UV light. 2.7. Analysis of data The MNV-1 titers were determined in triplicate. The average PFU obtained was calculated, and the standard error was expressed logarithmically. 3. Results At the end of the bioaccumulation period, the MNV-1 titers in the oysters in three separate trials were determined to be 1.56 104, 1.44 104, and 1.12 104 PFU per group. Since it is commonly believed that less than 100 norovirus particles are sufficient to cause illness (Caul, 1996), our results confirmed that shellfish have the ability of bioaccumulating viruses to a level far beyond the infectious dose. HHP treatment, with pressures ranging from 200 to 400 MPa and applied in 100-MPa increments, was administered to three treatment groups for 5 min at approximately 0 °C. The average titer obtained at each pressure is displayed in Table 1. The onset of inactivation was observed at 200 MPa, and treatment with 400 MPa was sufficient to inactivate MNV-1 within the oysters such that the viral load is undetectable. To confirm whether the viral load is undetectable only because of the inactivation of MNV-1, a residual infectivity test was performed. The results confirmed that
Table 1 Residual infectivity of MNV-1 detected after HHP treatment. Pressure (MPa)
Average log10 PFU (SEa)
Log10 PFU (SE a) for trial no. 1
2
3
0 200 300 400
4.13 (0.06) 3.56 (0.05) 3.18 (0.08) ND
4.19 (0.05) 3.57 (0.16) 3.18 (0.14) ND
4.15 (0.10) 3.50 (0.40) 3.08 (0) ND
4.04 (0.08) 3.62 (0.23) 3.28 (0.14) ND
Rate of reduction 0 13.8% 23.0% 100%b
ND: not detected. a SE: standard error. b Since the detection limit of the plaque assay is 200 PFU MNV-1/oyster, this result was verified by a residual infectivity test.
MNV-1 was indeed inactivated by the 400-MPa treatment. Moreover, the HPP treatment used in this study (200–400 MPa) did not affect the visual appearance of oysters and also facilitated their shucking. Viral RNA extraction was performed on all shellfish extracts. All controls (non-seeded oysters) tested negative for MNV-1 by RTPCR, indicating that all oysters were originally free of MNV-1. The 439-bp major amplification product was analyzed for MNV-contaminated samples in all oyster groups (Fig. 1; lanes 2, 4, 6, and 8). No genomic copies were detected after the addition of 10 ng of RNase A to 10 ll of RNA isolated from MNV-1, thus proving that RNase A was able to degrade free viral RNA. Pretreatment with both RNase A and proteinase K yielded mostly negative RT-PCR results for the 400-MPa inactivated MNV-1; however, positive results were obtained when only RNase A was used. This indicated that proteinase K effectively attacked the capsid (data not shown). After pretreatment with 200 ng of RNase A and 40 U of proteinase K at 47 °C for 30 min, the RT-PCR results of HPP-inactivated MNV-1 tend to present fainter band density along with the increase in the processing pressure (Fig. 1; lanes 5, 7, and 9), whereas those of the similarly treated infectious viruses remained positive (Fig. 1, lane 3).
4. Discussion Kingsley et al. (2007) reported that a 400-MPa treatment at 5 °C for 5 min was sufficient to inactivate 4.05 log10 PFU of MNV-1 within dissected oyster tissues. In our study, the 200MPa treatment resulted in partial inactivation, while the 400MPa treatment was sufficient to inactivate MNV-1 within oysters such that it is undetectable. To begin with, it should be noted that the rate at which the final pressure of our processor was attained was approximately 100 MPa/min; thus, the 400-MPa treatment actually lasted 9 min, which was much longer than that in the study conducted by Kingsley et al. (2007). In addition, we made a few modifications in the practical processing. Oysters are osmoconformers: their internal ionic strength mimics the ionic strength of the waters in which they reside. Since high salt concentrations can affect the rates of pathogen inactivation by HPP (Claci et al., 2005), we used artificial seawater whose salinity was maintained at 3.3% during the accumulation step. In order to make our experiment more relevant in terms of commercial application, we used whole oysters with intact shells rather than shucked ones. According to the study conducted by Kingsley et al. (2007), greater inactivation was observed for stock viruses at cooler temperatures. Before the HPP treatment, we packed the oysters with an ice–water mixture – a procedure feasible for commercial application. Ice was retained in the package at the end of the treatment, indicating that the