Identification of Effective and Practical Thermal and Non-thermal Processing
Technologies to Inactivate Major Foodborne Viruses in Oysters
DISSERTATION
Presented in Partial Fulfillment of the Requirements for the Degree Doctor of
Philosophy in the Graduate School of The Ohio State University
By
Elbashir Mohamed Araud
Graduate Program in Comparative Veterinary Medicine
The Ohio State University
2015
Ph.D. Examination Committee: Dr. Jianrong Li, Advisor Dr. Hua Wang Dr. Melvin Pascall Dr. Gireesh Rajashekara
Copyright by
Elbashir Mohamed Araud
2015
ABSTRACT
Human enteric viruses, such as human norovirus (HuNoV), hepatitis A virus
(HAV), and rotavirus (RV), are responsible for the majority of foodborne illnesses.
Seafood, particularly bivalve shellfish, is one of major high risk foods for enteric viruses contamination. Currently, the ecology, bioaccumulation, and persistence of enteric viruses in shellfish are poorly understood. There is no standard method to effectively inactivate these viruses in seafood. Therefore, there is an urgent need to develop effective thermal and non-thermal processing technologies to eliminate virus hazard in seafood.
The goals of this study are to: i) determine the natural bioaccumulation patterns of human enteric viruses in shellfish tissues, ii) to determine whether heat or high pressure processing (HPP) are capable of effectively inactivating enteric viruses in shellfish, iii) and to determine whether viruses can develop resistance to processing technologies.
Live oysters (Grassostrea gigas) were cultivated in a tank containing 4 liters of synthetic seawater and were artificially contaminated with HuNoV GII.4, HuNoV surrogates (Tulane virus, TV; murine norovirus, MNV-1), HAV, or RV at level of 1 ×
104 PFU/ml of TV, MNV-1, HAV, or 1 × 104 RNA copies/ml of HuNoV. At 24, 48, and
72 h post-inoculation, oysters were harvested, different portions of oyster tissue including gills, digestive glands, and muscles were isolated, and the presence of the viruses in each
ii tissue was determined by plaque assay or real time PCR (RT-qPCR). It was found that all viruses were bio-accumulated to a high titer within the oyster tissues within 72 h; however, the pattern of the bioaccumulation varied for each individual virus.
Caliciviruses (HuNoV, TV, and MNV-1) and HAV were localized in the stomach at a high level within the first 24 h, while RV was bio-accumulated to the highest level in gills after 24 h. In order to determine the thermal stability of the four cultivable viruses
(TV, MNV-1, HAV, and RV), each virus was diluted in cell culture medium, and the kinetics of viral inactivation was determined at temperatures of 62, 72, and 80 ˚C. It was found that the biphasic reduction model was the best fit to describe the virus inactivation curves at 62 and 72 ˚C. Decimal reduction time (D-values) of the low and high thermal resistant fractions of the four cultivatable viruses ranged from 0.13 to 1.81 min, and from
1.26 to 7.29 s at 62 and 72˚C, respectively. In contrast, the Weibull distribution was the best fit for the inactivation curves at 80 ˚C, and the time to first log10 reduction (TFL- value) ranged between 0.46 and 32 s. Within the oyster tissues, the TFL at 80 ˚C ranged between 0.61 to 19.99 min. The four viruses can be ranked from the most heat resistant to the least stable as the following: HAV>RV>TV>MNV-1. At 80 ˚C, time required for complete inactivation of HAV, RV, TV, and MNV-1 in cell culture medium is 12 s, 10 s,
10 s, and 6 s, respectively. However, it required 4, 3, and 3 min at 80 ˚C to completely inactivate RV, MNV-1, and TVin oysters, respectively, HAV survived the treatment even after 6 min. To decipher the mechanism underlying viral inactivation by heat, purified
TV was treated at 80°C for increasing time intervals. It was found that the integrity of the viral capsid was disrupted whereas viral genomic RNA remained intact. However, a iii lethal dose of heat treatment was not sufficient to disrupt the receptor binding activity of
HuNoV, HuNoV virus-like particles (VLPs), and TV. These data demonstrated that enteric viruses were efficiently bioaccumulated in oyster tissues and different viruses exhibited different distribution patterns. Although foodborne viruses have variable thermal stability to heat, 80˚C for 6 min was not sufficient to completely inactivate all the tested viruses in oysters.
High pressure processing (HPP) is a promising non-thermal technology that inactivates foodborne viruses while maintaining the organoleptic properties of processed foods. However, one interesting observation for HPP inactivation of viruses is that different viruses are variable in their susceptibility to high pressure. To date, the HPP sensitivity of different virus strains in the same genus, species or serotype is not known.
RVs are genetically diverse which makes it a good model to study the role of strain diversity in HPP inactivation of viruses. This study compared the baro-sensitivity of seven RV strains derived from four serotypes (G1: Wa, Ku, and K8, G2: S2, G3: SA-11 and YO, and G4:ST3) following high pressure treatment. It was found that RV strains showed varying responses to HPP based on the initial temperature and had different inactivation profiles. Ku, K8, S2, SA-11, YO, and ST3 showed enhanced inactivation at
4°C compared to 20°C. In contrast, Wa strain was not significantly impacted by the initial treatment temperature. Within serotype G1, Wa stain was significantly (p<0.05) more resistant to HPP compared to Ku and K8. Overall, the resistance of the human RV strains to HPP at 4°C can be ranked as Wa>Ku=K8>S2>YO>ST3 and in terms of
iv serotype G1>G2>G3>G4. In addition, pressure treatment of 400 MPa for 2 min was sufficient to eliminate the Wa strain, the most pressure resistant RV, from oyster tissues.
HPP disrupted virion structure, but did not degrade viral protein or RNA, providing insight into the mechanism of viral inactivation by HPP. Therefore, HPP is capable of inactivating RV at commercially acceptable pressures and the efficacy of inactivation is strain dependent.
In general, microorganisms are capable of acquiring resistance under stress.
However, whether enteric caliciviruses can develop resistance during thermal and nonthermal processing is not known. This study utilizes TV, an enteric primate calicivirus, as a surrogate for HuNoV, to screen for heat and high pressure (HPP) resistant strains. Wild type (WT) TV was subjected to either heat treatment at 70˚C for 6 s in culture medium for 5 passages or HPP at 300 MPa for 2 min at 20˚C for 3 passages.
Viral plaques were purified after each heat or HPP treatment and were subsequently re- treated or re-pressured and the change in log reduction after each treatment cycle was determined by plaque assay. At each passage, individual plaques were isolated, and tested for potential heat or pressure resistance. It was found that TV gradually developed resistance under heat and HPP stress. It appears that TV more rapidly developed resistance to HPP compared to development of resistance to heat. At passage 3, plaques isolated from HPP treated viruses had 2-3 logs more resistance compared to wild type
TV. At passage 5, plaques isolated following heat treatment had 1-3 logs more resistance compared to wild type TV. Interestingly, these heat or pressure resistant TVs had delayed
v growth replication kinetics, delayed cytopathic effects, and smaller plaque size, suggesting that they were attenuated in cell culture. Finally, the entire genome of two heat resistant and two pressure resistant TVs were amplified by RT-PCR and sequenced.
Interestingly, the majority of mutations were found in the major capsid protein (VP1) and the minor capsid protein (VP2) although some of mutations were located in RNA dependent RNA polymerase and the N-terminal domain of nonstructural polyprotein.
Collectively, we demonstrated that TV can easily develop resistance under both heat and
HPP, and mutations in the genome may be responsible for the resistance.
In summary, these data demonstrated that: (i) enteric viruses can be efficiently bio- accumulated in oyster tissues; (ii) different viruses have different bioaccumulation patterns in oyster tissues; (iii) enteric viruses have variable thermal stability and resistance to heat treatment can be ranked as HAV (most resistant)>RV>TV>MNV-
1(most sensitive); (iv) thermal treatment at 80˚C for 6, 4, 3, and 3 min was sufficient to completely inactivate HAV, RV, TV, and MNV-1 in oyster, respectively; (v) RV strains derived from different serotypes had different inactivation profiles and the resistance of the RV strains to HPP can be ranked as Wa>Ku=K8>S2>YO>ST3 and in terms of serotype G1>G2>G3>G4; (vi) TV can easily develop resistance to both heat and HPP treatments and majority of mutations responsible for resistance phenotype are located in major and minor capsid proteins; and (vii) heat and pressure resistant TVs are attenuated in cell culture. The findings in this study will facilitate the development of effective thermal and non-thermal processing technologies to eliminate enteric viruses in shellfish
vi and will facilitate the establishment of new standards for industry and regulatory agencies to improve seafood safety.
vii
Acknowledgments
It is my pleasure to thank all the people who made this work possible. I would like to thank my advisor Dr. Jianrong Li for his support and guidance in the execution of my research and pursuit of a career in science. I appreciate him giving me the opportunity to advance my education and for fostering my ability to think critically.
Besides my advisor, I would like to thank the rest of my dissertation committee: I would like to thank Dr. Hua Wang, Dr. Melvin Pascall, and Dr. Gireesh Rajashekara for acting as my committee members, their time and commitment is greatly appreciated. I also extend my gratitude to the members of Dr. Li’s research group who have been gracious in sharing their knowledge and expertise, as well as their kindness and humor, during my time in graduate school. All current and former lab members including Dr. Erin
DiCaprio, Dr. Yuanmei Ma, Dr. Yu Zhang, Dr. Hui Cai, Dr. Fangfei Lou, Yue Duan, Dr.
Junan Li, Ben Yeap, Dr. Yongwei Wei, Rongzhang Wang, Xueya Liang, and Dr. Mijia
Lu have all been instrumental in my success. In particular, I am very grateful to Erin
DiCaprio for her critical review of my dissertation and manuscripts.
I would like to thank my wife Rania for her help and support during all my study.
I would also like to thank my Kids Yasmin, Moftah, and Adam for their patient during viii those years of school. Finally, I would like to thank my big family, my parents, my brothers, and my sisters for their lifelong love, for always believing in me, and being more confident in my abilities than I ever have been.
ix
VITA
1987-1991………………………………….B.S. Food Science and technology
Higher Institute of Technology, Brack,
Libya.
1994-1999……………………………….… Microbiology Technician in the Biology
Department, College of Science, University
of El- Gabal El- Gahrby, Libya
2000-2004………………………………… MSc. in Agriculture Microbiology, Cairo
University, Cairo, Egypt.
2004-2009…………………………………... Assistant lecturer, Biology Department, the
University of El- Gabal, El- Gahrby, Libya
2010-2013………………………………....Graduate Research Associate, Department
of Food Science and Technology, The Ohio
State University, Columbus, OH
x 2013-2015………………………………....Graduate Research Associate, Department
of Veterinary Biosciences, The Ohio
State University, Columbus, OH
PUBLICATIONS
Elbashir Araud, Erin DiCaprio, Zhihong Yang, Xinhui Li, Fangfei Lou, John H. Hughes,
Haiqiang Chen, and Jianrong Li. High-pressure inactivation of rotaviruses: the role of treatment temperature and strain diversity in virus inactivation. Applied Environmental
Microbiology, AEM.01853-15.
Barakat Olfat, S., El-Gizawy, S A., Araud, EM, & Zahra, MK. (2005). Biological conversion of barley straw into glucose for baker’s yeast biomass production. Mansoura
Journal of Agricultural Science. 30(12) 8053-8073.
Fields of Study
Major Field: Comparative Veterinary Medicine
xi
Table of Contents
ABSTRACT……………………………………………………………………..………..ii
ACKNOWLEDGEMENTS………………………………………………………..……vii
VITA……………………………………………………………………………..………ix
LIST OF TABLES..………………………………………………………………...…..xxv
LIST OF FIGURES..………………………………………………………….………xxvii
ABBREVIATIONS………………………………………………………………...…..xxx
1. LITERATURE REVIEW……………………………………………………..……….1
1.1. Introduction………………………………………………………………...... 1
1.2. Foodborne viruses. ..………………………………………………………….5
1.3. Human norovirus (HuNoV)……………………………………...…………..7
1.3.1. Molecular biology of HuNoV …..………..……………………...……...8
1.3.2. HuNoV infection………………….……………………………...….…10
xii 1.3.3. HuNoV stability ..……………………………….……………………..11
1.3.4. Cell culture systems for HuNoV………...………….………………….12
1.3.5. HuNoV surrogates … ………………………….…………………….14
1.3.5.1. HuNoV virus-like particles (VLPs) . … ………….……..….………15
1.3.5.2. Tulane Virus (TV)…………………………….……………………..16
1.3.5.3. Comparing MNV-1 and TV as human norovirus surrogates ………19
1.4. Rotavirus (RV)………………………..…………………………………….20
1.4.1. Rotavirus molecular biology ….……………..….…………………….22
1.5. Hepatitis A virus (HAV)…………………………..………….……………..24
1.6. Limitations in current food safety controls to eliminate viruses from high risk foods……………………………………………………………………………..……….25
1.7. Detection methods for foodborne viruses………………….………………..26
1.7.1. HuNoV receptor binding assays for viral detection in foods…………..28
1.8. Shellfish are a high risk food for virus contamination………………………33
1.8.1. Seafood production practices …. ……….…..………………………..35
1.8.2. Seafood harvesting………………….……………..…………………..36
xiii 1.8.3. Post-harvest seafood processing ……….……..……………...……….38
1.8.4. Bioaccumulation and distribution of viruses in bivalve mollusk
tissues………………………………………………………………………38
1.9. Control strategies for foodborne viruses in shellfish...... 40
1.10. Depuration and relying …….. …………..………..………………….……40
1.11. Thermal processing . ………………………………….…………..……….44
1.11.1. The inactivation models of foodborne viruses……………….……….45
1.11.2. The comparison of the thermal stability of several foodborne
viruses………………..…………………………………………………….46
1.12. High hydrostatic pressure (HPP) ..………….……………………………..50
1.12.1. Mechanism of HPP inactivation of viruses ………………………….52
1.12.2. Intrinsic factors that influence virus inactivation by HPP ……..…… 53
1.12.3. Application of HPP in processing high-risk foods ……………….….56
2. Natural bioaccumulation of enteric foodborne viruses in live oysters and thermal inactivation of enteric viruses
2.1. Abstract …..……………………………………………………….………..58
2.2. Introduction………………………………………………………………….59
xiv 2.3. Materials and Methods……………….……………………………..……….63
2.3.1. Cell culture and virus propagation ..……………….………………….63
2.3.2. Viral plaque assays . …………………………………………………..64
2.3.3. Virus bio-accumulation and distribution in oysters …..……...………..64
2.3.4. Plaque assay and virus extraction from contaminated oyster tissues …65
2.3.5. Thermal treatment of viruses in culture medium………………………65
2.3.6. Heat inactivation of MNV-1, TV, HAV, and RV in oyster tissues …...66
2.3.7. Isolation and characterization of HuNoV GII.4 strain 5M . ..………..67
2.3.8. Production of HuNoV GII.4 virus-like particles (VLPs) ….….……….67
2.3.9. Preparation of porcine gastric mucin conjugated to magnetic beads
(PGM-MBs) ………………………………………………………..68
2.3.10. PGM-MB binding assay ….…. ……………………………………….69
2.3.11. Quantification of viral RNA by real-time RT-PCR ….…………..…….69
2.3.12. Thermal treatment of HuNoV GII.4 VLPs ….………………………….70
2.3.13. Purification of TV …….………………….……… …………………….71
2.3.14. Transmission electron microscopy………………. …….……………….71
xv 2.3.15. Reverse transcriptase polymerase chain reaction (RT-PCR) ..…..….71
2.3.16. Statistical analysis ……………………………………………………72
2.4. Results ..………………………………………………………………...... 72
2.4.1. Virus bio-accumulation and distribution in oysters.…………..……….72
2.4.2. Thermal inactivation of virus in cell culture medium………………….77
2.4.3. Thermal inactivation of viruses within oyster tissues …..…….……….83
2.4.4. The effect of the thermal treatment on HuNoV GII.4 VLPs ...……….84
2.4.5. Effect of heat treatment on HuNoV GII.4 and TV … ..………………87
2.4.6. Effect of heat treatment on TV RNA level. …………..……………….90
2.4.7. The effect of thermal treatment on purified TV ….…..….……………92
2.5. Discussion………………………………………………………………..….92
2.5.1. Virus bio-accumulation and distribution in oysters……..……………..94
2.5.2. Thermal inactivation of viruses ………….……………………………96
2.5.3. Insight into the mechanism underlying thermal inactivation of viruses………………………………………………….………………………………..99
3. High-pressure Inactivation of Rotaviruses: The Role of Treatment Temperature and Strain Diversity in Virus Inactivation
xvi 3.1. Abstract ...…………………………….…………………………………...102
3.2. Introduction…………………………………………………………..…….103
3.3. Materials and Methods …...……………………………………………106
3.3.1. Viruses and cell culture ….………..…………………..……….…….106
3.3.2. RV plaque assay …….……….…….…………………..….………....107
3.3.3. Pressure inactivation of different RV trains .….………….….……….107
3.3.4. Purification of RV . .... …………….………….…………………...... 108
3.3.5. Reverse transcriptase polymerase chain reaction (RT-PCR) ….…..…108
3.3.6. Transmission electron microscopy ……….…………….……………109
3.3.7. Analysis of RV proteins by SDS-PAGE ….……………………..….109
3.3.8. Bioaccumulation of RV in oyster tissues ..…………………………. 110
3.3.9. Statistical analysis ………………..………………………………..110
3.4. Results .……………………………………………………..…………...... 110
3.4.1. The effect of HPP initial temperature on different RV strains ………110
3.4.2. Comparing the baro-sensitivity of different human RV strains . ……117
3.4.3. The effect of HPP on RV capsid . ………..………………………..118
xvii 3.4.4. HPP effect on the viral proteins ……………………………………120
3.4.5. The effect of HPP on viral RNA… .………………………………… 122
3.4.6. Inactivation of RV in oyster tissues ………..………..……………..124
3.5. Dissussion ..…………………………………………………………….125
3.5.1. Role of initial temperature on the inactivation of seven RV
strains ……………………………………………….……………..…..126
3.5.2. Role of strain diversity on HPP inactivation ….……………………..128
3.5.3. Mechanism underlying HPP inactivation of viruses .……………….131
3.5.4. Inactivation of RV (Wa strain) in oyster tissues …………………….133