actual processing temperature was approximately 0 °C. Lastly, Schwab, Neill, Estes, Metcalf, and Atmar (1998) studied the distribution of NoVs within shellfish following bioaccumulation and subsequent depuration, and found that NoVs are mostly distributed in dissected stomachs and digestive diverticula. Therefore, we used dissected digestive tissues instead of whole organisms during viral extraction in order to improve the concentration of the virus and reduce the amount of RT-PCR inhibitors within the shellfish tissues; and thus achieved a more satisfying result. As previously demonstrated by Kingsley et al. (2002), viral inactivation by HPP might only be due to subtle alterations in viral capsid proteins, which prevents the viruses from attaching to homologous cell receptors, but their RNA remains protected. In our study, the RT-PCR results continued to be positive for MNV1 even after treatment with HPP, indicating that the integrity of the RNA was intact – a fact that needs further investigation. It was reported that infectious viral particles, which can withstand
D. Li et al. / Food Control 20 (2009) 992–996
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Fig. 1. Detection of MNV-1 in oysters. RT-PCR products were electrophoresed on a 1% agarose gel. Marker: DL 2,000 (Takara Biotechnology Co., Ltd.); lane 1, negative control from non-contaminated oysters; lane 2, 0 MPa; lane 3, 0 MPa with proteinase K and RNase A; lane 4, 200 MPa; lane 5, 200 MPa with proteinase K and RNase A; lane 6, 300 MPa; lane 7, 300 MPa with proteinase K and RNase A; lane 8, 400 MPa; and lane 9, 400 MPa with proteinase K and RNase A.
the acidity and proteolytic enzymes in the stomach and small intestine, are considerably resistant to proteinase K (Nuanualsuwan & Cliver, 2003). In order to assess the damage to the viral capsid proteins by HPP, pretreatment with proteinase K and RNase A before RT-PCR was performed. The purpose of using proteinase K was to attack the protein capsid and thus release the genomic RNA, which could be broken down by RNase A. After pretreatment with proteinase K and RNase A, the RT-PCR results of HPP-inactivated MNV-1 tend to show fainter band density along with the increase in the processing pressure, whereas those of the similarly treated infectious viruses remained positive. This observation could be a convincing evidence for the fact that HPP inactivation primarily targets the stability of capsid proteins, thus making these proteins more susceptible to digestion by proteinase K. In order to solve the problem of false-positive RT-PCR results, three single-factor tests were performed on the basis of a study conducted by (Nuanualsuwan & Cliver, 2002) for devising an effective enzymatic pretreatment (data not shown). Both the concentration of enzymes and the incubation temperature had a significant effect on the enzymatic digestion. The pretreatment with 200 ng of RNase A and 40 U of proteinase K at 47 °C for 30 min was chosen for the premise that infectious viral particles should not be affected. According to our results, although the digestion with proteinase K and RNase A could enable the differentiation between infectious and HPP-inactivated MNV-1 by RT-PCR, a precise correlation between the results of RT-PCR and plaque assay could not be attained. In summary, this study demonstrates that MNV-1 within shellfish can be inactivated readily by high pressure. Our study indicated that if human and murine NoVs have similar susceptibilities to high pressure, HPP could be a viable processing intervention to increase the safety of shellfish intended for raw consumption and/or cooking. Even with a 4-log reduction in MNV-1 infectivity, oysters highly contaminated with human NoV might still contain infectious particles after HPP treatment. Therefore, we suggest that treating oysters with HPP must be combined with harvesting from safe areas. Our results also showed that RTPCR could not distinguish between infectious and non-infectious viral genomes of MNV-1, which indicated that the problem of false-positive results should be taken into account when interpreting RT-PCR-positive results for NoV with relation to the threat to human health. Acknowledgements The authors acknowledge the financial support received from the research project HI-TECH Research and Development Program of China (2007AA091802), ‘‘948” Program of Ministry of Agriculture (2006-G42), and Special Fund for Public Welfare Industry (Agriculture) (nyhyzx07-047).
We also thank Dr. H. W. Virgin IV from Washington University for providing the virus strain murine norovirus 1. References Baert, L., Wobus, C. E., Coillie, E. V., Thackray, L. B., Debevere, J., & Uyttendaele, M. (2008). Detection of murine norovirus 1 by using plaque assay, transfection assay, and real-time reverse transcription-PCR before and after heat exposure. Applied and Environmental Microbiology, 74, 543–546. Beuret, C., Baumgartner, A., & Schluep, J. (2003). Virus-contaminated oysters: A three-month monitoring of oysters imported to Switzerland. Applied and Environmental Microbiology, 69, 2292–2297. Cannon, J. L., Papafragkou, E., Park, G. W., Osborne, J., Jaykus, L.-A., & Vinje´, J. (2006). Surrogates for the study of norovirus stability and inactivation in the environment: A comparison of murine norovirus and feline calicivirus. Journal of Food Protection, 69, 2761–2765. Caul, E. O. (1996). Viral gastroenteritis: Small round structured viruses, caliciviruses and astroviruses. Part II. The epidemiological perspective. Journal of Clinical Pathology, 49, 959–964. Claci, K. R., Meade, G. K., Tezloff, R. C., & Kingsley, D. H. (2005). High-pressure inactivation of hepatitis A virus within oysters. Applied and Environmental Microbiology, 71, 339–343. Doultree, J. C., Druce, J. D., Birch, C. J., Bowden, D. S., & Marshall, J. A. (1999). Inactivation of feline calicivirus, a Norwalk virus surrogate. Journal of Hospital Infection, 41, 51–57. Duizer, E., Schwab, K. J., Neill, F. H., Atmar, R. L., Koopmans, M. P. G., & Estes, M. K. (2004). Laboratory efforts to cultivate noroviruses. Journal of General Virology, 85, 79–87. Grove, S. F., Lee, A., Lewis, T., Stewart, C. M., Chen, H., & Hoover, D. G. (2006). Inactivation of foodborne viruses of significance by high pressure and other processes. Journal of Food Protection, 69, 957–968. Le Guyader, F., Haugarreau, L., Miossec, L., Dubois, E., & Pommepuy, M. (2000). Three-year study to assess human enteric viruses in shellfish. Applied and Environmental Microbiology, 66, 3241–3248. Karst, S. M., Wobus, C. E., Lay, M., Davidson, J., & Virgin, H. W. IV, (2003). STAT 1dependent innate immunity to a Norwalk-like virus. Science, 299, 1575–1578. Kingsley, D. H., Holliman, D. R., Claci, K. R., Chen, H., & Flick, G. J. (2007). Inactivation of a norovirus by high-pressure processing. Applied and Environmental Microbiology, 73, 581–585. Kingsley, D. H., Hoover, D., Papafragkou, E., & Richards, G. P. (2002). Inactivation of hepatitis A virus and a calicivirus by high hydrostatic pressure. Journal of Food Protection, 65, 1605–1609. Murchie, L. W., Cruz-Romero, M., Kerry, J. P., Linton, M., Patterson, M. F., Smiddy, M., et al. (2005). High pressure processing of shellfish: A review of microbiological and other quality aspects. Innovative Food Science and Emerging Technologies, 6, 257–270. Nishida, T., Kimura, H., Saitoh, M., Shinohara, M., Kato, M., Fukuda, S., et al. (2003). Detection, quantitation, and phylogenetic analysis of noroviruses in Japanese oysters. Applied and Environmental Microbiology, 69, 5782–5786. Noda, M., Fukuda, S., & Nishio, O. (2008). Statistical analysis of attack rate in norovirus foodborne outbreaks. International Journal of Food Microbiology, 122, 216–220. Nuanualsuwan, S., & Cliver, D. O. (2002). Pretreatment to avoid positive RT-PCR results with inactivated viruses. Journal of Virological Methods, 104, 217–225. Nuanualsuwan, S., & Cliver, D. O. (2003). Capsid functions of inactivated human picornaviruses and feline calicivirus. Applied and Environmental Microbiology, 69, 350–357. Richards, G. P. (1999). Limitations of molecular biological techniques for assessing the virological safety of foods. Journal of Food Protection, 62, 691–697. Schwab, K. J., Neill, F. H., Estes, M. K., Metcalf, T. G., & Atmar, R. L. (1998). Distribution of Norwalk virus within shellfish following bioaccumulation and subsequent depuration by detection using RT-PCR. Journal of Food Protection, 61, 1674–1680.
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Straub, T. M., Bentrup, K. H., Orosz-Coghlan, P., Dohnalkova, A., Mayer, B. K., Bartholomew, R. A., et al. (2007). In vitro cell culture infectivity assay for human noroviruses. Emerging Infectious Diseases, 13, 396–403. Thurston-Enriquez, J. A., Haas, C. N., Jacangelo, J., Riley, K., & Gerba, C. P. (2003). Inactivation of feline calicivirus and adenovirus type 40 by UV radiation. Applied and Environmental Microbiology, 69, 577–582.