4. Resistance of Tulane virus, an enteric primate calicivirus, to thermal and high pressure processing .………………………………………………………………………....136
4.1. Abstract ………………………………………………………………….136
4.2. Introduction…………………….…………………………………………..137
4.3. Materials and Methods……………………………………………………..140
4.3.1. Cell culture and virus propagation ...……………………..……..…140
4.3.2. TV plaque assays …………………..……………………………..141
xviii 4.3.3. Screening for heat resistant isolates ….……..………..…………….141
4.3.4. Screen for HPP resistant isolates .……..…….……………………...142
4.3.5. TV plaque purification .…………………..…………..……………143
4.3.6. Plaque size determination .….………………..………………….....143
4.3.7. Single-cycle growth curves of heat and HPP isolates ……..…….….143
4.3.8. Determination of the mutations in the heat and HPP treated isolates.. 144
4.3.9. Sequence Alignment ….…………………………….………………..145
4.3.10. Statistical analysis ………………...………………..……………...146
4.4. Results .………………………………………………………………...146
4.4.1. Screening for heat resistant isolates ……………………………....146
4.4.2. Screen for HPP resistant isolates ……………………..…………..149
4.4.3. Single-cycle growth curves of heat and HPP isolates ……………....151
4.4.4. Plaque size determination …………………………………….……154
4.4.5. Localization of mutations in heat and pressure resistant TVs ……….156
4.5. Discussion …………………………………………………………….157
4.5.1. Heat resistance …………………….………………..………………158
xix 4.5.2. HPP resistance …..….……………………….…………….………159
4.5.3. Phenotype of heat and pressure-resistant TV mutants …………….160
4.5.4. Genetic changes of the heat and HPP stable strains …..…………….161
5. CONCLUSION…...…………………………………………………………...... 164
5.1. Conclusion…………………………………………………………………164
5.2. Future Directions…………………………………………………………..166
LIST OF REFERENCES……………………………………………………………….171
xx
LIST OF TABLES
Table 1. Summary of the major foodborne viruses…………….………………………6
Table 2. HuNoV proteins and their functions……………………………………….….9
Table 3. Comparison of MNV-1 and TV to HuNoV…….…………………….………19
Table 4. RV structure and non-structure proteins……….…………..…………………22
Table 5. D-values and TFL for HAV, RV, TV, MNV-1 after heat treatment at 62˚C in
culture medium………………………………………………………………...73
Table 6. Comparing Log liner, Weibull distribution, biphasic models to best fit to the
survival curve of TV, MNV-1, HAV, and RV at 62, 72, and 80˚C in culture
medium………………………………………………………………………….75
Table 7. D-values and TFL-values of HAV, RV, TV, and MNV-1 at 72˚C in culture
medium ……….. ……………………………………….……………………….76
Table 8. D-values and TFL-values of HAV, RV, TV, and MNV-1 at 80˚C in culture
medium …………...……………………………………………………………..76
Table 9. TFL-values of HAV, RV, TV, and MNV-1 at 80˚C in the oyster tissues…..….78
Table 10: List of primers used for TV sequencing………..……………………………135
xxi Table 11: Localization of mutations in genome of heat and pressure resistant TVs…...147
xxii
LIST OF FIGURES
Figure1. Top pathogens leading to illness in the US ……………………….……………2
Figure 2. Norovirus genogroup and genotype characterization based on sequence
homology of the VP1 ………………………………………………..……….…8
Figure 3. Transmission electron microscopy image of human NoV virus-like particles
(VLPs) ………………………………………………………………..……….15
Figure 4. Transmission electron microscopy image of TV …………….…………. ….16
Figure 5. Transmission electron microscopy image of MNV-1 ..………..…..….……18
Figure 6. RV genome, proteins, and structure …… ……………………..…………….21
Figure 7. Bioaccumulation of Caliciviruses in oyster tissues …………………..………67
Figure 8. Bioaccumulation of HAV and RV in oyster tissues…………………………..70
Figure 9. Effect of heat treatment at 62, 72, and 80˚C on MNV-1, TV, HAV, and RV (Wa
strain) in culture medium …………….....…………………………………….71
Figure 10. The effect of heat at 80, and 100˚C on the binding ability of the VLPs of
HuNoV GII.4 to bind to the beads …………………….………………… …..80
xxiii Figure 11. TV heat inactivation at 62, 72, and 80˚C by using binding assay and RT-
qPCR………………………………………………………………....………82
Figure 12. HuNoV heat inactivation at 62, and 72˚C by using binding assay and RT-
qPCR……………….………………………………………………..………84
Figure 13.The effect of heat treatment at 80˚C on the capsid gene (VP1) of
TV……………………………….…………………………………………..85
Figure 14. - Visualize the damaged TV particles after heat treatment at 80˚C for 5s, 10s,
and 5 min..………………………………………………………………...86
Figure 15. Effect of temperature on inactivation of RV serotype G1 strains….……….104
Figure 16. Effect of temperature on inactivation of RV serotype G2 strain S2..………106
Figure 17. Effect of temperature on inactivation of RV serotype G3 strains.………….107
Figure 18. Effect of temperature on inactivation of RV serotype G4 strain ST3.. ……108
Figure 19. Direct comparison of pressure inactivation of six human RVs derived from
four serotypes…..……………………………………………………………109
Figure 20. HPP disrupts the integrity of RV particles..………………………..……….111
Figure 21. The effect of HPP on RV proteins…………………………………………..113
Figure 22. The effect of HPP on viral genomic RNA…………………….…………….115
xxiv Figure 23. Inactivation of RV Wa strain in oyster tissues by HPP…..…………………117
Figure 24. Models of viral inactivation by HPP………………………………………..124
Figure 25. Plaque sizes and log reduction of WT TV after heat treatment at 70˚C
for..…………………………………………………………….……….……137
Figure 26. Log reduction of TV heat-resistant variants ..……………….……………..138
Figure 27. Log reduction (log10 PFU/ml) of TV heat-resistant variants from passage 5
after heat treatment at 70˚C for 6s……………………….………………….139
Figure 28. Log reduction in TV titer after HPP treatment at 300 MPa for 2 min at
20˚C…………………………………………………………..…………….140
Figure 29. Log reduction (log10 PFU/ml) of TV HPP-resistant variants after HPP at 300
MPa for 2 min at 20˚C for 3 passages……………………………………..141
Figure 30. Growth kinetics and plaque morphologies of TV wild type and five isolates of the most heat and HPP resistant genotypes………….……………………..………….143
Figure 31. The infectivity of TV (WT) and two isolates from heat resistant, and three
HPP resistant genotypes in LLC-MK2 cells at MOI 0.01……………..…..144
Figure 32. Plaque size of TV (WT) and two isolates from heat resistant, and three HPP
resistant genotypes h…………….…………….…………………………..145
xxv
ABBREVIATIONS
ANOVA – Analysis of variance
CDC – Centers for Disease Control and Prevention
CPE – Cytopathic effect
DMEM – Dulbecco’s modified eagle medium
DNA – Deoxyribonucleic acid
EM – Electron Microscopy
FBS – Fetal Bovine Serum
FCV – Feline calicivirus
FDA – Food and Drug Administration
HAV – Hepatitis A virus
HBGA – Histo-blood group antigen
HPP – High pressure processing
HRV – Human rotavirus
HuNoV – Human norovirus
MA-104 - Rhesus monkey kidney cell line
MEM – Modified eagle medium
MK2-LLC – African green monkey kidney cell line
MNV-1 – Murine norovirus
MOI – Multiplicity of infection xxvi MPa - Mega Pascals
NIAID – National Institute of Allergy and Infectious Disease
ORF – Open reading frame
PCR – Polymerase chain reaction
PBS – Phosphate buffered saline
PFU – Plaque forming unit
PGM – Porcine gastric mucin
PGM-MB – Porcine gastric mucin conjugated magnetic beads qPCR – Quantative polymerase chain reaction
RAW 264.7 – Mouse macrophage cell line
RdRp – RNA dependent RNA polymerase
RNA – Ribonucleic acid
RT-PCR – Reverse transcriptase polymerase chain reaction
SDS-PAGE – Sodium dodecyl sulfate polyacrylamide gel electrophoresis
TEM – Transmission electron microscopy
TV – Tulane virus
VLP – Virus-like particle
WHO – World Health Organization
xxvii
CHAPTER 1 LITERATURE REVIEW
1.1. Introduction
Although foodborne outbreaks and illnesses have decreased in the last 10 years, they remain an important public health concern. An estimated 48 million illnesses each year are attributed to the consumption of contaminated foods in the U.S., resulting in 55,961 hospitalizations and more than 1,300 deaths. Human norovirus (HuNoV) is the leading cause of foodborne illness in the U.S. (Scallan et al., 2011; Scallan, Hoekstra, et al.,
2011). Despite an increased effort to document the number food outbreaks, the vehicle food, and the pathogen, only in a small portion of the foodborne outbreaks are reported to the CDC, and in many of the reported outbreaks the causative food and pathogen cannot be identified (CDC, 2014).
According to the CDC, during 2012 the reported foodborne outbreaks in U.S. acceded
830 and caused near 15,000 individual illnesses and 23 deaths (CDC, 2014). HuNoV was the cause of 41% of the cases in which a single etiological agent could be identified.
Seafood consumption was associated with 23% of the reported outbreaks (CDC, 2014).
1
Figure 1. Top pathogens leading to illness in the US (adapted from CDC, 2014)
Viruses in general were responsible for more than 50% of the identified outbreaks. Seafood, fresh produce, poultry, and meat accounted for more than 50% of the total outbreaks from 2001 to 2010. Fresh produce and seafood contributed to the highest number of outbreaks, responsible for 696 and 657 outbreaks, respectively. By comparing outbreaks‐per‐pound of food consumed, seafood is the highest risk food with approximately 19 times higher risk than fresh produce (CSPI, 2014). HuNoV and
Salmonella spp. are the most commonly reported agents in food associated outbreaks
(CSPI, 2014). Food contamination with enteric viruses can occur during pre-harvesting or post-harvesting stages, including growing, harvesting, processing, transportation, or from the food handler.
Seafood is a nutritionally rich food containing high-protein, omega-3 fatty acids, vitamin B12, zinc, and has a low-fat content, which provides a wide range of health benefits
2 (CSPI, 2014). In the U.S., the daily consumption of seafood has increased during the last two decades by approximately 37% (U.S. Census Bureau, 2011). Economically, the U.S. imports 16.6 billion dollars’ worth of seafood and fishery products and importation increased by more than 10% in 2011. In addition, the U.S. exported more than $5 billion worth of seafood the same year (U.S. Census Bureau, 2011). Despite the economic value attributed to seafood, it remains a public health concern because numerous outbreaks related to the consumption of the contaminated seafood have been reported.
It has been estimated that seafood contributes 10-19% of all foodborne illness outbreaks in the U.S. Shellfish, as a subset of seafood, are particularly susceptible to contamination with pathogens. These bivalve filter-feeders sieve gallons of water a day through their gills which can lead to the bio-accumulation of both bacterial and viral pathogens within shellfish tissues. Approximately 7.6-14.5 million illnesses in the U.S. are attributed to the consumption of contaminated seafood each year, and foodborne viruses were responsible for more than 50% of these cases (Butt et al., 2004). Human norovirus (HuNoV), hepatitis A virus (HAV), rotavirus (RV), and human sapovirus are the most common viruses associated with shellfish related outbreaks. Therefore, the FDA must balance the risks and benefits of seafood consumption when making recommendations to the public about the health benefits of a diet rich in seafood. In fact, the FDA classifies seafood as one of the highest risk foods for consumption in the U.S.
Other government agencies, such as the USDA, also consider the safety of seafood an important issue in the U.S. (Allshouse et al., 2003).
HuNoVs account for more than 58 % of all foodborne outbreaks and result in
11% of the foodborne illness related deaths reported each year (Scallan, Hoekstra, et al.,
3 2011). HuNoV causes severe gastroenteritis characterized by vomiting, diarrhea, and stomach cramps. The mode of transmission of HuNoV is the fecal-oral route, with most disease transmission through contaminated food, fomites, or water. It has been a challenge to study HuNoV because it cannot be grown in a cell culture system and lacks a small animal model (Teunis et al., 2008). Therefore, cultivable viral surrogates that are closely related to HuNoV, such as murine norovirus (MNV-1), Tulane virus (TV), and feline calicivirus (FCV), have been used to study the survival of HuNoVs in foods and the environment (Bozkurt et al., 2014; Topping et al., 2009).
HAV causes gastroenteritis, liver damage, and jaundice. HAV is typically transmitted by food, although water is also a major transmission vehicle recognized in the last decade. HAV outbreaks have declined due to the improvements in drinking water quality and sanitation practices, as well as the availability of effective vaccines in developed countries. However, HAV remains endemic in developing countries, many of which export food to the U.S.(Jacobsen & Koopman, 2004). The presence of HAV in imported seafood remains a high concern for U.S. regulatory agencies.
RV is the major causative agent of gastroenteritis in children and infants worldwide, and in the U.S., 90% of children have experienced a RV infection by the age of 5. RV causes gastroenteritis primarily in children aged 6 months to 2 years old; most people above this age have already developed immunity. Dehydration caused by diarrhea often leads to death, so a vaccine has been developed for this virus and is recommended for children at 2 months of age (Ghazanfar et al., 2014). The common transmission route is fecal-oral, but fomites are often found to harbor and then disseminate the virus. Since
4 RV is transmitted mainly via the fecal-oral route, so consumption of contaminated food or water sources often lead to disease (Kittigul et al., 2014; Quiroz-Santiago et al., 2014) .
Despite major efforts, the survival of foodborne viruses in seafood is still poorly understood. It is known that foodborne viruses are highly stable in foods and the environment and only a few particles (10-1000 particles) are required to cause disease.
Previous work has evaluated the distribution of bacterial pathogens in shellfish, however little is known about the bioaccumulation and distribution of viral pathogens in shellfish.
Therefore, an understanding of the ecology and persistence of enteric viruses in shellfish is needed to help prevent future outbreaks. Overall, viruses are much more resistant than bacteria to methods of inactivation and removal from foods (Marino et al., 2005; Polo et al., 2014; Ramos et al., 2012). To date, most thermal and non-thermal inactivation methods for shellfish have been standardized to target bacteria. However, these inactivation parameters (e.g., time, temperature, dose, and pressure levels) may not be sufficient to inactivate foodborne viruses. Thus, there is urgent need to identify effective and feasible thermal and non-thermal food processing technologies to inactivate foodborne viruses in seafood.
1.2. Foodborne viruses.
Viruses transmitted by food are defined as food-borne viruses. They include
viruses from many families; however these viruses share many common
characteristics. The main unifying trait of food-borne viruses is that they are non-
enveloped viruses, lacking a lipid envelope. Table (1) shows the most prevalence
foodborne viruses.
5 Table 1. Summary of the major foodborne viruses.
Virus Genome Envelope Disease
Norovirus +ssRNA No Gastroenteritis
Adenovirus dsDNA No Gastroenteritis
Rotavirus dsRNA No Gastroenteritis
Sapovirus +ssRNA No Gastroenteritis
Astrovirus +ssRNA No Gastroenteritis
Aichivirus +ssRNA No Gastroenteritis
Hepatitis A +ssRNA No Jaundice, Hepatitis, Gastroenteritis
Hepatitis E +ssRNA No Juandice, Hepatitis, Gastroenteritis
Polio virus +ssRNA No Poliomyelitis
Note: +ssRNA: single-stranded positive-sense RNA virus; dsDNA: double-strandedDNA virus; dsRNA: double-stranded RNA viruses
Non-enveloped viruses, in general, are more resistant to heat, pH, drying, and organic solvents than enveloped viruses (Kovac et al. 2012, Kotwal and Cannon 2014). This environmental stability allows the non-enveloped viruses to be maintained in the food for long periods of time and to survive the acidic conditions found in the digestive tract. The stability of non-enveloped viruses also makes them more resistant to common sanitation methods and food processing technologies (Lou et al. 201a; Nowak et al. 2011; Bozkurt et al. 2014)
6 1.3. Human Norovirus (HuNoV)
HuNoV is the leading causative agent of nonbacterial gastroenteritis worldwide
(Glass et al., 2009; Verhoef et al., 2013). In the U.S., it is estimated that HuNoV accounts for more than 60% of foodborne illnesses each year (Scallan, Hoekstra, et al.,
2011). HuNoV infects individuals of all ages and outbreaks frequently occur in crowded places, such as cruise ships, hospitals, restaurants, hotels, day care centers, schools, nursing homes, cruise ships, swimming pools, and military installations (Holtby et al.,
2001; Kassa, 2001). HuNoVs are estimated to cause over 200,000 deaths a year among children in the developing countries (Patel et al., 2008). In the U.S., HuNoVs are the second leading cause of gastroenteritis-related mortality, causing 797 deaths annually
(Patel et al., 2009; Clair & Patel, 2008) .
HuNoV is highly stable and contagious, and a few particles (usually less than 10) can cause disease (Teunis et al., 2008). Therefore, HuNoV is considered as a category B biodefense agent by the National Institute of Allergy and Infectious Diseases (NIAID). It has been a challenge to study HuNoV as it cannot be grown in cell culture and lacks a small animal model. As a consequence, the survival of HuNoV in food and environment is poorly understood. Currently, there is no anti-viral drug or vaccine against HuNoV.
1.3.1. Molecular biology of HuNoV
HuNoV is a non-enveloped, single-stranded positive-sense RNA virus. HuNoV belongs to the Caliciviridae family under the genus Norovirus. The Caliciviridae family includes other five genera: Vesivirus, Lagovirus, Becovirus, Recovirus, and Sapovirus, which contain viruses causing a wide array of diseases in animals (respiratory disease,
7 hemorrhagic fever, and gastroenteritis), however the viruses in this family which infect humans cause mainly gastroenteritis. The genus Norovirus is divided into five genogroups, GI-GV. All noroviruses that infect humans are found in genogroups GI, GII, and GIV; genogroup II (GII) and III (GIII) noroviruses infect pigs, sheep and cattle, and genogroup V (GV) noroviruses infect mice (Karstet al., 2003; Liu et al., 1999; Sugieda et al., 1998). Recently, Mesquita et al. (2010) isolated a new norovirus strain from dogs with diarrhea which is proposed to belong to a new genogroup, GVI (Mesquita et al.,
2010).