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Contents lists available at ScienceDirect
Food Control journal homepage: www.elsevier.com/locate/foodcont
Effects of high-pressure processing on murine norovirus-1 in oysters (Crassostrea gigas) in situ Dan Li1, Qingjuan Tang1, Jingfeng Wang, Yuming Wang, Qin Zhao, Changhu Xue * College of Food Science and Engineering, Ocean University of China, No. 5, Yushan Road, Qingdao, Shandong Province 266003, PR China
a r t i c l e
i n f o
Article history: Received 3 July 2008 Received in revised form 20 November 2008 Accepted 25 November 2008
Keywords: High-pressure processing Murine norovirus-1 Oyster RT-PCR
a b s t r a c t We assessed the effects of high-pressure processing (HPP) at 200–400 MPa on murine norovirus-1 (MNV1)-contaminated oysters by using a flow-through seawater system. Plaque assays demonstrated that a 5-min 400-MPa treatment at 0 °C inactivated MNV-1 within oysters to undetectable levels. No correlation was found between MNV-1 RNA detection by reverse transcription-PCR (RT-PCR) and the infectivity determined by the plaque assay before and after pressure exposure. Pretreatment with proteinase K and RNase A enabled the differentiation between infectious and HPP-inactivated MNV-1 by RT-PCR, indicating that HPP might subtly alter the viral capsid proteins but that the RNA remains protected. Ó 2008 Published by Elsevier Ltd.
1. Introduction Marine organisms consumed as seafood, particularly bivalve shellfish such as oysters, can bioaccumulate microbial pathogens at high levels from surrounding marine and estuarine waters (Murchie et al., 2005). Thorough cooking is an absolutely safe way of eliminating pathogens; however, this technique also alters the organoleptic qualities of shellfish. Hence, the consumption of raw or minimally cooked oysters is a common custom practiced worldwide. An alternate technology, high-pressure processing (HPP), has recently been developed as a potential means for reducing pathogens within raw shellfish. With the use of this technology, the appearance, flavor, texture, and nutritional qualities of unprocessed foods like oysters can be retained. In addition to enhancing the safety and extending the shelf-life of seafood, HPP treatment has the additional advantage of shucking shellfish, thus rendering this technology particularly useful to the shellfish-processing industry and consumers alike. HPP has become a commercial success in the American and European markets with the availability of several ready-to-eat meat products and seafoods, as reviewed in Grove et al. (2006) and Murchie et al. (2005). As the single most common cause of outbreaks as well as sporadic cases of acute gastroenteritis, noroviruses (NoVs) are a great threat to the safety of edible shellfish worldwide (Beuret, Baumgartner, & Schluep, 2003; Guyader, Haugarreau, Miossec, Dubois, & Pommepuy, 2000; Nishida et al., 2003; Noda, Fukuda, & Nishio, * Corresponding author. Tel.: +86 532 82032468; fax: +86 532 82032468. E-mail address: [email protected] (C. Xue). 1 These authors contributed equally to this work. 0956-7135/$ - see front matter Ó 2008 Published by Elsevier Ltd. doi:10.1016/j.foodcont.2008.11.012
2008). NoVs were first detected by immune electron microscopy during the 1968 outbreak of gastroenteritis in Norwalk, Ohio, US; the lack of suitable animal models and the inability to propagate NoVs in cell cultures have hampered further study on NoVs (Duizer et al., 2004). Straub et al. (2007) have described a highly differentiated 3D cell culture model that supports the natural growth of human NoVs. However, this system is rather complex; therefore, surrogate viruses that share common pathological and molecular features with human NoVs are employed instead. Feline calicivirus (FCV) was almost exclusively used as a surrogate virus in previous studies (Doultree, Druce, Birch, Bowden, & Marshall, 1999; Thurston-Enriquez, Haas, Jacangelo, Riley, & Gerba, 2003). Recently, a NoV infecting mice, namely, murine norovirus-1 (MNV-1), has been identified (Karst, Wobus, Lay, Davidson, & Virgin, 2003; Wobus, Thackray, &Virgin, 2006; Wobus et al., 