Figure 2. Norovirus genogroup and genotype characterization based on sequence
homology of the VP1 (major capsid protein) (Adapted from CDC, 2011).
8 The genome of HuNoV is 7.5-7.7 kb in length and is divided into three open reading frames (ORFs) (Hennessyet al., 2003; Jianget al., 1993). The genome is linked to the VPg protein at the 5’ end and is polyadenylated at the 3’ end (Karst et al., 2003;
Pletneva et al., 2001). Both ends of the genome have conserved regions that play a role in virus replication and pathogenicity (McFadden et al., 2011; Simmonds et al., 2008).
Murine norovirus, the only cell culture cultivable norovirus, has an additional ORF,
ORF4, located in the subgenomic RNA used for translation of VP1 and VP2. ORF4 encodes virulence factor 1 (VF1) which is translated by a ribosomal frame shift from the subgenomic RNA (Herod et al., 2014; McFadden et al., 2011).
Table 2. HuNoV proteins and their functions
Protein Function p48 Inhibits host protein trafficking Formation of replication complex
NTPase NTPase activity p22 Inhibits host protein secretion Involved in formation fo the replication complex
VPg Viral RNA replication primer Recruitment of translation initiation factors
3C Protease RdRp RNA dependent RNA polymerase VP1 Major capsid protein VP2 Minor capsid protein Particle stabilization
9 1.3.2. HuNoV infection:
The primary symptoms of a HuNoV infection include diarrhea, vomiting, nausea, and mild fever. Infected individuals can shed up to 9 log10 RNA copies/g of feces during symptomatic infection (Atmar et al., 2008). HuNoV can also be shed during asymptomatic infection or after recovery from symptoms for over a month (Leon et al.,
2011). The duration of fecal shedding is longer for immunocompromised individuals and children (Furuya et al., 2011). Milbrath et al. (2013) conducted a study to correlate norovirus-shedding duration of infected individuals with the potential and severity of norovirus outbreaks. The study divided the shedding duration into two types: regular shedding period (between two to three weeks) and long shedding period (from 100 to 140 days) and developed a norovirus transmission model. The model suggests that the presence of individuals with longer HuNoV shedding duration in a community increased the chance of an outbreak by 50 to 80%, the severity of illness symptoms by 20%, and subsequent viral transmission among the population by 100% (Milbrath et al., 2013).
This study highlights that individuals with prolonged HuNoV shedding
(immunocompromised individuals and children) may be “super-spreaders” leading to increases in disease propagation and severity.
Not all individuals are susceptible to HuNoV infections. Volunteer studies have shown that individuals with secretor status, who have mutated FUT2 gene at positions
428 and 571, are more susceptible to HuNoVs infections. For instance, Kindberg et al.
(2007) found a strong correlation between secretor status of 61 patients from five HuNoV outbreaks in Denmark and the susceptibility to the infection. The study included individuals from five outbreaks occurring between 2004 and 2005 in Denmark. The
10 symptoms included vomiting diarrhea and nausea and RT-PCR results of the stool samples of the affected patients were positive for norovirus G.II.4. DNA from infected individuals as well as uninfected contacts was evaluated for polymorphisms in FUT2.
The results showed a strong correlation (p<0.003 by using Fisher exact test, two-tailed,
95% confidence interval) between the susceptibility to the HuNoV infection and secretor status (i.e. FUT2 mutations). Thus individuals with non-secretor status are less susceptible to the human norovirus infection(Kindberg et al., 2007).
1.3.3. Human norovirus stability:
Studies have shown that HuNoV is highly stable in the environment. For instance,
Ngazoa et al. (2008) studied the persistence of GII.4 HuNoV in different types of water
(treated sewage, mineral water, tap water, and river water (St Lawrence River, Canada)) at different storage temperatures (-20, 4, and 25˚C) for 20, 40, 60, 80, and 100 days. The initial virus concentration and the level of survived virus after storage were evaluated by qRT-PCR. The results showed that, in general, at low temperature storage (-20 and 4˚C)
HuNoV was more stable than at 25˚C. The virus was stable in mineral and tap water for
60 days under all experimental temperatures, however the virus genome was degraded after 100 days. The stability of HuNoV in sewage treated water and river water was reduced compared to tap water and river water. The viral genomic RNA was degraded after one month of storage in sewage and river water at all storage temperatures. This study highlights that HuNoV is stable in aqueous solutions for long periods of time. In addition, other studies have indicated that HuNoV (GI) can persist for 3 years in water
(Seitz et al., 2011).
11 1.3.4. Cell culture systems for HuNoV:
Despite exhaustive efforts, attempts to cultivate HuNoV have failed to adapt the virus to in vitro culture systems. Early studies found that Norwalk virus like particles
(VLPs) were able to attach to differentiated Caco-2 cells (White et al., 1996).
Differentiated Caco-2 cells resemble mature enterocytes, express H antigen, and were originally derived from an individual with blood type O (Amano & Oshima, 1999).
However, replication of HuNoV clinical isolates in differentiated Caco-2 cells has remained unsuccessful. More recent attempts to cultivate HuNoV have involved the use of cell culture systems using combinations of gastric cells, duodenal cells, and small intestine enterocyte-like cells (Duizer et al., 2004). These combined cell culture systems have failed to replicate HuNoV, even with the addition of additives such as proteases, hormones, and intestinal contents to further mimic the intestinal micro-environment
(Duizer et al., 2004).
Guix et al. (2007) reported that the transfection of HuNoV RNA into human hepatoma Huh-7 cells lead to viral replication. Increases in VPg and VP1 protein expression were used as indicators of viral replication. The study showed some viral particles were released to the medium, however the resulting progeny viruses were unable to infect new cells (Guix et al., 2007).
Most recently, a study conducted by Jones et al. (2014) attempted to cultivate human (HuNoV) and murine norovirus (MNV) in human and mice B cells in the presence of gut bacteria in vitro. Mouse B cell lines M12 and WEHI-231, as well as
RAW264.7 macrophages were infected with MNV-1 and MNV-3.The virus titer of
MNV-1 and 3 were detected by using standard TCID50 assay. The results showed that
12 mice B cell lines supported the growth of MNV-1 and 3.Based on the fact that MNV was found to replicate in B cells, the BJAB human B cell line was infected with HuNoV genogroup II, genotype 4 (GII.4) Sydney strain. Filtered and unfiltered stool samples containing HuNoV were used to infect the B cells. RT-qPCR was used to monitor viral
RNA levels following infections. Western blotting and an immunofluorescence assay were used to detect HuNoV GII.4 structural and non-structural proteins. The results showed that the level of HuNoV RNA increased by 10-fold and 25-fold genome copies in the BJAB human B cell line after 3 and 5 days post infection when unfiltered inoculum was used. However, when filtered samples were used as the inoculum the virus replication was decreased. This suggested that there was a filterable cofactor required for
HuNoV replication in B cells. It was found that bacteria expressing HBGAs were required for successful replication of HuNoV in B cells, and that adding the bacteria or synthetic HBGAs to filtered HuNoV inoculum rescued the ability of the virus to replicate in B cells (Jones et al., 2014). These results are highly promising, however, it remains unknown whether this system is robust enough to test the survival of HuNoV in vitro.
1.3.5. HuNoV surrogates.
Currently, the survival of HuNoV during food processing is poorly understood because of its inability to grow in cell culture. Therefore, research on the effectiveness of food processing technologies to inactivate HuNoV has relied on surrogates. These surrogates include viruses that are closely related to HuNoV in terms of genetic makeup, size, receptor binding, pathogenicity, and environmental stability. Other surrogates used for the study of HuNoV include virus-like particles (VLPs) and P domain-particles (P-
13 particles). These particles resemble portions of the HuNoV protein capsid, which are important for receptor binding of the virus to the host cell and antigenic recognition of the virus by the immune system. The particles are non-infectious due to the fact that they are composed only of protein and lack the viral genome component of the native virus.
While the use of surrogates has aided in the understanding of HuNoV, there are several limitations in comparing data generated from the use of surrogates to HuNoV (Bozkurt et al., 2015; Bozkurt et al., 2014).
1.3.5.1. HuNoV virus-like particles (VLPs)
The norovirus virion is not enveloped and the protein capsid is composed of 180 monomeric protein units of VP1. These 180 proteins are further organized into 90 dimers. The VP1 protein can be divided into three domains; N, S, and P. While N and S function to stabilize the viral particle and are found internally, the P domain is positioned externally on the viral particle and can be further subdivided into P1 and P2 (de
Rougemont et al., 2011). The P2 domain of VP1 is responsible for the receptor binding and the attachment to cells. While HuNoV cannot be grown in cell culture, the use of recombinant baculovirus expression systems has allowed for the production of the VP1 protein in an insect cell line. The VP1 protein can then self-assemble to form HuNoV virus-like particles (VLPs), which retain the same capsid structure and antigenicity as the native virus (Fig.3). The HuNoV VLPs lack the genome component of the native virus and therefore are noninfectious. Interestingly, the expression of VP1 results in the formation of two sizes of VLPs. The diameters of the large and small particles were between 30-38 nm and 20-23 nm, respectively (Kamata et al., 2005). The VLPs have
14 been found to bind readily to the HuNoV cellular receptor, the histo-blood group antigens
(HBGAs) (Gandhi et al., 2010).
There are a number of advantages of using VLPs as a surrogate to study HuNoV.
First, HuNoV VLPs can be produced in large quantities by expressing VP1 in a number of systems such as the recombinant baculovirus grown in insect cells, as described above.
Secondly, damage to VLPs can be evaluated using biophysical and biochemistry methods such as electron microscopy (EM), sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), and immunoblotting assays. Finally, VLPs possess authentic receptor binding activity which is essential for viral infection. Hence, VLPs can be assayed for receptor binding activity as an indicator of virus survival.
Figure 3. Transmission electron microscopy image of human NoV virus-like
particles (VLPs) (Adapted from Feng et al., 2011).
15 1.3.5.2. Tulane Virus
Tulane virus (TV) is a cultivable calicivirus that was recently isolated from the stool of rhesus macaques with symptoms of gastroenteritis and represents a new genus in the family Calicivirus, Recovirus (Farkas et al., 2008). Despite the fact that TV is not a member of norovirus genus, it is genetically closely related to GII HuNoVs and utilizes the histo-blood group antigens (HBGAs) as a cellular receptor, as HuNoV does
(Farkas et al., 2010). TV is a single-stranded, positive-sense RNA virus, with a genome size of 6714 bp, similar to HuNoV. The genome of TV contains three open reading frames (ORFs). ORF1 encodes the nonstructural (NS) polyprotein, ORF2 encodes the major capsid protein (VP1), and ORF3 encodes the minor capsid protein (Farkas et al.,
2008). TV is a promising surrogate for HuNoV for several reasons. First, TV is an enteric virus similar to HuNoV and it recognizes the authentic receptor for HuNoV, HBGAs. In addition, Cryo-Electron Microscopy (Cryo-EM) results show that the structure and the organization of the TV major capsid protein is similar to HuNoV (Yu et al., 2013).
Figure 4. Transmission electron microscopy image of Tulane virus (TV). 16 1.3.5.3. Murine Norovirus
Murine norovirus (MNV) was first isolated from immunocompromised mice in
2003 (Karst et al., 2003). MNV is genetically closely related to HuNoV, and to date it is the only virus in the genus Norovirus that can be grown in cell culture. Therefore, it has been widely used to understand norovirus gene expression and as a surrogate for HuNoV.
MNV belongs to genogroup V (GV) noroviruses and its infection is lethal to immunocompromised (STAT-/-) mice (Karst et al., 2003). MNV recognizes a different receptor, sialic acid, compared to HuNoV. The capsid of MNV has a size of 28-35 nm in diameter with icosahedral structure. MNV is a single-stranded, positive-sense RNA virus with a genome size of 7,382 kb. The genome contains three ORFs. ORF1 with a length of
182.5-kD encodes the nonstructural proteins. ORF2, 58.9-kD, encodes for the major capsid protein (VP1) and ORF3, 22.1-kD, encodes for the minor capsid protein (Karst et al., 2003). Recent studies have provided evidence that MNV contains an extra ORF,
ORF4. ORF4 encodes virulence factor 1 (VF1), which has been found in MNV and in saproviruses as well (Clarke & Lambden, 2000; McFadden et al., 2011). ORF4 is encoded in an alternate reading frame in the subgenomic RNA. ORF4 has been shown to modulate the innate immune response and mutant MNV-1 lacking ORF4 had decreased virulence in mice (McFadden et al., 2011).
17
Figure 5. Transmission electron microscopy image of murine norovirus (MNV-1).
1.3.5.3. Comparing murine norovirus and Tulane virus as human norovirus surrogates
MNV and TV are widely used as surrogates to mimic the HuNoV response to several thermal and non-thermal treatments (Bozkurt et al., 2013; Bozkurt et al., 2015).
Hirneisen and Kalmia (2013) compared the stability of the most commonly used human norovirus surrogates, MNV and TV, to heat and chlorine treatment. MNV and TV were exposed to low temperature treatments ranging between 20-75˚C in PCR tubes for 2 min.
The chlorine treatment was conducted at concentrations from 0.2-2,000 ppm for 5 min at the room temperature. The environmental stability of MNV and TV was evaluated in tap water stored at either 20 or 4 ˚C. The stability of the viruses at a range of pHs (2-10) was also determined. The level of surviving viruses following each treatment was determined by plaque assay. In tap water at 4˚C, MNV was stable for 30 days, whereas TV lost its infectivity after 5 days of storage. In addition, MNV was found to be more stable than TV
18 at low chlorine concentrations (<2 ppm), however this enhancement in stability of MNV over TV was abolished at higher chlorine concentrations. MNV was also more stable than TV at low and high pH values, however both viruses had less than a 1 log reduction at low pH values (Hirneisen & Kniel, 2013).
Table 3. Comparison of MNV-1 and TV to HuNoV
MNV-1 TV Human Norovirus
Genetic material Positive single- Positive single- Positive single- stranded RNA stranded RNA (6.7 stranded RNA (7.3 kb) kb) (7.7 kb) Enteric vs. Non- Non-enteric Enteric Enteric enteric Particle size 27-38 nm 36 nm 28-35 nm
Enveloped vs. Non-enveloped Non-enveloped Non-enveloped non-enveloped Receptor Sialic acid Histo-blood group Histo-blood group antigens antigens
1.4. Rotavirus (RV)
Rotaviruses (RVs) are the major etiological agent of acute gastroenteritis in infants worldwide, and account for 27% of deaths in children, according to the WHO
(Lanata et al., 2013). It is estimated that RVs cause 500,000 deaths each year among children worldwide (CDC, 2013; WHO, 2011; WHO, 2013). The virus is transmitted via 19 the fecal-oral route, secondary transmission often occurs via person-to-person contact, contaminated food and water sources, and also through contaminated fomites. The symptoms include moderate to severe watery diarrhea, vomiting, fever, and abdominal discomfort; with onset time ranges between 24 h to one week (R. M. Lee et al., 2013).
Despite the success of vaccines against RVs in the reduction of gastroenteritis among infants, RVs are still of major public health importance and the leading cause of severe diarrhea in the U.S. in children under five (CDC, 2013). In addition, RVs are highly stable in food, water, and other fomites. RV is highly infectious among children due to its low infectious dose (<10 particles can lead to disease and the fact that the virus is shed at a very high titer (1010 to 1012 virus particles/g of stool) (Bishop, 1996; Ward et al.,
1986).
RV contamination of shellfish growing areas, drinking water, and irrigation water sources can easily occur from sewage discharges. The contamination of these water sources with RV can then lead to virus contamination of foods. For instance, RV was detected in drinking water and vegetables in France, Kenya, South Africa, and
Netherlands (Gratacap-Cavallier et al., 2000; Kiulia et al., 2010; Rutjes, Lodder et al.,
2009; van Zyl et al., 2006), which may increase the risk of RV outbreaks. Moreover, new strains of RV are constantly emerging because of intra-genogroup reassortment of RV strains and may impact vaccine effectiveness in the future. These data strongly suggest the importance of implementing a new prevention approach to insure the safety of food and drinking water.
20 1.4.1. Rotavirus molecular biology
RV was first isolated from humans in the 1970s by electron microscopic imaging of the duodenums of infected children (Bishop et al., 1976). RV belongs to the
Reoviridae family. The mature virion is a non-enveloped icosahedral particle 100 nm in diameter (including the spikes). The RV particle is comprised of a triple-layered capsid containing 11 segments of double-stranded RNA with size of 16,500-21,000 nucleotides.
The genome encodes six structural proteins, classified based on their molecular weight into VP1, VP2, VP3, VP4, VP6, and VP7 and six non-structural proteins (NSP1-6).
RVs are highly diverse, both antigenically and genetically. There are eight species
(groups) of rotavirus, referred to as A, B, C, D, E, F, G, and H. Humans are primarily infected by species A, B and C, most commonly by species A. Rotavirus species A can be further divided into different serotypes. RV is a segmented double-stranded RNA virus with a triple-layered icosahedral capsid. The outer capsid glycoprotein (VP7) and the spike protein (VP4) differentiate RVs into 14 G (Glycoprotein) serotypes and 27 different P (Protease sensitivity) genotypes (van Zyl et al., 2006).
21
Figure 6. Rotavirus genome, proteins, and structure (adapted from Pesavento,
2006). a) PAGE showing rotavirus RNA segments and gene–protein assignments. b)
Surface representation of the mature rotavirus (triple layered particle TLP). c) Cut-away
of the TLP structure showing the internal structural features.
Currently, five serotypes (G1–G4, and G9) are the predominant circulating viruses, accounting for almost 95% of strains worldwide. Recently, commercial RV vaccines have been used in children to provide immunity against the most commonly circulating strains (Pacilli et al., 2015). Despite major efforts, RV outbreaks still occur worldwide due to the high genetic diversity of RVs and lack of cross-protection.
Therefore, alternative strategies for the prevention of RV infection must be established.
22 Table 4. RV structure and non-structure proteins
(Adapted from Pesavento, 2006)
1.5. Hepatitis A virus (HAV)
HAV is transmitted mainly by the fecal-oral route via contaminated food or water.
The symptoms include fatigue, nausea, vomiting, abdominal pain or discomfort, low- grade fever, and yellow skin and eyes. The symptoms can last between two weeks to several months. HAV is a picornavirus, and it has only one serotype. The genome of
HAV is a positive-sense RNA molecule with a central open reading frame encoding a polyprotein that is flanked by highly structured, non-translated regions (5′ NTR and 3′
NTR) and a 3′ poly(A) tail (Balayan, 1989; Balayan, 1992; Kusov, 1990). Despite the
23 fact that a vaccine is available for HAV, outbreaks of HAV continue to occur, especially in developing countries. In the developed world, an individual may acquire HAV from international travels and outbreaks can usually be traced to a contaminated high risk food
(such as seafood and fresh produce). From 1991 to 2007 a total of 268 HAV outbreaks in the US were reported (Pereira, 2004; Craig et al., 2007; Torner et al., 2012).
1.6. Limitations in current food safety controls to eliminate viruses from high risk foods
Despite the fact that viruses contribute to more than 50% of the foodborne illnesses, most food handling and microbial inactivation regulations are based on bacterial pathogens. However, based on the inherent differences between bacteria and viruses there are several limitations when applying control strategies developed for bacteria to eliminate viruses in the food supply.
First, bacteria and viruses have different structural and biological properties.
Viruses are Akaryotic organisms that are obligate parasites. The viral genomic material can be RNA or DNA wrapped in a protein structure, called the viral capsid. Viruses are small in size, lack metabolic machinery, and cannot replicate within the food matrix.
Thus, the regular food preservation strategies, such as refrigeration and modified atmospheric packaging that used to inhibit bacterial growth, are not effective to control or eliminate foodborne viruses from food during transportation and storage.
Second, the levels of viral contamination in foods are usually very low. There is no method for enrichment of viruses in vitro prior the detection as there are for bacteria.
24 This requires that sensitive methods and procedures that are different from those used for bacterial numeration in foods are needed for viral detection.
Third, only a small number of viral particles are sufficient to cause disease (<100 particles). Moreover, most bacteria species can only grow in a narrow spectrum of food
(eg. low acid food vs. acid food), that provides the bacteria the optimal growth conditions. Foodborne viruses can persist in most food types regardless of pH, aw, oxygen conditions, and temperature. Therefore, foodborne viruses are highly infectious and can be transmitted via most types of contaminated food.
Most of foodborne bacteria can be grown in synthetic media, which facilitates detection and identification. The most common detection methods for foodborne pathogenic bacteria are PCR, ELISA, API systems, culture-based methods, specialized enzyme substrates, antibodies, other DNA based methods, and serological approaches. In contrast, several foodborne viruses are very difficult or impossible to cultivate in vitro.
Detection of the viral genomic material via PCR based methods is also difficult due to the low level of virus contamination and the presence of PCR inhibitors in the food matrix.
Overall, the removal and detection of viruses in foods has proved more challenging than for foodborne bacterial pathogens.
1.7. Detection methods for foodborne viruses
In contrast to bacteria, viruses have a simple structure and lack the ability to replicate in synthetic medium which limits the detection approaches for food samples to the
25 nucleic acid detection methods (Kamata et al., 2005; Morton et al., 2009; Baert et al.,
2008).
PCR or RT-PCR is the most widely used tool to detect viruses in food and environmental samples. PCR based methods are simple, sensitive, cost effective, and can discriminate between genetically closely related viruses. The disadvantage of traditional
PCR or RT-PCR is that they are qualitative methods and at best semi-quantitative.
Inhibitors from the food matrix can also affect the results, and this often requires extraction and concentration of the virus before the test is conducted. RT-qPCR is used to overcome some of the disadvantages of traditional PCR and RT-PCR. To date, RT-qPCR is the only quantitative method to detect the HuNoV. However, a major limitation of nucleic based detection methods is that they cannot discriminate between viral nucleic acid from an infectious or non-infectious particle. Therefore, viral RNA detection in foods may overestimate the level of infectious virus present in the commodity.
Some immunological assays are available for foodborne virus detection such as
ELISA, however these assays are hampered by the limited cross-reactivity of antibodies for HuNoV. Immunological assays also require complicated method for viral extraction and concentration before the assay can be performed. Electron microscopy can also be used to detect viruses. However this method requires a high number of viral particles in the sample, a trained observer is need to identify the virus, and is expensive. Therefore,
EM may be used for clinical diagnosis but is not used for detection of viruses in foods.
Cell culture based assays such as plaque assay and TCID50 are an important method of detection for viruses as they are able to detect infectious virus. The limitations of these cell culture based assays is that not all viruses can be cultivated in vitro, and those viruses
26 that are cultivatable require long incubation times to obtain results (7-9 days for HAV and 3 days for RV) (Arnold et al., 2009; Baert et al., 2008).
Viral detection in food and environmental samples requires viral extraction and concentration prior the execution of the detection method. For instance, Haramotoa et al.
(2009) developed two virus concentration methods and compared the newly developed methods to a conventional concentration method that uses an electropositive filter
(1MDS-method). The two new methods used electronegative filters (Mg-Method and Al- method). The three methods were evaluated for the ability to recover a HuNoV GII. 4 strain and poliovirus (PV) type 1 from different types of water. The level of recovered viruses using each method was determined using RT-qPCR. The new methods (Mg- method and Al-method) showed comparable results to those obtained by the 1MDS- method for both viruses tested (HuNoV and PV). The results provide an indicator of the benefits of using the concentration methods for concentrate and detect the non-cultivable
HuNoV (Haramotoa et al., 2009).
1.7.1. HuNoV receptor binding assays for viral detection in foods
Since human NoVs cannot be grown in cell culture, viral RNA, viral proteins, or viral particles are targets for detection. Limitations for NoV detection are low concentration of viruses in a sample and extreme genetic and antigenic diversity seen within the genus Norovirus. There are no cross-reactive antibodies which can detect all circulating strains using enzyme immunoassays (EIAs). Likewise, nucleic acid detection assays are also hampered by low sequence homology because of genetic diversity. Thus, a single primer pair is insufficient for detecting all NoV strains and yet be free of false
27 positive reactions. For viral particle detection, electron microscopy (EM), immune electron microscopy (IEM), and solid-phase immune electron microscopy (SPIEM) can all be used but are expensive, require a highly trained observer to distinguish NoVs from other enteric viruses, and a large number of outbreak specimens cannot be rapidly examined.
Initially, RNA detection methods for HuNoVs were reverse transcription polymerase chain reaction (RT-PCR) assays (Atmar & Estes, 2001; Green et al., 1993).
Currently, RT quantitative PCR (RT-qPCR) assays are considered to be the “gold standard” for huNoVs detection and are used in many public health, clinical, food, environmental and research laboratories (Antonishyn et al., 2006; Kageyama et al., 2003;
Richards et al., 2004). In addition to RT-PCR and RT-qPCR other amplification variations, such as RT multiplex PCR, RT-nested PCR, direct RT-PCR, RT-nested, real time PCR, RT-booster PCR, and nucleic acid sequence-based amplifications – NASBAs, have been used for the detection of NoVs in various specimens (Antonishyn et al., 2006;
Boxman et al., 2011; De Medici et al., 2007; Pang et al., 2005). Recently, a reverse transcription loop-mediated isothermal amplification (RT-LAMP) approach has also been used for the rapid detection of HuNoVs (Fukuda et al., 2006; Yoda et al., 2009). In order to have a 90% probability for detecting a HuNoV as an etiological agent for an outbreak, at least three samples need to be tested using a standard RT-PCR assay (Duizer et al.,
2007). However, a major limitation of nucleic acid based detection methods is that they do not discriminate between infectious and non-infectious viral particles.
An intact capsid and genome are required for HuNoV replication and infectivity.
However, for some single stranded positive sense RNA viruses, such as polio virus, viral
28 genome RNA alone can be infectious in vitro (Holland et al., 1960). In the host and the environment, the viral capsid is necessary for protection of the viral genome from degradation. The viral capsid is also responsible for attachment to host cells, receptor binding, entry, and subsequent release of the viral genome into the host cell. Therefore, determining viral capsid integrity along with the detection of genomic RNA is a more accurate method for estimating viral infectivity compared to either one alone.
Several methods have been employed to estimate viral capsid integrity. A combined enzyme approach using Proteinase K, to degrade damaged capsid protein, and
RNase A, to degrade free viral RNA, was described to evaluate virus survival following heat treatment (Nuanualsuwan & Cliver, 2002). Heat inactivated polio virus RNA was still detected using RT-qPCR and using RNase A pre-treatment alone reduced the RNA detected, but did not completely abolish it (Nuanualsuwan & Cliver, 2002). However, this treatment did not abolish residual RNA detection of MNV-1 RNA after complete inactivation of the virus by heat (Baert et al., 2008). It was theorized that the residual virus RNA was being protected from RNase degradation by partially degraded capsid protein, so RNase A pre-treatment was combined with Proteinase K pre-treatment. The proteinase treatment approach was later abandoned because efficient Proteinase K digestion requires Ca ions for enzyme stability and prevention of self-proteolysis, the enzyme is also capable of degrading intact virus particles, and the enzyme can degrade
RNase.
RNase pre-treatment alone has been utilized to evaluate viral capsid integrity. In simple matrices, it has been found that HuNoV surrogates have good correlation between
RNase pre-treatment RT-qPCR and infectivity assays (Nowak et al., 2011; Topping et al.,
29 2009). However in stool suspensions, HuNoV GII.4 RNA detection was not affected by
RNase pre-treatment and this was comparable to surrogate viruses in stool suspensions
(Nowak et al., 2011; Topping et al., 2009). The complexity of the matrix in which the virus is suspended can affect enzyme function and provide an abundance of alternative enzyme substrates and can therefore interfere with the effectiveness of the assay.
A more direct approach to determining the integrity of the HuNoV capsid is to evaluate the capsid for its binding ability to the functional HuNoV receptors, the HBGAs.
HuNoVs have a high degree of binding affinity for the glycopeptide moieties associated with the HBGAs (Knight et al., 2013). Synthetic HBGAs have been used as HuNoV capture ligands in several studies and Caco-2 cells, which express HBGAs on their surface, have been used to evaluate HuNoV receptor binding ability (Cannon & Vinje,
2008; Knight et al., 2013; Morton et al., 2009). Mucins found in human breast milk obtained from Se+ individuals were found to inhibit the binding of Norwalk virus VLPs to their receptors, indicating these mucins contained HBGA-like molecules (Ruvoen-
Clouet et al., 2006). Similarly, porcine gastric mucin was found to bind to Norwalk virus
VLPs (Tian et al., 2005). Porcine gastric mucins are large heterogeneous extracellular glycoproteins with molecular weights ranging from 0.5-20 kDa. It has been determined that porcine gastric mucin contains a mixture of type A, type H1, and Lewis b HBGAs
(Tian et al., 2010). Theoretically porcine gastric mucin can be used to evaluate the ability of the HuNoV capsid to bind to its cellular receptor and estimate the potential of the viral particle to be infectious when used in combination with viral RNA detection.
Norwalk virus (GI.1) VLPs (rNVLPs) pre-incubated with varying concentrations of porcine gastric mucin (PGM) and incubation of rNVLPs with PGM at a concentration
30 of 20ng/mL was found to inhibit rNVLP binding to HBGAs by 85% (Tian et al., 2005).
Using an ELISA based assay, the binding of rNVLPs to PGM was found to be dose dependent and rNVLP binding to PGM was blocked by pre-incubation of the rNVLPs with HBGAs from human saliva or synthetic Lewis b and Lewis d antigens (Tian et al.,
2005). Additionally, pre-incubation of rNVLPs with PGM also significantly reduced rNVLP binding to Caco-2 cells (Tian et al., 2005). These results indicate that HuNoV binds to PGM in a specific manner and the interaction of HuNoV with PGM inhibits the ability of the virus to bind to its functional cellular receptor, the HBGAs.
Subsequently, PGM was conjugated to magnetic beads (MBs) and was used to capture and concentrate a HuNoV GII.4 strain from simple mediums and viral RNA was then detected by RT-qPCR. It was observed that using the PGM-MB binding assay lead to a 2 log increase in the viral RNA detected compared to the RNA detected from unbound HuNoV GII.4 samples (Tian et al., 2008). In oyster homogenate inoculated with HuNoV GII.4 the PGM-MB binding assay lead to RNA detection similar to the input virus level. However, when direct RT-qPCR was used as the method of detection, no HuNoV GII.4 RNA was detected (Tian et al., 2008). These results indicate that the
PGM-MBs concentrate HuNoV and increase the sensitivity of viral detection and also effectively remove PCR inhibitors in complex matrices.
The PGM-MB binding assay was then employed to determine the survival of
HuNoV GI.1 following high pressure processing (HPP) (Dancho et al., 2012). The level of HPP required to inhibit HuNoV GI.1 receptor binding ability as detected by the PGM-
MB binding assay was similar to the level of HPP required to inactivate HuNoV GI.1 seeded in oysters as determined in a human volunteer study (Dancho et al., 2012; Leon et
31 al., 2011). Based on these results, it was concluded that the PGM-MB binding assay was able to detect loss of human NoV receptor binding ability and was an improved method for determining HuNoV survival following inactivation treatments. The PGM-MB assay has also been used to determine HuNoV survival following heat treatment, freeze-that cycling, and exposure to sanitizers (Kingsley et al., 2014;Li et al., 2012; Tian et al.,
2012). Different HuNoV strain susceptibility to HPP has also been investigated using the
PGM-MB binding assay (Li & Chen, 2015; Li et al., 2013). Overall, the PGM-MB binding assay is an improved method over direct RT-qPCR for detecting HuNoV inactivation.
1.8. Shellfish are a high risk food for virus contamination
Shellfish are subjected to enteric virus contamination during production, harvesting, processing, or from food handlers (Santo and Edge, 2010). It has been shown that viruses persist in shellfish for significantly longer periods of time than bacteria
(Cook and Ellender 1986; Power and Collins, 1990). The bacterial instability in the shellfish may be due to both humoral and cellular defense mechanisms, such as the phagocytic process that poses humoral defense factors such as lectins, lysosomal enzymes, and various antimicrobial peptides (Canesi, et al., 2002). Most enteric viruses can persist in food under normal production and storage conditions. In the case of shellfish, contamination of the growing area by sewage waste, because of poor sewage treatments or from flood and rainfall, is the major cause of virus accumulation in shellfish tissues. For instance, in 2013, Keller et al. conducted a study to investigate the unusually
32 high rates of annual gastroenteritis in Vitória Bay, Espírito Santo, Brazil that have been associated with consumption of shellfish (oysters, mussels, and crabs) for 14 months
(February 2008 to March 2009). The collected samples included samples of the growing waters and shellfish samples from Mangroves area, from where most of the consumed shellfish in Espírito Santo are harvested. The target microorganisms included the indicator bacteria Escherichia coli (E. coli) and gastroenteric viruses including HuNoV,
RV, and adenovirus. Surprisingly, there results showed that 100% of the examined shellfish samples were positive for E. coli, RV, and adenovirus. However, only 80% of the water samples were positive for these pathogens. Moreover, the viral and bacterial concentration in shellfish meats was 400 times higher than levels in the growing water.
HuNoV, which is responsible for more than 95% of nonbacterial food outbreaks in the U.S., was present in less than 5% of the samples. The study concluded that the reason underlying the high microbial contamination of the water and the shellfish was due to discharge of home sewage to the growing water, intentionally or by flood and rainfall (Keller et al., 2013). In another survey, Bellou et al. (2013) systematically reviewed reported food outbreaks to Medline, Embase, Scopus, PubMed,
Eurosurveillance Journal and Spingerlink, and a global electronic reporting system
(ProMED) that are related to shellfish and viruses from 1980 to July 2012. The investigation showed that more than 11 shellfish-borne viral were reported each year worldwide. HuNoV and HAV were the leading cause of the outbreaks (83% and 12%, respectively). Among shellfish, oysters were associated with more than 50% of the reported illnesses. Contamination of growing water by sewage was the most reported source of the viral contamination (Bellou et al., 2013). In the same trend, Polo et al.
33 (2015) studied the virus contamination levels of class B growing areas (conditionally approved, according to NSSP classification in the US) of the most important shellfish production area in the European Union, the Galician Rias area, for a year and a half
(2013-2015). The investigated viruses were HAV and HuNoV GI and GII in mussels, clams, and cockles. HuNoV was detected by RT-qPCR and HAV was detected using standard plaque assay. The results showed that more than 55% of shellfish samples
(mussels, clams, cockles) were contaminated with enteric viruses. HuNoV GI and GII, and HAV were detected in 58% and 10% of the positive samples, respectively. Moreover,
11% of the shellfish samples were positive for the three enteric viruses. The viral titer in contaminated samples ranged from 2-3 log10 RNA copies. In contrast with other studies, the study reported that the seasonal peak for viral contamination was during the warm months. The results emphasize the magnitude of enteric virus contamination in shellfish and may help in the adoption of new legislation that sets viral critical values to improve food safety (Polo et al., 2015).
1.8.1. Seafood production practices
As defined by the FDA, seafood includes all fishery and seafood products, refrigerated, frozen, and fresh products, fin fish, shell fish (Mollusks and Crustaceans), turtles, frogs, and alligator. Seafood is a high-protein and low-fat food which is rich in omega-3 fatty acids, vitamin B12, and zinc. Thus, the consumption of seafood provides a range of health benefits such as improving infant neurological development and adult cardiovascular conditions. On the other hand, 7.6 to 14.5 million illnesses in the U.S. are
34 attributed to the consumption of seafood each year, and viruses were responsible for 50% of these cases (Butt et al., 2004).
Shellfish or bivalve molluscan shellfish, such as oysters, clams, mussels, and cockles, are filter feeders and are easily contaminated by a wide range of enteric pathogens, particularly viruses and bacteria through contaminated growing waters or food handlers.