2004). A report investigating the applicability of MNV-1 and FCV as surrogates for human NoV in studies on the stability and inactivation of NoV demonstrated that MNV-1 was more acid-tolerant than FCV, thus making it a more suitable surrogate for human NoV (Cannon et al., 2006). Currently, RT-PCR is the most commonly used detection method because of its rapidness and high sensitivity. However, since it cannot discriminate between inactivated and infectious viral particles, it may provide potentially false-positive results for foods that have actually been properly treated (Richards, 1999). The inclusion of a pretreatment with enzymes prior to RNA extraction has been investigated to solve this problem and has proved effective in the case of human hepatitis A virus, vaccine poliovirus 1, and feline calicivirus inactivated by ultraviolet light, hypochlorite, or heating (Nuanualsuwan & Cliver, 2002). Baert et al. (2008) examined MNV1 following heat exposure but observed no correlation between the
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number of infectious particles and viral genomes after heat treatment, regardless of the presence or nature of the enzyme treatment. However, whether MNV-1 inactivated by HPP can be detected by this method has not been reported yet. In this study, we aimed to examine the efficacy of HPP in inactivating MNV-1 within shellfish without altering the organoleptic qualities of oysters, and investigated the correlation between the number of infectious particles and viral genomes after HPP along with enzyme treatment. Oysters were bioaccumulated with MNV-1 in a flow-through seawater system before the inactivation by HPP; MNV-1 was detected by both plaque assay and RT-PCR. Pretreatment with proteinase K and RNase A was also performed. 2. Materials and methods 2.1. Virus and shellfish Working stocks of MNV-1 were prepared using confluent monolayers of RAW 264.7 mouse monocytes/macrophages (Shanghai Cell Bank, Shanghai, China) cultured in high-glucose Dulbecco’s modified Eagle medium (DMEM; Gibco-Invitrogen Co., Grand Island, NY, USA) supplemented with 10% fetal bovine serum (FBS; Hyclone Co., Logan, Utah, USA), 100 U/ml of penicillin (North China Pharmaceutical Group Corporation, Shijiazhuang, China), and 100 lg/ml of streptomycin sulfate (Amresco, Solon, OH, USA). After lysis, the lysate was freeze-thawed twice, centrifuged at 3500g for 10 min, and filtered through a membrane with a pore size of 0.22 lm. The MNV-1 stocks were stored at 80 °C. In order to determine the titer of the MNV-1 stock, plaque assays were performed in triplicate as described by Kingsley, Holliman, Claci, Chen, and Flick (2007) by using serially 10-fold diluted viral stocks in DMEM and assayed using confluent monolayers of RAW 264.7 mouse monocytes/macrophages in 6-well plates (Corning Inc., Corning NY, USA). Inoculation with 0.5 ml of MNV-1 was carried out for 2 h at 37 °C; this was followed by overlay with 2 ml of DMEM containing 1% low-melting-point agarose (BBI Co., Ltd., Boston, MA, USA) with 2% FBS. After 48 h of incubation, plaques were visualized by staining with 0.03% neutral red (Sigma Chemical Co., St. Louis, MO, USA) for 30 min at room temperature. The titer of the MNV-1 stock that was used to contaminate the oysters was approximately 2 1011 plaque forming units (PFUs)/ml. Pacific oysters (Crassostrea gigas) were harvested from an approved aquafarm in Jiaozhou Bay, Shandong Province, PR China. Considering the fact that the shellfish matrix composition, i.e., its fat content, water content, and salt concentration etc., can affect the pathogen inactivation rates of HPP and inhibit detection by RT-PCR (Claci, Meade, Tezloff, & Kingsley, 2005), oysters were bioaccumulated with MNV-1 in a flow-through seawater system as according to a method described by Kingsley, Hoover, Papafragkou, and Richards (2002) with a few modifications. Eight hours prior to viral accumulation, 24 commercial-size oysters were placed in an accumulation tank containing artificial seawater. The salinity of the seawater was 3.3%. An aquarium air pump was used to supply sufficient amounts of oxygen for the shellfish. MNV-1 (2 1011 PFU) was added to 4 l of double-distilled Milli-Q water (Millipore Corp., Bedford, MA, USA), which was continually mixed at 4 °C. Peristaltic pumps were adjusted to combine 4 ml of the viral suspension with 200 ml of UV-treated artificial seawater per minute in a cytostir vessel. After 16 h, the oysters were divided into 4 groups of 6 oysters each. 2.2. High-pressure treatment The oyster samples (6 oysters per treatment group) were transferred into aluminum foil pouches (30 cm 30 cm) containing an