Shellfish can bio-accumulate enteric viruses in their tissues and can concentrate the viruses up to 400 times in tissues compared to the level in the overlying water (Keller et al., 2013). It has been shown that viruses are persistent in shellfish and can survive most of the cleansing approaches that have been shown to be effective in eliminating pathogenic bacteria, such as depuration and relaying (Richards et al., 2010). Depending on the sanitary and fecal coliform levels, the growing water of shellfish can be divided into approved, restricted, or prohibited areas. Shellfish from approved growing areas are allowed to be sold directly to the public whereas shellfish harvested from restricted growing areas must be cleansed before sale, according to National Shellfish Sanitation
Program guidance (NSSP).
1.8.2. Seafood harvesting
Shellfish (especially bivalve Mollusks) are filter feeders, circulating the growing water through their gills. A single oyster, for example, can circulate up to 15 gallons of water per 24 h. By straining suspended food particles, shellfish may bio-accumulate large numbers of pathogenic viruses and bacteria from the harvesting waters and may act
35 as a passive vehicle for human pathogens. It has been shown that shellfish growing areas are the main source leading to enteric virus contamination of shellfish.
In the US, the National Shellfish Sanitation Program (NSSP) classifies shellfish growing waters based on generic E. coli or total fecal coliform levels into three classes.
Approved, from which the harvested shellfish may be directly distributed and consumed;
Conditionally Approved and Restricted, the harvested shellfish from these areas must be cleansed by pasteurization (by heat or pressure) or be subjected depuration before distribution; and Prohibited areas from which the harvested shellfish may not be harvested or distributed (Anonymous, 1999). In the European Union, the shellfish regulations ((EC) 852/2004, 853/2004 and 854/2004) classify the growing areas into four classes (A to D) based on the levels of generic E. coli or total fecal coliforms in harvested shellfish tissues. Shellfish harvested from class A areas may be directly consumed and distributed; class B shellfish must be cleansed before marketing; shellfish from class C harvesting areas must be subjected to prolonged cleansing procedures before distribution; and class D, where shellfish harvesting and distribution are prohibited
(Anonymous, 2004).
In both the US and the EU shellfish growing areas are regulated based on the total fecal coliforms or Escherichia coli (E. coli). However, it has been shown that using E. coli and/or fecal coliforms as criterion to evaluate the safety of the seafood does not predict virus contamination. To date, there is no good indicator organism for detecting enteric virus contamination in environmental or food samples. Nevertheless, the first step in ensuring the safety of shellfish is to restrict shellfish growing waters in accordance
36 with the applicable standards which aim to determine whether human waste contaminates the growing areas.
1.8.3. Post-harvest seafood processing
Post-harvest handling practices for shellfish can pose a risk of contamination and can also lead to outbreaks. Since enteric viruses are transmitted via the fecal-oral route, food handlers are often considered a point of contamination during shellfish sucking and packaging. Food handlers recovering from virus infection or those with asymptomatic viral infections have been shown to be leading cause of HuNoV contamination in foods.
In addition, contaminated ice used during storage and transport has been reported to be a source of shellfish contamination with both HAV and HuNoV (Beller 1992; Khan et al.
1994).
1.8.4. Bioaccumulation and distribution of viruses in bivalve mollusk tissues
Studies have shown that HuNoV bioaccumulation in shellfish is linked to the presence of carbohydrate ligands present in the shellfish tissues that closely resemble the cellular receptor of the virus (Maalouf et al., 2010, Maalouf et al., 2011). Histo-blood group antigens (HBGAs), the HuNoV receptor, have been detected in the digestive tissues of oysters, clams, and mussels. Schwab et al. (1998) showed that HuNoV is accumulated naturally in oyster and clam tissues within 24 h. The digestive system showed a high concentration of the viruses compared to muscles and gills throughout the
24 h period (Schwab et al., 1998).
37 Maalouf et al. (2010) compared the bioaccumulation of three HuNoV strains,
GI.1, GII.4, and GIII.3 in oyster (Crassostrea gigas) tissues. The GI.1 strain had the highest bioaccumulation level compared to the GII.4 strain in the digestive organs. This effect was increased in the winter season where the HBGA-like carbohydrate ligand expressed at a higher level in the oyster digestive organs. The GII.4 strain was detected in all tissues. This result is thought to be due to the fact that the GII.4 strain can recognize a sialic acid-containing ligand as an alternative receptor, which allowed for the bio- accumulation in all shellfish tissues. This may explain the high number of outbreaks caused by GI.1 strains in the winter and spring seasons (Maalouf et al., 2010). GIII.3 was accumulated most in the gills and mantle and was subsequently concentrated in digestive gland. These results demonstrate that different genotypes of HuNoV may have different bioaccumulation and distribution profiles in shellfish. .
To compare the distribution of different viruses in oyster tissues, a survey was conducted by Drouaz et al. (2015) comparing the distribution of HuNoV GI, TV, and mengovirus (MgV) in oyster tissues after 24 h of oyster cultivation in a contaminated seawater containing 7, 8.5, or 9 log10 RNA copies of HuNoV, TV, or MgV, respectively.
The results showed that HuNoV and TV were more accumulated in digestive glands, while MgV was more concentrated in gills. However, all three viruses persisted in oyster tissues for more than 50 days. HuNoV had the highest half-life in the tissues of 7.5 days compared to 4.65 days and 2.17 days for TV and MgV, respectively. These results indicate that different types of viruses bio-accumulate in different oyster tissues, and TV had a similar bioaccumulation profile as HuNoV in oysters and may serve as a good surrogate to study HuNoV bioacculumation in oysters (Drouaz et al., 2015). Hepatitis E
38 virus (HEV), which is an enteric pathogen that infects both humans and animals, has shown to be bio-accumulated and concentrated in the digestive tissues of shellfish
(oysters, flat oysters, mussels, and clams). Mussels and clams were more susceptible to
HEV contamination compared to oysters, indicating that the type of shellfish also influences viral bioaccumulation. The level of HEV bioacculuation in shellfish also showed a trend based on the seasonality, with the highest rate viral accumulation detected in March (Grodzki et al., 2014).
Taken together, this data indicates that viruses can attach to shellfish tissues via specific and non-specific binding, different viruses have different shellfish tissues in which they bioaccumulate, the level of virus bioaccumulation is also dependent on the type of shellfish, and finally that the season influences virus bioaccumulation in shellfish.
1.9. Control strategies for foodborne viruses in shellfish
Prevention is the most effective control strategy for control of virus outbreaks.
Since person to person contact is a common way to transmit the disease, a good hygiene plan can reduce virus infection and related illnesses. Although vaccines are available against some enteric foodborne viruses such as RVs and HAV, strain diversity limits vaccine efficacy to control foodborne outbreaks. In addition, HuNoV, the most prevalent foodborne pathogen, has no vaccine or antiviral drug to limit infection and outbreaks.
1.10. Depuration and relying
The depuration process has been used for shellfish cleansing for over a century.
It was developed as a method of cleansing oysters in response to typhoid outbreaks in
39 Europe and North America in the late 1800’s (Herdman & Scott 1896, reviewed in
Richards et al., 2010). In the US as early as 1911, it was demonstrated that depuration completely eliminated fecal coliforms in contaminated oysters (Phelps, 1911). The development of the process was a result of a collaboration between the oyster industry and the government in response to a large scale oyster related outbreak that occurred in
New York City.
Depuration is a commercial process in which the harvested shellfish are placed in flowing high quality seawater for specific time (at least 36 hours) under controlled temperature. This process allows the shellfish to purge contaminants from their gut using their natural filtration activity. Choosing healthy and active oysters, maintenance of water salinity, temperature, and dissolved oxygen are necessary to reach the highest oyster filtration activity. The depuration process is a batch process where the circulated seawater is filtered and sterilized using UV, ozone, or chlorine before it enters to the system
(reviewed in Richards, 2010).
Research on the conditions that affect the depuration efficacy to remove pathogenic bacteria and viruses from shellfish has been conducted. For instance, Love et al. (2010) conducted an experiment to study the effect of depuration temperature, turbidity, and algae concentration on the cleansing of artificially contaminated oysters
(Crassostrea virginica) and hard shell clams (Mercinaria mercinaria) in a flow-through system for 5 days. The examined depuration temperatures were 12, 18, and 25°C, turbidity levels of 1 NTU, 10 NTU, and 20 NTU, and algae concentration of 0 and 50,000 cells/ml. Two bacterial species (Escherichia coli and Enterococcus faecalis) and three viruses (coliphage MS2, Poliovirus type-1, and HAV) were used as indicators to evaluate
40 the effectiveness of the depuration system. Overall, the results showed that the algae concentration and turbidity did not affect the depuration of both oysters and shell clams, whereas, increasing the treatment temperature reduced the efficacy of the oyster cleansing process. In addition, it was found that the bacterial species were eliminated faster than the viruses in oyster and clam samples (Love et al., 2010).
Another study conducted by Chinnadurai et al. (2014) examined the effect of oyster load and oyster tray position in a fill-draw system on the elimination of the indicator bacteria, E. coli by depuration. The artificially contaminated oysters were cleansed by relaying the oysters in a flow of clean water, for 44 h. The system consisted of cartridge-filters to remove particulates from the cleansing water, a UV source to inactivate microorganisms, a temperature control, and a pH control. Oysters were placed in trays at either the bottom or the surface of the system, for 0 to 44 h. Before cleansing, the levels of pathogenic bacteria and viruses levels in the oysters was higher than the
NSSP recommended levels. The results showed that the oysters in the bottom of the system needed 2 days to achieve acceptable E. coli, levels, whereas the surface oysters were cleansed within 24 h. Moreover, the results showed that reducing the load of oysters in the depuration system (2 oysters/liter of water) resulted in a significant improvement in the depuration process. The results point out that using one layer of trays and reducing the number of oysters loaded in the system reduced the cleansing time from
48 h to 24 h (Chinnadurai et al, 2014). However, reducing the number of oysters per batch in the depuration system also increases cost which may not be commercially acceptable.
41 The effect of temperature and water salinity on the depuration process has also been studied. High water salinity (35 psu) enhanced the elimination of two pathogenic bacteria, Vibrio vulnificus (Vv) and Vibrio parahaemolyticus (Vp), from oyster tissues after 3 days of depuration. In contrast with other studies, the results showed that bacterial elimination was significantly higher at a high temperature compared to low temperature.
The study concluded that using high salinity water enhanced bacterial elimination.
However, the level of salinity may influence the optimal depuration temperature which should be considered when designing a depuration process (Larsen et al., 2015).
Although commercial depuration and relaying procedures are common approaches to remove pathogenic bacteria such as E. coli, it has some limitations. For instance, it has been shown that depuration at the optimal salinity and temperature can lead to an increase in Vibrio concentrations in shellfish tissues. It has also been demonstrated that depuration and relying are less effective in eliminating enteric viruses
(Schwab et al., 1998; Love et al., 2010; Chinnadurai et al, 2014; Garcia et al., 2015).
Histo-blood group antigens (HBGAs) the specific HuNoV receptor have been detected in oysters, clams, and mussels (Maalouf et al., 2010). Thus the limitation of the depuration process to remove enteric viruses, especially HuNoV, from shellfish may be due to non- specific binding and specific binding of HuNoVs to HBGAs present in shellfish tissues.
Moreover, the high cost of the depuration process limits its wide use commercially. Thus, it should be noted that focusing on new growing water quality standards that consider enteric viruses is the most effective strategy to reduce seafood related outbreaks.
42 1.11. Thermal processing:
Even though other processing methods such as high pressure processing (HPP) and food irradiation are widely accepted as a non-thermal technologies to eliminate microorganisms from food, classic thermal processing remains the most popular method to inactivate pathogenic and spoilage microorganisms by the food industry. That popularity has been supported by the long history of food safety in the canning industry.
In general, thermal processing is a combination of temperature and time to inactivate a certain amount of a target pathogen in the processed food. Based on the treatment temperature, time, and the target microorganism, thermal processing can be divided into pasteurization and commercial sterilization (retorting). Pasteurization mainly targets pathogenic viruses and bacteria (in vegetative state) by using low temperature followed by rapid cooling. The advantage of pasteurization is that it has a minimal effect on the organoleptic properties of the processed food and prevents the survival of spoilage microorganisms from growing during storage (Hasting, 1992). Disadvantages include the fact that pasteurization does not affect heat resistant viruses or bacterial spores. During pasteurization, the processing temperature and time depends on the type of food, target microorganism, and the local regulations. In general, the most common pasteurization standard temperatures and treatment durations are: 62-65˚, 72˚, and 80-95˚C for 30 min,
15-40 s, and 2-10 s, respectively (Hasting, 1992).
The principal of retorting is to reach a target sterilization level, a 12 log reduction of a target microorganism, using a high temperature, such as 121 °C, for a given time.
Retorting requires that all particles in the food matrix reach the desired temperature and time. Despite the reliability of commercial retorting in providing a long shelf life for the
43 food products, the undesirable effect of the severe thermal treatment on the flavor, color, and texture of the treated food is still unavoidable. Moreover, retorting requires a large amount of energy for steam generation and rapid cooling (Barbosa-Cánovasa et al.,
2014).
The study of the thermal inactivation of foodborne viruses is still under development due to difficulty to extract, cultivate, and quantify the most important foodborne viruses, such as HuNoV (Grove et al., 2006). Heat inactivation of viruses depends on several factors: 1) the treated virus (enveloped vs. non-enveloped viruses, virus strain, genogroup, and family); 2) the chemical and physical properties of the processed food such as the moisture, fat, salt, and protein content and pH, which can confer protection to the treated virus during the heat treatment; 3) the treatment parameters (temperature, time, type of the retort etc.). All these factors need to be considered when the thermal inactivation processes are designed.
1.11.1. The inactivation models of foodborne viruses:
It has been shown that virus inactivation by thermal processing does not follow a linear inactivation model (first-order kinetics) that assumes a linear logarithmic reduction of the treated virus with time. In a linear logarithmic model the slope of the inactivation curve can be used to calculate the decimal reduction value (D- value), the time required to reach one log reduction of the virus at a specific temperature (Chick, 1908; Tuladhar, et al., 2012). A virus thermal inactivation curve usually shows shouldering and tailing at the beginning and the end of the treatment curve, respectively, which limits the ability of using D-values to establish the inactivation parameters. Therefore, alternative models are
44 being applied to overcome the nonlinear inactivation curve issues. For example, the biphasic reduction model is used to calculate the time to the first log10 reduction (TFL- value), instead of calculating D-value, from the inactivation curve based on two different rates of inactivation (de Roda Husman, et al., 2009; Tuladhar, et al., 2012). The Weibull model was described by van Boekel (2002) and it has been shown as the best fit to express virus inactivation in many studies (van Boekel, 2002). In the Weibull model the inactivation parameters α and β and the scale and shape of the probability density function are used to describe the time required for a desired amount of inactivation at a specific temperature (Bozkurt et al., 2014). The application of these statistical models is helpful in estimating virus survival during thermal processing, however the application of these models requires comparable data on the stability of different viruses to build the model on and appropriately describe the inactivation curve (Tuladhar, et al., 2012).
1.11.2. The comparison of the thermal stability of several foodborne viruses:
The efficacy of thermal treatment of viruses is complex and depends on the target virus. In general, the systematic comparison of heat inactivation of viruses has showed that viruses of different families, genogroups, genotypes, and even strains behave differently under heat treatment. For example within the Calicivirus family, MNV-1 and
FCV, commonly used surrogates for HuNoV, were compared for thermal stability at pasteurization temperatures of 56°C, 63°C, and 72°C in hot bars, and the results showed that at 56°C MNV-1 was more sensitive to the heat treatment than FCV. One log reduction of MNV-1 and FCV required 3.5, and 6.7 min treatment time, respectively.
Increasing the treatment temperature to 63°C reduced the required time to achieve a 1-log reduction of both viruses in <25 seconds (Cannon et al., 2006). On the other hand,
45 Bozkurt, et al. (2013) calculated the D-values of MNV-1 and FCV at heat treatment temperatures ranging between 50 to 72°C. The study concluded that MNV-1 was more stable at 72°C than FCV. The D72 value of MNV-1 and FCV was 0.15 and 0.11 min, respectively (Bozkurt et al., 2013). The thermo-stability of TV was also compared to
MNV-1 at 70 and 75°C in culture medium (Hirneisen & Kniel 2013). The complete elimination (6-log reduction) of both viruses was achieved after a 2 min holding time at
70 and 75°C. Reducing the treatment temperature from 70 to 60 resulted in only 3-log reduction after a 2 min holding time (Hirneisen & Kniel, 2013).
Thermal stability studies of HuNoV are hampered because of the lack of suitable cell culture system and a small animal model to support HuNoV growth. However,
HuNoV receptor binding and RNase pre-treatment coupled with RT-qPCR is widely used to assess the survival HuNoV after heat treatment (Topping, et al., 2009, Li, et al., 2011).
Li et al. (2012) studied the stability of HuNoV at 70 and 85°C for 2 min. At 70°C, 1.62 log10 RNA copies reduction was achieved after a 2 min holding time, while at 85°C an average of 5 log10 RNA copies were eliminated after 2 min treatment (Li et al., 2012).
The results were supported using HuNoV VLPs in the binding assay after heat treatment.
For instance, Li et al., (2012) showed that HuNoV GII.4 VLPs completely lost its ability to bind to the HBAGs at 82°C after 2 min of holding time. The same study compared the heat stability of VLPs of different HuNoV strains, GI.1, GI.4, GII.4, and GII.9. The results indicated that GI.1 was the most sensitive strain and GII.9 was the most resistant to the heat treatment (Li, et al., 2012). The data may indicate that different viruses of the same genotypes, families, and even strains may have differing thermal stabilities.
46 Picornaviruses are highly thermal stabile. Tuladhar, et al. (2012) compared the stability of adenovirus type 5 with two picornaviruses, poliovirus Sabin 1 and parechovirus 1 under the heat treatment at 56 and 73°C in culture medium and a stool suspension. The time to first log (TFL) reduction values showed that parcehovirus 1 was highly resistant to heat treatment at 56°C, with TFL of 27 min in culture medium and no reduction in the stool suspension, compared to a TFL < 0.30 and 0.1min for both adenovirus type 5 and poliovirus Sabin 1 in culture medium and stool suspension, respectively (Tuladhar, et al., 2012). HAV, another food transmitted picornavirus, showed high heat resistance, in a survey conducted by Bidawid, et al. (2000). To investigate the effect of fat content of dairy products on the thermal stability of HAV, three different dairy products (skim milk with 0% fat, homogenized milk with 3.5% fat, and table cream with 18% fat) were inoculated with HAV at a level of 2.3 ×106 PFU/ml.
The inoculated products were heated in micro capillary tubes in a water bath at temperatures between 65-85˚C for intervals of holding times between 0.5-6 min. The results showed that at the highest treatment temperature (85°C), the fat content did not significantly affect virus inactivation. At this temperature a 5-log reduction was achieved
(the required log reduction by FDA). At low treatment temperature (<85°C), the fat content provided a protection to HAV during heat treatment. The virus was more stable in table cream compared to the skim milk and homogenized milk (Bidawid, et al., 2000).
Bozkurt, et al. (2014) performed an experiment to study HAV stability during the blanching of chopped spinach. The spinach samples were inoculated with 7-log10 PFU/ml of HAV and homogenized. The samples were heated at 50 -72°C for holding times between 0 and 6 min. The log reduction was calculated using plaque assay and the first-
47 order kinetic model was used to calculate the change in the virus titer. The results showed that the D-value of HAV at 50 °C was 34.40 ± 4.08 min. increasing the blanching temperature to 72°C reduced the D-value to 0.91 ± 012 min. The key findings are that
HAV is highly stable to heat treatment and the food matrix plays an important role in virus inactivation (Bozkurt et al., 2014).
To compare HAV to HuNoV surrogates, a study showed that HAV required longer exposure to heat than did MNV-1 for complete inactivation in soft-shell clam tissues, which has also been observed in studies done with other shellfish. For cockles, immersion in water at 90˚C for 180 seconds or 95 ˚C for 120 seconds was sufficient to inactivate HAV (Millard et al., 1987), whereas 108˚C for 120 seconds (2 minutes) was necessary for complete inactivation of this virus in mussel homogenates (Croci et al.,
1999). The treatment applied by Hewitt et al. (2009) in milk and water (72˚C for 60 seconds) was equally effective for HAV and MNV-1. A recent study by Mormann et al.
(2010) showed that complete inactivation of HuNoV GII could be achieved by boiling for
30 minutes at 100 ˚C in liquid with reduced water activity (mincemeat) but not by dry heating (oven-roasting) for 30 minutes at 200 ˚C (Mormann et al., 2010). It is therefore possible that an internal temperature of 90 ˚C for 180 seconds might be sufficient for inactivation of HuNoVin soft-shell clams or other food matrices. Bivalve shellfish are offered in the market in either shucked (i.e., with the meats removed from the shell) or the whole-shell form. For inactivation of HAV in New Zealand Greenshell whole mussels, immersion in boiling water for 260 seconds (170 seconds to achieve 90 ˚C and
90 seconds maintained at this temperature) has been suggested.
48 1.12. High hydrostatic pressure (HPP):
A non-thermal process is a “cold” process, which can be used for decontamination, pasteurization, and sterilization. One of the key attributes of the non- thermal processed product is the excellent quality, wherein the products maintain a
“fresh” characteristic. The advantages include better nutritional values (e.g. vitamin, enzyme, and protein), better sensory (e.g. texture and color) and microbiological quality, and minimal or no use of preservatives (Grove et al., 2006; Rastogi, Raghavarao et al.,
2007). Commonly known non-thermal technologies include high pressure processing
(HPP), irradiation, ultraviolet (UV) light, ultrasound, pulsed electric field, and cold plasma. The application of a particular non-thermal processing technology depends on the type of targeted matrix, the location of the pathogens, and the effectiveness of the technology. For viruses located within the food matrix, technologies targeting food surface areas (such as UV, ultrasound, and cold plasma) are not applicable. Some technologies (such as pulsed electric field) are only suitable for liquid matrices. Ionizing radiation (gamma rays and electron beam) has the ability to penetrate the food matrix and therefore could be applicable to target internalized viral pathogens. High pressure processing (HPP) evenly distributes pressure throughout the food being treatmed and therefore may also be effective to eliminate internalized viruses.
With the increasing demand for minimally processed food, there is an urgent need for an effective non-thermal processing technology which can eliminate pathogens while retaining the organoleptic properties of the processed food. High pressure processing
(HPP) or High Hydrostatic Pressure processing (HHP) is one the most promising approaches to achieve this goal. HPP is able to inactivate or reduce foodborne pathogens
49 including bacteria, molds, and viruses by applying the high hydrostatic pressure evenly from all sides of the processed food without additional heat. Studies showed that microorganisms respond differently to HPP. For example, gram-negative bacteria are more susceptible to HPP than gram-positive bacteria. As for viruses, it is reported that there is no correlation between virus barosensitivity and the presence of a viral envelope
(Lou et al, 2011b). The enteric viruses, non-enveloped viruses, also have shown heterogeneous responses to HPP. For instance, HuNoV surrogates such as FCV, TV, and
MNV-1 showed different susceptibilities to HPP (Kingsley et al., 2002; Kingsley et al.,
2007; Sánchez et al., 2011).
In 2002, Kingsley et al. preformed an experiment to inactivate HAV and poliovirus in cell culture medium by HPP. HAV is one the most resistant viruses in response to heat treatment while poliovirus is one of the most sensitive. Treatment pressure levels ranged between 300 and 600 MPa (Mega Pascal is the unit used to measure the pressure, and one MPa= 145 pounds per square inch) for 5 min. The results showed a 7-log reduction in HAV titer can be achieved at 450 MPa, whereas poliovirus had high stability to HPP treatment and remained viable after 600 MPa for 5 min. Adding salt to the culture medium enhanced HAV stability (Kingsley et al., 2002). In the same trend, Lou et al. (2011) studied the inactivation of MNV-1 by HPP in culture medium and fresh produce. The study investigated the effect of the treatment temperature (4 and
20⁰C), the pH (between 2.5 to 6.3), and the pressure levels (ranged between 200 to 450
MPa). The results showed that a treatment of 400 MPa for 2 min at 4°C was enough to completely eliminate MNV-1 from the fresh produce. HPP treatment at low temperature
50 (4°C) was more effective in the virus inactivation. MNV-1 was more susceptible to HPP at neutral pH (7) than the acid pH (Lou et al., 2011).
1.12.1. Mechanism of HPP inactivation of viruses:
Lou et al. (Lou et al., 2011) performed a systematic study to determine the mechanism of inactivation of norovirus by HPP using MNV-1 as a model. After pressure treatment, the virion structure changed from a discrete small round shape to a large amount of undefined protein debris as visualized by electron microscopy. The capsid proteins of MNV-1 were not degraded and were still capable of reacting with antibodies, indicating that the primary and secondary structures of viral proteins remained intact, although the quaternary and tertiary structures of viral capsid proteins were completely distorted (Lou et al., 2011). In a subsequent study by Lou et al. (2012), HPP also impaired the receptor binding capacity of HuNoV VLPs. This is also supported by another study showing a significant decrease in the ability of HuNoV to bind its functional receptor, the histo-blood group antigens (HBGAs), using the PGM-MB binding assay after treatment at 400 MPa at 5oC for 5 min (Dancho et al., 2012). It was also shown that HPP does not degrade MNV-1 genomic RNA (Lou et al., 2011).
Subsequent studies have shown that HPP treatment does not lead to lower levels of
HuNoV or TV RNA as detected by RT-qPCR although receptor binding ability or infectivity has been abolished (Cromeans et al., 2014; Li & Chen, 2015). This lack of
RNA degradation is due to the fact that HPP does not break covalent bonds at the level applied for food processing (Lou et al., 2011; Tang et al., 2010). Therefore, the primary
51 mechanism of norovirus inactivation is the disruption of the virion capsid structure and receptor binding activity but not degradation of viral proteins or RNA.
1.12.2. Intrinsic factors that influence virus inactivation by HPP
To effectively inactivate pathogens, it is critical to optimize the conditions for pressure treatment. The effectiveness of HPP is influenced by many factors such as processing parameters (applied pressure, holding time, and initial temperature) and non- processing parameters (virus structure itself, food matrix, pH, and aw of foods) (Chen, et al, 2005; Grove et al., 2006; Grove et al., 2008; Lou, et al., 2011a; Lou et al., 2011b).
Generally, the extent of virus inactivation increases commensurate with the pressure level and holding time. But it was discovered that virus inactivation is more pressure-dependent and a first-order relationship was exhibited between pressure levels and inactivation outcome (Dancho et al., 2012). As for treatment time, the HPP inactivation curves of FCV, MNV, and HAV showed variation of virus titer as a function of treatment time, exhibiting pronounced tailing, indicating that longer holding time did not significantly enhance inactivation (Chen et al., 2005; Kingsleyet al., 2006; Kingsley et al., 2007; Lou et al., 2011).
The initial temperature at which pressure is applied can also have a significant impact on pressure inactivation of viruses whereby temperature can either work synergistically or antagonistically with pressure. It is reported that pressure inactivation of MNV-1 was favored at refrigerated temperatures (Lou et al., 2011); a 350-MPa treatment held for 5 min at 30°C inactivated only 1.2 logs of virus, while the same treatment achieved a reduction of 5.6 logs at 5°C (Kingsley et al., 2007). This is
52 consistent with the temperature-sensitivity of another recently developed HuNoV surrogate, Tulane virus (TV), during HPP treatment (Li et al., 2013). In contrast, other studies have shown that certain viruses were more sensitive to HPP at room temperature than at lower temperatures. For HAV, pressure inactivation was enhanced as temperatures increased above 30°C compared to temperatures ranging from 5°C to 30°C
(Kingsley et al., 2006). FCV was found to be minimally affected by pressure at room temperature and inactivation was enhanced at either below or above 20°C (Kingsley &
Chen, 2008). These observations suggest that the optimal temperature for pressure inactivation is specific to the virus of interest.
The pH of the suspending medium or food is an important consideration in the pressure inactivation of viruses. HAV is known to be pH-stable at atmospheric pressure; however its inactivation is significantly enhanced in an acidic environment (pH 3) compared to a neural environment (pH 7) at 400 MPa (Kingsley & Chen, 2009). This synergistic effect of pH and HPP could be beneficial for processing selected acidic food products such as salsa and strawberry puree (Kingsley et al., 2005). On the other hand,
MNV-1, TV, FCV, and RV were all shown to be more easily inactivated at neutral pH compared to acidic pH (Chen et al., 2005; Kingsley & Chen, 2008; Lou et al., 2011b;
Lou, et al., 2011a). As surrogates for enteric HuNoV, MNV-1 and TV both exhibit high stability at low pH (Li et al., 2013; Lou et al., 2011). For MNV-1, an 8.1 log reduction of virus titer was achieved at 350 MPa for 2 min at pH 7.0, whereas only a 6.0-log virus reduction was achieved at pH 4.0 at the same pressure level and holding time (Lou et al.,
2011b). A treatment of 350 MPa for 2 min at 21°C resulted in a 3.8-log reduction of TV at neutral pH, but the same treatment only reduced TV by 2.4 logs at pH 4 (Li et al.,
53 2013). The titer of HRV was reduced by 3.4 logs following exposure to 250 MPa of pressure at pH 7.0, but only 1.2 logs at pH 4.0 (Lou et al., 2011a). The mechanism by which pH influences the pressure inactivation of different non-enveloped viruses is unknown. It is likely that the susceptibility of virus to pH is dependent on the nature of the viral capsid protein.
Finally, the composition of a medium or food matrix is another important factor that affects viral inactivation during pressure treatment. It has been reported that carbohydrates, fats, salts, proteins, ions, and other food constituents can protect viruses from inactivation (Baert et al., 2009; Balny et al., 2002; Gross and Jaenicke, 1994;
Kingsley and Chen, 2008, and 2009). For instance, sucrose (40%) provided a baroprotective effect against HPP inactivation of FCV and resulted in <1 log inactivation at 250 MPa at 20°C for 5 min (Kingsley and Chen, 2008). Murchie et al. (2007) pressure- treated several viruses including FCV in shellfish, seawater, and culture medium and the viruses were found to be most resistant when treated in oysters and mussels (Murchie,
2007). Comparisons of the effect of HAV inactivation in oyster homogenates and 0.3%
NaCl solutions at a similar pH revealed an increased pressure resistance in oyster homogenates compared to salt solutions, suggesting the baroprotective characteristic of other oyster components (Kingsley and Chen, 2009). Consistent results were observed in a study on MNV-1 demonstrating that viral reduction was higher in medium (8 logs) than in strawberries (5.8 logs) and in strawberry puree (4.7 logs) under 450 MPa for 2 min at
4°C (Lou et al., 2011b). These observations demonstrate that the food matrix confers protection from inactivation of viruses by HPP. Thus, it is necessary to optimize the
54 processing parameters for each product since the efficiency of viral inactivation varies with the food matrix.
1.12.3. Application of HPP in processing high-risk foods:
Currently, HPP is employed for the commercial processing of shellfish, oysters in particular. Not only does HPP significantly reduce pathogen levels in oysters, it also causes the oysters to be more voluminous, more flavorful, and more texture-attractive compared to untreated oysters (Cruz-Romero, 2004; He, 2002; Hoover, 1989; Lopez-
Caballero et al., 2000). Moreover, HPP treatment causes the adductor muscle of oysters detach from the shell thus opening (shucking) the oyster. This “self-shucking” aspect of
HPP can drastically cut down on labor requirements associated with manual shucking, thus constituting an important economic advantage for oyster processors (He, 2002;
Murchie, 2007).
Although Molluskan shellfish are recognized as a high risk food for viral contamination and have led to several high profile outbreaks, the bioaccumulation profiles for different viruses and different strains of viruses is unknown. In addition, the ability of both thermal and non-thermal processing to inactivate different viruses and different viral strains is poorly understood. In this research we will evaluate the distribution profile of different viruses including MNV-1, TV, HuNoV, RV, and HAV in oyster tissues to determine whether each virus has a unique distribution profile. In addition we will compare the inactivation of these different viruses in oyster tissues by heat and HPP to determine the optimal inactivation profile for each individual virus. To evaluate inactivation variation at the strain level, seven different RV strains derived from
55 four genotypes will be subjected to HPP treatment to determine if different viral strains are more resistant/sensitive to HPP treatment. In addition, it is well known that bacterial pathogens easily develop resistance under environmental stress. However, whether foodborne viruses can develop such resistance and mechanism involved in developing resistance are poorly understood. This research will determine whether calicivirus can develop resistance upon heat and HPP treatment. This research will determine optimal treatment parameters to inactivate viruses in high risk foods, specifically molusckan shellfish. These results can be used by the shellfish industry to significantly improve the safety of seafood, and thus limit outbreaks.
56
CHAPTER 2
Natural bioaccumulation of enteric foodborne viruses in live oysters and thermal
inactivation of enteric viruses
2.1. Abstract
The consumption of contaminated bivalve shellfish continues to pose a public health risk. Human enteric viruses have been reported as one of the main causative agents associated with bivalve mollusk outbreaks. The objective of this study is to determine the kinetics of bioaccumulation, and heat stability of the most predominant enteric viruses, specifically human norovirus and its surrogates (murine norovirus, MNV-1; and Tulane virus, TV), hepatitis A virus (HAV), and human rotavirus (RV) in aqueous medium and oyster tissues. Our results demonstrate that all viruses were bio-accumulated to a high titer within the oyster tissues, however, the pattern of the bioaccumulation substantially varied for each individual virus. Caliciviruses and HAV were localized in the stomach at a high level within the first 24 h, while RV was bio-accumulated in gills. The biphasic reduction model showed the best fit to describe the virus inactivation curves at 62 and 72
˚C in culture medium. Decimal reduction time (D-values) of the low and high thermal resistance fractions of the four viruses ranged from 0.13 to 1.81 min, and from 1.26 to
7.29s at 62 and 72˚C, respectively. The Weibull distribution was the best fit for the inactivation curves at 80 ˚C, and the first log10 reduction (TFL-value) ranged between
0.46 and 32 s. Within the oyster tissues, the TFL at 80 ˚C ranged between 0.61 to 19.99
57 min. The four viruses can be ranked from the most heat resistant to the least stable as following: HAV>RV>TV>MNV-1. At 80 ˚C, time requires for complete inactivation of
HAV, RV, TV, and MNV-1 in cell culture medium is 12, 10, 10, and 6 s respectively.
However, it requires 3, 3, and 4 min at 80˚C to inactivate MNV-1, TV, and RV, respectively in oyster tissues; HAV required more than 6 min to be activated in oysters.
To decipher the mechanism underlying viral inactivation by heat, purified TV was treated at 80°C for increasing time intervals. It was found that the integrity of viral capsid was disrupted whereas viral genomic RNA remains intact. However, a lethal dose of heat treatment was not sufficient to disrupt the receptor binding activity of HuNoV, virus-like particles, and TV. Collectively, we demonstrate that enteric viruses were efficiently bioaccumulated in oyster tissues and different viruses exhibited different distribution patterns. Although foodborne viruses have variable thermal stability to heat, 80oC for more than 6 min was sufficient to completely inactivate major enteric viruses in oysters.
2.2. Introduction
The consumption of contaminated bivalve shellfish continues to pose a public health risk. These bivalve filter-feeders sieve gallons of water a day through their gills which can lead to the bio-accumulation of both bacterial and viral pathogens within shellfish tissues. Human enteric viruses have been reported as one of the main causative agents associated with bivalve mollusk outbreaks. Human norovirus (HuNoV), hepatitis
A virus (HAV), and rotavirus (RV) account for the majority of these shellfish outbreaks
(Bellou et al., 2013; Boxman et al., 2006; Brake et al., 2014; Le Guyader et al., 2006;
58 Loury et al., 2015) . Pioneer studies have shown that these enteric viruses persist and can be bio-accumulated to high titers within shellfish tissues (Richards, 1988).