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ice–water mixture and sealed using a DZQ-600 vacuum packaging machine (Zhangjiagang Deshun Machinery Co., China). An overpack pouch was sealed over the inner pouch. Pressurization of the oyster samples was carried out using a high-pressure food processor (Ningbo Branch, Institute of China Enginery, Ningbo, China). The range of the HPP treatment condition was chosen on the basis of the conditions generally maintained in commercial processing (Murchie et al., 2005). All samples were pressurized at 200, 300, and 400 MPa for 5 min using di-n-octyl sebacate (DOS) medium. The pressure vessel had a capacity of 5 l and an internal diameter of 120 mm. The rate at which the final pressure was attained was approximately 100 MPa/min, and pressure release was almost instantaneous. Ice bags were used during the shipment to ensure that the samples were maintained at a low temperature. Negative (uncontaminated) and positive (contaminated and nonpressurized) controls were processed in a manner similar to the HPP-treated oysters, except that they did not undergo the HPP treatment. 2.3. Viral extraction and plaque assays The stomachs and digestive diverticula of the oysters (61 g from each of the 6 oysters per treatment group) were removed by dissection, placed in 50-ml conical tubes that contained 6 ml of glycine buffer (0.1 M glycine, 0.3 M NaCl; pH 9.5, 4 °C), and homogenized by a high-performance disperser (IKA T18 basic ULTRA-TURRAXÒ, German) at 18,000 rpm for approximately 3 min. An ice bath was used to maintain the dispersion temperature below 10 °C. The homogenized extracts were centrifuged at 12,000g for 15 min at 4 °C in a Jouan MR23i centrifuge (Thermo Scientific). The resultant supernatant was neutralized with 2 N HCl and mixed with penicillin (1000 U/ml) and streptomycin sulfate (1000 lg/ ml); this was followed by incubation at 4 °C for 1 h. Tenfold serial dilutions were made in DMEM, and plaque assays were performed in triplicate using the same method as that used for determining the titer of the MNV-1 stock. 2.4. Residual infectivity No interference was observed in the cell culture and MNV-1 plaque assay until the viral extraction was diluted at least 100 times in DMEM. The detection limit of the MNV-1 assay was consequently raised to 200 PFU MNV-1/oyster. Thus, a residual infectivity test was carried out to verify whether all viral particles in the suspension were inactivated. The viral extraction (0.1 ml) was inoculated into confluent monolayers of RAW 264.7 mouse monocytes/macrophages in 100-mm tissue culture dishes and incubated at 37 °C for 2 h. After 48 h, the viral extraction was observed in order to determine whether it could induce a cytopathic effect in the mouse monocytes/macrophages. After 48 h, the lysate was freeze-thawed twice and diluted tenfold. Plaque assays were performed using the same method as that stated before. 2.5. Composite enzymatic pretreatment RNase A (BBI Co., Ltd., Boston, MA, USA) was diluted in TrisEDTA buffer (1.0 M Tris–HCl, 0.1 M EDTA; pH 8.0) and maintained at 20 °C. Proteinase K (BBI Co., Ltd., Boston, MA, USA) was dissolved in 0.01 M PBS and prepared freshly for each experiment. Both 40 U of proteinase K and 200 ng of RNase A were added to 100 ll of the viral extraction and incubated at 47 °C for 30 min. To 10 ll of RNA isolated from MNV-1, 10 ng of RNase A was added, and this solution served as a positive control for RNase A activity. In order to test the enzyme activity of proteinase K, pretreatments were performed with both RNase A and RNase A combined with proteinase K.