HuNoV causes severe gastroenteritis characterized by vomiting, diarrhea, and stomach cramps (Atmar et al., 2008). In the US, it is estimated that HuNoV accounts for more than 60% of foodborne illnesses and it is the second leading cause of gastroenteritis-related mortality, causing 797 deaths annually (Scallan et al., 2011, Hall et al., 2013). It has been a challenge to study HuNoV because it cannot be grown in a cell culture system and lacks a small animal model. Therefore, cultivable viral surrogates that are closely related to HuNoV, such as murine norovirus (MNV) and Tulane virus (TV) have been used to study the survival of HuNoVs in foods and the environment (Bozkurt et al., 2013; Bozkurt et al., 2014; Drouaz et al., 2015; Li et al., 2013; Predmore et al.,
2015). HAV causes gastroenteritis, liver damage, and jaundice. HAV outbreaks have declined due to improvements in drinking water quality and sanitation practices, as well as the availability of effective vaccines in developed countries. However, HAV remains endemic in developing countries (Jacobsen and Koopman, 2004). Thus the presence of
HAV in imported seafood remains an important concern for US regulatory agencies.
Rotaviruses (RVs) are the major etiological agent of acute gastroenteritis in infants worldwide, and account for 27% of deaths in children, according to the WHO (Lanata et al., 2013). RV is highly infectious among children due to its low infectious dose (<10 particles can lead to disease) and the fact that the virus is shed at a very high titer (1010 to
1012 virus particles/g of stool) (Ward et al., 1986; Bishop, 1996). All three of these viruses are highly stable in foods, water, and the environment. In addition, all three of
59 these viruses are transmitted mainly via the fecal-oral route, so the consumption of contaminated food or water sources often leads to disease.
The current standards used to monitor the safety of shellfish and the quality of growing waters depends on the levels of E. coli or total fecal coliforms (Lee & Reese,
2014). These standards (Marino et al., 2005) have succeeded in reducing bacterial shellfish related outbreaks, however it has been shown that the bacterial standards are inadequate for estimating the presence of the enteric viruses in shellfish and growing waters (Dowell, et al., 1995). Moreover, the depuration process for shellfish harvested from conditionally approved (or category B) growing areas has been shown to be less effective in eliminating enteric viruses from contaminated shellfish (de Abreu Correa et al., 2012;Dore et al., 2000; Garcia et al., 2015). Therefore, there is a need to understand the interaction of viruses and shellfish and also to develop efficient methods to inactivate/remove the viruses from this food commodity.
Previous work has evaluated the distribution of bacterial pathogens in shellfish, however little is known about the bioaccumulation and distribution of viral pathogens
(Marino et al., 2005;Powellet al., 2013; Wang & Shi, 2011). In general, viral localization in shellfish has been shown to vary based on the exposure time and the virus type. It has been demonstrated that the type of shellfish and also the season affect the rate of viral bioaccumulation (Maalouf et al., 2010; Grodzki et al., 2014). Previous work has demonstrated that a primate calicivirus, Tulane virus, can be bioaccumulated in shellfish and can serve as a potential surrogate to mimic norovirus behavior in oysters (Drouaz et al., 2015). However, no study has directly compared the bioaccumulation profiles of the three most prevalent viruses (HuNoV, HAV, RV) causing shellfish outbreaks directly.
60 Thermal treatment is still regarded as one of the most effective means to inactivate spoilage and pathogenic microorganisms. Most thermal inactivation methods for shellfish have been standardized to target bacterial species. However, these inactivation parameters (e.g., time, temperature) may not be sufficient to inactivate foodborne viruses. Therefore, there is an urgent need to identify effective thermal processing parameters to inactivate foodborne viruses in shellfish (Bertrand et al. 2012).
To date, there is no standard method that has been established to study the thermal stability of enteric viruses (Arthur & Gibson, 2015). Treatment conditions such as temperature, holding time, sample matrix, and data modeling method, as well as virus type tested were found to influence the kinetics of thermal inactivation and differ between research studies (Arthur & Gibson, 2015).
Additionally, it has been shown that virus inactivation by thermal processing does not follow a linear inactivation model (first-order kinetics) that assumes a linear logarithmic reduction of the treated virus with time (Bozkurt et al., 2015). A virus thermal inactivation curve usually shows shouldering and tailing at the beginning and the end of the survival curve, respectively, which limits the ability of using D-values to establish the inactivation parameters. Therefore, alternative models, such as the biphasic reduction model and Weibull model, are being applied to overcome the nonlinear inactivation curve issues (de Roda Husman, et al., 2009; Tuladhar et al., 2012). Although the application of these models is helpful in estimating virus survival during thermal processing, the application requires comparable data on the stability of different viruses to build the model on and appropriately describe the inactivation kinetics (Tuladhar, et al., 2012).
61 The aim of this study is to obtain comprehensive data on the localization, bioaccumulation, and heat stability of the most predominant enteric viruses, specifically
HuNoV surrogates (MNV-1 and TV), HAV, and RV in oyster tissues. Comparable data on the thermal stability of different enteric viruses will provide insight into the kinetics of inactivation and aid in establishing new standards that are affective against all enteric viruses.
2.3. Materials and methods
2.3.1. Cell culture and virus propagation. MNV-1, a generous gift from Herbert W.
Virgin IV, Washington University School of Medicine (Karst et al., 2003), was propagated in confluent flasks of murine macrophage cell line RAW264.7 (ATCC,
Manassas, VA) grown in high glucose Dulbecco's Modified Eagles Medium (DMEM) supplemented with 10% of fetal bovine serum (FBS) (Invitrogen), as described by (Lou et al., 2011). TV was provided by Xi Jiang at Children's Hospital Medical Center,
Cincinnati, Ohio, and propagated in monkey kidney cells (MK2-LLC) which was cultured in Opti-MEM supplemented with L-glutamine and 2% of FBS as described by
(DiCaprio et al., 2012). HAV, a generously gift from David H. Kingsley, Delaware State
University, Dover, Delaware, was cultivated in fetal rhesus monkey kidney cells (FRhK-
4). FRhK-4 cells was grown in Eagle's minimal essential medium (EMEM) containing
10% of FBS. Human RV (Wa strain, provided by John Hughes, The Ohio State
University Medical Center) was propagated in rhesus monkey kidney cells (MA-104)
62 cultured in Eagle's minimum essential medium (MEM) supplemented with 6µl/ml trypsin.
2.3.2. Viral plaque assays. MNV-1, TV, HAV, and RV plaque assay were performed in six-well plates containing RAW 264.7, LLC-MK2, FRHK-4, or MA104 cells, respectively, as described by Lou et al. (2011). A monolayer of 2 ×106 cells/well in 6- well plates (Corning Life Sciences, Wilkes-Barre, PA) were infected with 400 μl of a series 10-fold dilutions of the virus and incubated at 37°C and 5% CO2 atmosphere for 1 h for MNV-1, TV, and RV, and for 90 min for HAV with hand agitation every 10 to 15 min. An overlay solution containing 2× Eagle minimum essential medium (MEM), 1% agarose, 2% FBS, 1% sodium bicarbonate, 0.1 mg of kanamycin/ml, 0.05 mg of gentamicin/ml, 15 mM HEPES (pH 7.7), and 2 mM L-glutamine (Invitrogen) were added to each well for MNV-1, TV, and HAV and incubated for 48 h for MNV-1 and TV, and for 9 days for HAV plates. For the RV plates, the overlay solution contained 2.5 µl/ml trypsin and no FBS was added to the overlay and plates were incubated for 72 h.
Following incubation, the cell monolayer was fixed by 10% formaldehyde and stained with 0.05% crystal violet to visualize the viral plaques. Viral titer was expressed as mean plaque forming unit (PFU)/ml ± 1 standard deviation.
2.3.3. Virus bio-accumulation and distribution in oysters. Live oysters (Grassostrea gigas) were obtained from a local grocery store (Kroger Inc.) and were cultivated in a tank containing 4 liters of artificial sea water (1.5% sea salt) under aeration conditions.
Phytoplankton was added to the tank to feed the growing oysters. Prior to the
63 experiments, three oysters from each batch were shucked and homogenized and examined for the presence of enteric viruses including HAV and RV. Five batches of oysters of 25 oysters per batch were grown separately as described above, and the salt water was artificially contaminated with either TV, MNV-1, HAV, or RV at a virus level of ~ 1× 104 PFU /ml or ~ 1× 104 RNA copies/ml of HuNoV GII.4. The oysters were grown for 72 h in the contaminated tank. At 24, 48, and 72 h, three oysters from each tank were aseptically dissected. Different portions of the oyster tissue including gills, digestive gland, and muscles were isolated and examined for the presence of virus by plaque assay or RT-qPCR.
2.3.4. Plaque assay and virus extraction from contaminated oyster tissues. TV,
MNV-1, HAV, and RV virus extraction and plaque assay were conducted as described by
Kingsley and Richards (2003) with some modifications. In brief, three contaminated oysters after 24, 48, or 72 h were harvested and shucked and each targeted tissue (gills, digestive gland, and muscles) was aseptically separated in a weight boat. Two grams of each separated tissue was homogenized in 5 ml of Hank's Balanced Salt Solution (HBSS) and the homogenized tissues were centrifuged at 2,300 × g for 10 min at 4°C. The supernatant was collected and ten-fold serial dilutions were made in the suitable medium for each virus. The plaque assay was conducted as described previously.
2.3.5. Thermal treatment of viruses in culture medium. To minimize the effect of the heating vessel on the temperature come-up time, capillary tubes were used to conduct the heat treatment. Eighty microliters of viral suspension of HAV, RV (Wa), MNV-1, or TV
64 in culture medium was inserted in capillary tubes (Kimble Chase, 1.5-1.8 × 100 mm) and sealed with a vinyl plastic cover (Leica Critoseal). The sealed tubes were heated in a circulating thermostatically controlled water bath at treatment temperatures of (62, 72, or
80°C) for different treatment times (ranging from 2 s to 30 min). The water bath temperature was monitored using a mercury-in-glass thermometer. The come-up times at each temperature was determined using a thermocouple probe inserted inside a capillary tube filled with culture medium. The come-up time was less than one second at all treatment temperatures, therefore it was ignored in the calculations of Decimal reduction time (D-values) or TFL. Following the heat treatment, the capillary tubes were immediately cooled on ice. The treated viruses were transferred to sterilized Eppendorf tubes and 50 µl of each treated virus was used for plaque assay.
2.3.6. Heat inactivation of MNV-1, TV, HAV, and RV in oyster tissues. Before the experiment, oysters were adapted to grow in the salt water for 24 h and dead oysters were removed before the experiment. For the contaminated oyster experiment, 25 oysters were cultivated in four liters of artificially contaminated salt water (1.5% sea salts and a phytoplankton) containing 1 × 105 PFU /ml of MNV-1, TV, HAV, or RV for 24 h. The water was aerated and kept at room temperature. The contaminated oysters were harvested and three oysters were randomly assigned for each heat treatment time point.
Contaminated oysters were treated in an 80˚C water bath for treatment times ranging from 1 to 6 min. The water bath temperature was monitored using a mercury-in-glass thermometer. The heat treated oysters were aseptically shucked, and the oyster tissues were homogenized in 5ml of HBSS. The virus survival was determined by standard
65 plaque assay. The thermal inactivation kinetics for each virus in oyster tissues was determined and the TFL for each virus was calculated as described above.
2.3.7. Isolation and characterization of HuNoV GII.4 strain 5M. Human NoV clinical isolate 5M was originally isolated from an outbreak of acute gastroenteritis in Ohio. The stool samples were diluted 1:10 in PBS, shaken vigorously for 10 min at 4ºC and centrifuged for 10 min at 5,000×g. The sample was filtered through a 0.45 μm filter, aliquoted and stored at −80ºC until use. The entire genomic cDNA of the human NoV strain 5M was amplified by RT-PCR using five to six overlapping fragments. The PCR products were then purified and cloned into a pGEM-T-easy vector (Promega), and sequenced at the Plant Microbe Genetics Facility at The Ohio State University. The full- length genome of the viral isolate was 7558 nt in length and has been deposited into
GeneBank at accession number JQ798158. Sequence comparison found that the strain belongs to the norovirus genotype GII.4. The genomic RNA was then quantified by real- time RT-PCR and the GII.4 isolate 5M was found to have 6.7×106 genomic RNA copies/ml.
2.3.8. Production of HuNoV GII.4 virus-like particles (VLPs). The capsid VP1 gene of a human NoV GII.4 strain (HS66) was amplified by high fidelity PCR and cloned into a pFast-Bac-Dual expression vector (Invitrogen) at Sma I and Xho I sites under the control of the p10 promoter, which resulted in construction of the expression vector, pFast-Bac-
Dual-VP1. The correct insertion of the VP1 gene was confirmed by DNA sequencing.
Subsequently, pFast-Bac-Dual-VP1 was transformed into DH10Bac and the baculovirus
66 expressing VP1 protein was generated by transfection of bacmids into Spodoptera frugiperda (Sf9) cells (ATCC no. CRL-1711™, Manassas, VA) using a Cell-fectin
Transfection kit (Invitrogen), according to the manufacturer’s instructions. Human NoV
VLPs were purified from insect cells as previously described with minor modifications
(Ma & Li, 2011). Briefly, Sf9 cells were infected with baculovirus at a MOI of 10, and the infected Sf9 cells and cell culture supernatants were harvested at 6 days post-inoculation.
The VLPs were purified from cell culture supernatants and cell lysates by ultracentrifugation through a 40% (w/v) sucrose cushion, followed by CsCl isoponic gradient (0.39 g/cm3) ultracentrifugation. Purified VLPs were analyzed by SDS-PAGE,
Western blot, and electron microscopy. The protein concentration of the VLPs was determined using Bradford reagent (Sigma Chemical Co., St. Louis, MO).
2.3.9. Preparation of porcine gastric mucin conjugated to magnetic beads (PGM-
MBs). PGM-MBs were prepared as described previously (Li et al., 2013; Dancho et al.,
2012; Tian et al., 2008). In short, 1 ml of MagnaBind carboxyl-derivatized beads
(Thermo Scientific, Rockford, IL) were washed three times with 1 ml of PBS on a bead attractor (EMD Millipore), to separate the beads. Porcine gastric mucin (PGM) solution of type III mucin was prepared by dissolving 10 milligram of type III mucin from porcine stomach (Sigma, St. Louis, MO) in 1 ml of conjugation buffer (0.1 M MES (2-(N- morpholino) ethanesulfonic acid), 0.9% NaCl, pH 4.7). The suspension was shaken well, and 0.1 ml of 10 mg/ml 1-ethyl-3-[3-dimethylaminopropyl]carbodiimide hydrochloride
(EDC) in the same conjugation buffer was add to the type III mucin solution. The mixture was added to the washed beads and shaken for 30 min at the room temperature on a
67 bunch top rotor. After rotation for 30 min, the beads were washed three times with 1 ml of PBS, and re-suspended in 1 ml PBS containing 0.05% sodium azide. The beads were stored in the refrigerator until use.
2.3.10. PGM-MB binding assay. Porcine gastric mucin beads assay followed by real time PCR was used to discriminate between infectious and non-infectious particles of
HuNoV. The PGM-MB binding assay was executed as described by Dancho et al.
(2012). Fifty microliter of heat treated or untreated TV, HuNoV GII.4, or HuNoV GII.4
VLPs were added to 100 µl of PGM-MBs, and the volume was brought up to 1 ml by adding 850 µl of PBS in low adhesive 1.5 ml centrifuge tube. The mixture was shaken for
30 min at room temperature. After incubation the beads were separated from the mixture by the magnetic bead attractor. The beads were washed three times by 1 ml of PBS. The beads were re-suspended in 100 μl PBS and used for RNA extraction for virus detection by RT-qPCR or loaded to SDS-PAGE for VLP analysis.
2.3.11. Quantification of viral RNA by real-time RT-PCR. Total RNA was extracted from HuNoV GII.4 and TV either untreated or heat treated at 62, 72, or 80˚C for holding time ranging between 2 s to 36 min using the RNeasy kit (Qiagen). The first strand of cDNA of the VP1 gene of HuNoV GII.4 or TV was synthesized with SuperScript III reverse transcriptase (Invitrogen) in RT-PCR. Two different primers (5’-
TTATAATACACGTCTGCGCCC-3’) and (5’-AATTCCACCTTCAACCCAAGTG-3’) were used to amplify the targeted VP1 of HuNoV GII.4 and TV, respectively. The first strand cDNA of HuNoV GII.4 or TV was quantified by real-time PCR using custom
68 TaqMan primers and probes (Forward primer, 5’-CACCGCCGGGAAAATCA-3’, reverse primer, 5’-GCCTTCAGTTGGGAAATTTGG-3’, and reporter 5’-
ATTTGCAGCAGTCCC-NFQ-3=) for HuNoV; and (forward primer, 5’-
TTGCAGGAGGGTTTCAAGATG-3’, reverse primer, 5’-
CACGGTTTCATTGTCCCCATA-3’, and probe, 5’-FAM-
TGATGCACACATGTGGGA-NFQ-3’) for TV on a StepOne real-time PCR machine
(Applied Biosystems, Foster City, CA). PCR and cycling parameters was according to the manufacturer’s protocol (Invitrogen). Cycling parameters were as the following holding stage at 95°C for 20 s prior to cycling, followed by 40 cycles of 95°C for 1 s for annealing and 60°C for 20 s for extension. Standard curves were used to convert CT values into log10 RNA copies. The RNA copies are expressed as genomic RNA copies/ml. Error bars represent mean of three replicates ± 1 standard deviation.
2.3.12. Thermal treatment of HuNoV GII.4 VLPs. To investigate the effect of heat treatment on the ability of HuNoV to bind to PGM-MBs, 20 µl of VLPs of HuNoV GII.4 were treated either at 80˚C for 10, 30, 60 s, and 5 min, or at 100 ˚C for 5 s in a capillary tube sealed with a vinyl plastic cover (Leica Critoseal) followed by rapid cooling on ice.