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2.6. RNA extraction and RT-PCR Regardless of enzyme pretreatment, 100 ll of viral extraction and 500 ll of TRIzol Reagent (Invitrogen, Carlsbad, CA, USA) were vigorously mixed with each sample by repipetting. After incubation for 5 min at room temperature, 100 ll of chloroform was added to each sample. The samples were then vortexed vigorously for 30 s, incubated at room temperature for 5 min, and centrifuged at 12,000g for 5 min. The upper aqueous layer, which contained the RNA, was precipitated by the addition of 0.5 volume of isopropanol for 5 min at room temperature; this was followed by centrifugation at 12,000g for 5 min. The resulting white pellets were washed with cold 75% ethanol, and each pellet was then resuspended in 50 ll of RNase-free water. RT-PCR amplification of the MNV-1 sequences was performed in a two-step format. The primer 3311 50 -GATTGGTAGT GCCGTTGTG-30 and the antisense primer 3732 50 -TGACCTGCTGAAGGGAAG-30 were used. Reverse transcription of all viral RNA was performed at 37 °C for 60 min using MMLV reverse transcriptase (Promega, Madison, WI, USA). We performed 40-cycle PCR reactions that involved annealing at 55 °C for 1 min, extension at 72 °C for 1 min, and denaturation at 95 °C for 30 s. For the final cycle, the annealing time was extended to 2 min, and the extension was performed for 10 min. The final concentration of each of the primers used was approximately 0.25 lM. The PCR products were separated by 1% agarose gel electrophoresis, stained with ethidium bromide, and visualized by UV light. 2.7. Analysis of data The MNV-1 titers were determined in triplicate. The average PFU obtained was calculated, and the standard error was expressed logarithmically. 3. Results At the end of the bioaccumulation period, the MNV-1 titers in the oysters in three separate trials were determined to be 1.56 104, 1.44 104, and 1.12 104 PFU per group. Since it is commonly believed that less than 100 norovirus particles are sufficient to cause illness (Caul, 1996), our results confirmed that shellfish have the ability of bioaccumulating viruses to a level far beyond the infectious dose. HHP treatment, with pressures ranging from 200 to 400 MPa and applied in 100-MPa increments, was administered to three treatment groups for 5 min at approximately 0 °C. The average titer obtained at each pressure is displayed in Table 1. The onset of inactivation was observed at 200 MPa, and treatment with 400 MPa was sufficient to inactivate MNV-1 within the oysters such that the viral load is undetectable. To confirm whether the viral load is undetectable only because of the inactivation of MNV-1, a residual infectivity test was performed. The results confirmed that
Table 1 Residual infectivity of MNV-1 detected after HHP treatment. Pressure (MPa)
Average log10 PFU (SEa)
Log10 PFU (SE a) for trial no. 1
2
3
0 200 300 400
4.13 (0.06) 3.56 (0.05) 3.18 (0.08) ND
4.19 (0.05) 3.57 (0.16) 3.18 (0.14) ND
4.15 (0.10) 3.50 (0.40) 3.08 (0) ND
4.04 (0.08) 3.62 (0.23) 3.28 (0.14) ND
Rate of reduction 0 13.8% 23.0% 100%b
ND: not detected. a SE: standard error. b Since the detection limit of the plaque assay is 200 PFU MNV-1/oyster, this result was verified by a residual infectivity test.
MNV-1 was indeed inactivated by the 400-MPa treatment. Moreover, the HPP treatment used in this study (200–400 MPa) did not affect the visual appearance of oysters and also facilitated their shucking. Viral RNA extraction was performed on all shellfish extracts. All controls (non-seeded oysters) tested negative for MNV-1 by RTPCR, indicating that all oysters were originally free of MNV-1. The 439-bp major amplification product was analyzed for MNV-contaminated samples in all oyster groups (Fig. 1; lanes 2, 4, 6, and 8). No genomic copies were detected after the addition of 10 ng of RNase A to 10 ll of RNA isolated from MNV-1, thus proving that RNase A was able to degrade free viral RNA. Pretreatment with both RNase A and proteinase K yielded mostly negative RT-PCR results for the 400-MPa inactivated MNV-1; however, positive results were obtained when only RNase A was used. This indicated that proteinase K effectively attacked the capsid (data not shown). After pretreatment with 200 ng of RNase A and 40 U of proteinase K at 47 °C for 30 min, the RT-PCR results of HPP-inactivated MNV-1 tend to present fainter band density along with the increase in the processing pressure (Fig. 1; lanes 5, 7, and 9), whereas those of the similarly treated infectious viruses remained positive (Fig. 1, lane 3).
4. Discussion Kingsley et al. (2007) reported that a 400-MPa treatment at 5 °C for 5 min was sufficient to inactivate 4.05 log10 PFU of MNV-1 within dissected oyster tissues. In our study, the 200MPa treatment resulted in partial inactivation, while the 400MPa treatment was sufficient to inactivate MNV-1 within oysters such that it is undetectable. To begin with, it should be noted that the rate at which the final pressure of our processor was attained was approximately 100 MPa/min; thus, the 400-MPa treatment actually lasted 9 min, which was much longer than that in the study conducted by Kingsley et al. (2007). In addition, we made a few modifications in the practical processing. Oysters are osmoconformers: their internal ionic strength mimics the ionic strength of the waters in which they reside. Since high salt concentrations can affect the rates of pathogen inactivation by HPP (Claci et al., 2005), we used artificial seawater whose salinity was maintained at 3.3% during the accumulation step. In order to make our experiment more relevant in terms of commercial application, we used whole oysters with intact shells rather than shucked ones. According to the study conducted by Kingsley et al. (2007), greater inactivation was observed for stock viruses at cooler temperatures. Before the HPP treatment, we packed the oysters with an ice–water mixture – a procedure feasible for commercial application. Ice was retained in the package at the end of the treatment, indicating that the actual processing temperature was approximately 0 °C. Lastly, Schwab, Neill, Estes, Metcalf, and Atmar (1998) studied the distribution of NoVs within shellfish following bioaccumulation and subsequent depuration, and found that NoVs are mostly distributed in dissected stomachs and digestive diverticula. Therefore, we used dissected digestive tissues instead of whole organisms during viral extraction in order to improve the concentration of the virus and reduce the amount of RT-PCR inhibitors within the shellfish tissues; and thus achieved a more satisfying result. As previously demonstrated by Kingsley et al. (2002), viral inactivation by HPP might only be due to subtle alterations in viral capsid proteins, which prevents the viruses from attaching to homologous cell receptors, but their RNA remains protected. In our study, the RT-PCR results continued to be positive for MNV1 even after treatment with HPP, indicating that the integrity of the RNA was intact – a fact that needs further investigation. It was reported that infectious viral particles, which can withstand
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Fig. 1. Detection of MNV-1 in oysters. RT-PCR products were electrophoresed on a 1% agarose gel. Marker: DL 2,000 (Takara Biotechnology Co., Ltd.); lane 1, negative control from non-contaminated oysters; lane 2, 0 MPa; lane 3, 0 MPa with proteinase K and RNase A; lane 4, 200 MPa; lane 5, 200 MPa with proteinase K and RNase A; lane 6, 300 MPa; lane 7, 300 MPa with proteinase K and RNase A; lane 8, 400 MPa; and lane 9, 400 MPa with proteinase K and RNase A.
the acidity and proteolytic enzymes in the stomach and small intestine, are considerably resistant to proteinase K (Nuanualsuwan & Cliver, 2003). In order to assess the damage to the viral capsid proteins by HPP, pretreatment with proteinase K and RNase A before RT-PCR was performed. The purpose of using proteinase K was to attack the protein capsid and thus release the genomic RNA, which could be broken down by RNase A. After pretreatment with proteinase K and RNase A, the RT-PCR results of HPP-inactivated MNV-1 tend to show fainter band density along with the increase in the processing pressure, whereas those of the similarly treated infectious viruses remained positive. This observation could be a convincing evidence for the fact that HPP inactivation primarily targets the stability of capsid proteins, thus making these proteins more susceptible to digestion by proteinase K. In order to solve the problem of false-positive RT-PCR results, three single-factor tests were performed on the basis of a study conducted by (Nuanualsuwan & Cliver, 2002) for devising an effective enzymatic pretreatment (data not shown). Both the concentration of enzymes and the incubation temperature had a significant effect on the enzymatic digestion. The pretreatment with 200 ng of RNase A and 40 U of proteinase K at 47 °C for 30 min was chosen for the premise that infectious viral particles should not be affected. According to our results, although the digestion with proteinase K and RNase A could enable the differentiation between infectious and HPP-inactivated MNV-1 by RT-PCR, a precise correlation between the results of RT-PCR and plaque assay could not be attained. In summary, this study demonstrates that MNV-1 within shellfish can be inactivated readily by high pressure. Our study indicated that if human and murine NoVs have similar susceptibilities to high pressure, HPP could be a viable processing intervention to increase the safety of shellfish intended for raw consumption and/or cooking. Even with a 4-log reduction in MNV-1 infectivity, oysters highly contaminated with human NoV might still contain infectious particles after HPP treatment. Therefore, we suggest that treating oysters with HPP must be combined with harvesting from safe areas. Our results also showed that RTPCR could not distinguish between infectious and non-infectious viral genomes of MNV-1, which indicated that the problem of false-positive results should be taken into account when interpreting RT-PCR-positive results for NoV with relation to the threat to human health. Acknowledgements The authors acknowledge the financial support received from the research project HI-TECH Research and Development Program of China (2007AA091802), ‘‘948” Program of Ministry of Agriculture (2006-G42), and Special Fund for Public Welfare Industry (Agriculture) (nyhyzx07-047).
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