Treated or untreated VLPs were bound to the beads for 30 min at room temperature. The beads were washed on the beads attractor three times with 1 ml of PBS and re-suspended in 10 µl of PBS. Treated and untreated VLPs were mixed (1:4 V/V) with SDS-PAGE loading buffer which consists of 1% SDS, 2.5% β-mercaptoethanol, 6.25 mM Tris-HCl
(pH 6.8), and 5% glycerol. The mixture of each sample was boiled for 5 min and loaded
69 to a 15% polyacrylamide gel. The protein bands were visualized by Coomassie blue staining.
2.3.13. Purification of TV. Purification of TV was performed using the protocol described previously by Lou et al., (2011a). Approximately 180 ml of TV stock (1.5 ×
107 PFU/ml) was centrifuged at 82,000 × g through a 40% (wt/vol) sucrose cushion at
4°C for 3h, in a Ty50.2 rotor (Beckman Coulter, Fullerton, CA). The virus pellet was re- suspended in 300 µl of TNC buffer (100 mM NaCl, 10 mM Tris, 1 mM) on ice overnight.
A CsCl isoponic gradient (1.37 g/ml) was used to purify TV in a SW50.1 rotor (Beckman
Coulter) centrifuging at 115,000 × g at 4°C for 18 h. The TV band was collected (close to
1.73 g/ml density) and re-suspended in TNC buffer (0.05 M Tris-HCl, 0.15 M NaCl, 15 mM CaCl2, pH 6.5). The final TV pellets were re-suspended in 300 μl of TNC buffer.
2.3.14. Transmission electron microscopy. 10 µl of purified TV were heated at 80˚C for 5, 30 s, and 5 and 10 min. The samples were then fixed to copper grids (Electron
Microscopy Sciences, Inc.) and subjected to negative staining using uranyl acetate. Virus particles were then visualized by a FEI Tecnai G2 Spirit Transmission Microscope at the
Microscopy and Imagining Facility at the Ohio State University.
2.3.15. Reverse transcriptase polymerase chain reaction (RT-PCR). Reverse transcription polymerase chain reaction (RT-PCR) and RNase treatment were combined to determine whether the viral capsid of TV was degraded by heat at 80˚C for 5, 30s, and
5 and 10 min. Eighty microliters of TV was heated at 80˚C for 5 and 30s, and 5 and 10
70 min in capillary tubes and followed by RNase treatment of 10 μl (0.5 μg/μl) of RNase
(Invitrogen) for 30 min at 37 ˚C. Total viral RNA was extracted from TV heat treated and untreated virus using the RNeasy Mini Kit (Qiagen, Valencia, CA). Two primers were used to amplify the VP1 gene (5′-TTATAATACACGTCTGCGCCC-3′ and 5’-
GCCAGCCATTATCTAAAGA-3’). Bands were visualized using 1% gel electrophoresis.
2.3.16. Statistical analysis. All experiments were performed in triplicate. Virus titer was expressed as mean log PFU/ml ± 1 standard deviation. Statistical analysis was performed by one-way multiple comparisons using SPSS 8.0 statistical analysis software (SPSS Inc.,
Chicago, IL). A P value of <0.05 was considered statistically significant.
2.4. Results:
2.4.1. Virus bio-accumulation and distribution in oysters. To monitor the uptake and localization of enteric viruses within the oyster tissues, three caliciviruses including
HuNoV GII.4, and two of its surrogates (MNV-1 and TV), HAV, and RV (Wa strain) were compared for magnitude of uptake and localization within different oyster tissues.
Briefly, live oysters (Grassostrea gigas) were cultivated in a tank containing 4 liters of synthetic seawater (1.5% sea salt) and the salt water was artificially contaminated with
TV, MNV-1, HAV, or RV at virus level of 1× 104 PFU/ml of MNV-1, TV, HAV, or RV or 1 × 104 RNA copies of HuNoV. The oysters were grown for 72 h in the contaminated tank. Three oysters from each tank were aseptically dissected. Different portions of oyster tissue including gills, digestive gland, and muscles were isolated and examined for the
71 presence of the examined viruses. The observed accumulation pattern of the three caliciviruses (HuNoV GII.4 strain, MNV-1, and TV) is presented in figure 7.
Figure 7. Bioaccumulation of Caliciviruses HuNoV, MNV-1 and TV in oyster
tissues. (A) Bioaccumulation of MNV-1 in oyster gills, stomach, and muscles after 24,
48, and 72 h. (B) Bioaccumulation of TV in oyster gills, stomach, and muscles after 24,
48, and 72 h. MNV-1 and TV titer was assessed by plaque assay. (C) Bioaccumulation of
HuNoV GII.4 in oyster gills, stomach, and muscles after 24, 48, and 72 h. HuNoV titer
was assessed by RT-qPCR. The titer is mean of three replicates and error bars is ± one
standard deviation. Statistically significant (P< 0.05)
Continued
72 Figure 7 continued.
In general, HuNoV GII.4, MNV-1, and TV were bio-accumulated and detected very efficiently in the all oyster tissues through the three day experimental period. At 24
73 h, HuNoV GII.4, TV, and MNV-1 were accumulated at a significantly higher level
(p<0.05) in the digestive gland compared to the gills and muscles. The virus titer in the digestive gland ranged between 4.3 and 4.6 log10 PFU/g for MNV-1 and TV, respectively, and 5.0 genomic RNA copies/g for HuNoV. In the gills and muscles the titer of MNV-1 and TV ranged between 3 and 4.5 logs PFU/g, and 4.3 and 3.8 RNA copies/g for HuNoV. At the 48 and 72 h time points, HuNoV GII.4, TV, and MNV-1 were distributed equally in the oyster tissues with no significant difference in the level of the viruses between the different oyster tissues (P>0.05).
The results for HAV bioaccumulation in oysters is presented in Fig.8a. HAV was detected at a high titer in all oyster tissues after 24 h, with the highest level detected in the digestive gland. The virus titer in stomach reached 4.7 log10 PFU/g compared to 3.7 and 3.6 log10 PFU/g in gills and muscles, respectively. After 48 and 72 h, HAV was detected at similar levels in all oyster tissues, with no significant differences in HAV titer between different oyster tissues (P>0.05). Compared to TV, MNV-1, and HAV, RV
(Fig.8b) was accumulated at a relatively low titer and showed a different distribution pattern. After 24 h, RV was detected in the gills at a level of 3 log10 PFU/g and in the muscles at a titer of 2.3 log10 PFU/g. At the 24 h time point no RV was detected in the digestive gland. At 48 h, the RV titer increased in the digestive gland, however the titer detected was less than 1 log10 PFU/g. After 72 h, the level of RV detected in the gills and muscles was 2.5 and 3.5 log10 PFU/g, respectively, and the level of RV detected in the digestive gland increased to 1.7 log10 PFU/g.
74
Figure 8. Bioaccumulation of HAV and RV in oyster tissues: (A) bioaccumulation of
HAV in oyster gills, stomach, and muscles after 24, 48, and 72 h. (B) bioaccumulation of
RV in oyster gills, stomach, and muscles after 24, 48, and 72 h. Virus titer was assessed
by plaque assay. The titer is mean of three replicates and error bars is ± one standard
deviation. Statistically significant (P< 0.05)
75 2.4.2. Thermal inactivation of viruses in cell culture medium. To provide more precise data about the thermal stability of enteric viruses, thermal inactivation of HAV, RV, TV, and MNV-1 were systematically investigated at the pasteurization temperatures of 62, 72, and 80˚C in culture medium. The viruses were heated in capillary tubes in a water bath for holding times ranging between 2 s to 30 min. The virus survival following heat treatment was determined by plaque assay. The log10 PFU/ml of survived virus was plotted against the treatment time at each temperature as shown in figure 9.
Figure 9. Effect of heat treatment at 62, 72, and 80˚C on MNV-1, TV, HAV, and RV
(Wa strain) in culture medium: (A) The effect of heat treatment at 62˚C on the four
treated viruses. (B) The effect of heat treatment at 72˚C on the four treated viruses. The data presents the Log10 PFU/ml of the surviving viruses determined by plaque assay. (C)
The effect of heat treatment at 80˚C on the four treated viruses. Data are the means of
three replicates. Error bars is ± one standard deviation. Continued
76 Figure 9 continued.
77 The log linear, Weibull distribution, and biphasic models were compared to describe survival curve kinetics. The regression coefficients (R2) and root mean square error (RMSE) values were used to evaluate the model fitness. The D-values or the first log10 reduction (TFL-value) were calculated from the best fit model.
The inactivation data at 62˚C is presented in Table 5, based on the shape factor
(P) of the Weibull distribution (P<1) the four viruses showed monotonic upward concave
(tailing) behavior, which may indicate that the thermal sensitive members of the viral quasi-species were quickly inactivated while the more resistant quasi-species survived the mild treatment.
Table 5. D-value and TFL for HAV, RV, TV, MNV-1 after heat treatment at
62˚C in culture medium
62˚C Log liner Weibull Biphasic D 72 (min) TFL (min) D value1 (min) D value2 (min) HAV 5.46 0.09 0.13 7.58 RV 5.43 0.25 0.36 2.81 TV 2.38 0.49 0.48 2.07 MNV-1 2.17 1.53 0.55 1.81
The log linear model showed the worst fit of the experimental data with the lowest R2 and the highest RMSE. At this temperature, the D62˚C values, as calculated from the biphasic model, for the low and highly resistant quasi-species of the four viruses were 0.13, 7.58; 0.36, 2.81; 0.48, 2.07; and 0.55, 1.81 min for HAV, RV, TV, and MNV-
1, respectively. The biphasic model showed the best fit to the experimental data for the
78 four viruses at this treatment temperature, which can be concluded from the highest R2 and the lowest RMSE values of the biphasic model compared to log linear and Weibull distribution (Table.6).
By increasing the treatment temperature to 72˚C the inactivation curve gained more linearity and the tailing effect became less apparent, as shown by the increase of the
P value in the Weibull model (Table 6). The biphasic model showed the best curve fitness. However, the log linear regression had the lowest R2 and RMSS values among the three models used and it showed some improvement in the curve fitness. The D72˚C values for the low and highly resistant quasi-species of the four viruses were calculated from the biphasic model and were 1.26, 13.38; 1.87, 11.83; 1.46, 9.73; and 1.13, 7.29 s for HAV, RV, TV, and MNV-1, respectively (Table 7) .
79 Table 6. Comparing Log liner, Weibull distribution, biphasic models to best fit to the survival curve of TV, MNV-1, HAV, and RV at 62, 72, and 80˚C in culture
medium
62˚C Log liner Weibull Biphasic
RMSE R2 RMSE R2 P RMSE R2
HVA 0.74 0.583 0.18 0.980 0.21 0.09 0.996
RV 0.96 0.459 0.43 0.910 0.23 0.17 0.988
TV 0.87 0.844 0.42 0.969 0.44 0.38 0.980
MNV-1 0.92 0.851 0.65 0.939 0.59 0.38 0.983
72˚C Log liner Weibull Biphasic
RMSE R2 RMSE R2 P RMSE R2
HVA 0.91 0.734 0.54 0.923 0.51 0.26 0.986
RV 0.65 0.898 0.22 0.987 0.50 0.05 0.999
TV 0.90 0.861 0.62 0.943 0.58 0.46 0.979
MNV1 1.14 0.791 0.64 0.951 0.39 0.25 0.995
80˚C Log liner Weibull Biphasic
RMSE R2 RMSE R2 P RMSE R2
HVA 0.95 0.865 0.83 0.910 0.57 0.81 0.945
RV 0.72 0.925 0.634 0.958 0.67 0.68 0.960
TV 0.85 0.892 0.78 0.937 0.59 ND ND
MNV-1 0.60 0.964 0 1 0.52 ND ND
80 Table 7. D-values and TFL-values of HAV, RV, TV, and MNV-1 at 72˚C in culture medium
72˚C Log liner Weibull Biphasic D value (s) TFL (s) D value1 (s) D value2 (s) HAV 7.57 3.27 1.26 13.85 RV 5.3 3.11 1.87 11.83 TV 4.58 2.71 1.46 9.73 MNV-1 4.63 0.72 1.13 7.29
At 80˚C, the application of the biphasic model was not suitable for MNV-1 and
TV, because it requires more data points to be applied. The results showed that the
Weibull distribution was the most appropriate for modeling virus inactivation compared to the log linear model. In general, within 12 s all of the viruses were completely inactivated at 80˚C. TFL-values, as calculated from the Weibull model, were as the following: 0.46, 0.72, 0.32, 0.32 seconds for HAV, RV, TV, and MNV-1, respectively.
Based on the D-value or TFL-values for the four viruses they can be ranked from the most heat resistant to the least stable as: HAV>RV>TV>MNV-1.
Table 8. D-values and TFL-values of HAV, RV, TV, and MNV-1 at 80˚C in culture
medium
80˚C Log liner Weibull Biphasic D value (s) D 72 (s) TFL (s) D value1 (s) HAV 0.89 0.46 0.44 1.79 RV 0.87 0.72 0.39 1.53 TV 0.84 0.32 ND ND MNV-1 0.39 0.32 ND ND
81
2.4.3. Thermal inactivation of viruses within oyster tissues.
To study the feasibility of thermal treatment at 80˚C to inactivate enteric viruses in oyster tissues, 25 oysters were artificially contaminated with HAV, RV,TV or MNV-1, by growing the oysters in a contaminated salt water contained 1 × 105 PFU /ml of the tested viruses for 24 h. After the virus bio-accumulation, three oysters were randomly selected and assigned for each treatment time point. Heat treated and untreated oysters were shucked and homogenized, and the level of survived virus was quantified by plaque assay. The log10 PFU/g of virus survival of HAV, RV, TV, and MNV-1 is presented in
Table 9.
The Weibull Model showed the best fit for virus inactivation in oyster tissues. The
TFL for the tested viruses was calculated from the Weibull model and is presented in
Table 9. TV, MNV-1, and RV showed a decline in the virus titer commensurate with an increase in treatment time. TV and MNV-1 titers were at undetectable levels (TV 4.8 log10 reduction and MNV-1 4.7 log10 reduction) in the oysters following treatment for 3 min at 80°C. A holding time of 4 min at 80˚C was required to eliminate 3.1 logs of RV
(complete inactivation). However, no significant decrease in HAV titer was observed even after 6 min of treatment at 80°C. The longest TFL-value was for HAV and the lowest value was to MNV-1, and based on the TFL the four viruses can be raked from the more heat stable to the less stable as the following: HAV>RV>TV>MNV-1.
82 Table 9. TFL-values of HAV, RV, TV, and MNV-1 at 80˚C the oyster tissues
TFL(min) R2 RMSE
MNV-1 0.61 ±0.47 0.946 0.66
RV 2.99 ±0.38 0.975 0.24
TV 1.46 ±0.26 0.989 0.27
HAV 19.99 ±1.0 0.73 0.14
2.4.4. The effect of the thermal treatment on HuNoV GII.4 VLPs: The most prevalent foodborne virus, HuNoV, cannot be grown in cell culture. Therefore, there is need to develop a surrogate assay to estimate the survival of HuNoV. One novel approach is to utilize viral receptor-binding activity as an indicator for HuNoV infectivity. Disruption of receptor binding activity will likely be lethal to virus as it is the first step in the virus life cycle. The cellular receptor for HuNoVs is the histo-blood group antigen (HBGA), a carbohydrate moiety that can be cross-linked to the surface of magnetic beads. Interestingly, it was found that porcine gastric mucin (PGM) contains
HBGAs which can be used as a source of HuNoV receptors. Based on this theory, a
PGM-magnetic bead (PGM-MB) binding assay was developed to detect intact HuNoV treated by high pressure processing. To determine whether this assay can be used to evaluate the survival of HuNoV treated by heat, we first test this using HuNoV VLPs as a model because VLPs processes authentic receptor binding activity as native virions. To do this, VLPs of HuNoV GII.4 were treated either at 80˚C for 10 s, 30 s, 60 s, and 5 min,
83 or at 100 ˚C for 5s. Treated and untreated VLPs were incubated with PGM-MBs for 30 min at room temperature. The beads were washed three times by PBS and re-suspended in 10 µl of PBS. The beads and the bound VLPs were loaded to a 15% SDS-PAGE gel and visualized by Coomassie blue staining. The protein density of VLPs bound to the beads after treatment at 80 ˚C for 10, 30, and 60 s and was similar to untreated VLPs
(Fig. 10a). A slight reduction in the protein density of the VLPs bound to the PGM-MBs was observed after heat treatment at 80 ˚C for 5 min compared to the original input VLPs
(Fig. 10b). The VLPs completely lost ability to bind to the PGM-MBs after treatment at
100 ˚C of 5s (Fig. 10c).
The data may indicate that thermal treatment completely inactivated HuNoV surrogates (TV and MNV-1) was not sufficient to inactivate the receptor binding activity of VLPs. However, higher temperature (100 ˚C of 5s) or longer holding time (80 ˚C for 5 min) can damage the receptor binding activity.
84
Figure 10. The effect of heat at 80, and 100˚C on the binding ability of the VLPs of
HuNoV GII.4 to bind to the beads, analyzed by 15% SDS-PAGE and stained by
Coomassie staining. Bound were the heat treated or untreated VLPs bound to the beads
after 30 min of shucking at room temperature. Input were the heat treated or untreated
VLPs loaded directly to the gel. Unbound were heat treated or untreated VLPs in the
washing solution after the beads were washed with PBS. (A) Purified VLPs treated at
80˚C for 10 and 30 s. (B) Purified VLPs treated at 80˚C for 1 min and 5min. (C) Purified
VLPs treated at 100˚C for 5 s. (D) % of VLPs binding to BGM-MB.
85 2.4.5. Effect of heat treatment on HuNoV GII.4 and TV. The above result showed that
HuNoV VLPs were much more stable than HuNoV surrogates (TV and MNV), based on the PGM-MB assay. One possibility is that the empty VLPs (without genome) were more difficult to be inactivated than native virions which contain capsid and genomic RNA. To test this hypothesis, we directly compared the stability of TV and a HuNoV GII.4 clinical isolate upon heat treatment followed by PGM-MB assay. TV was used as a control because it also utilizes HBGAs as receptors. Briefly, TV and HuNoV GII.4 were heated at 62, 72, and 80 °C for holding times between 2 s to 36 min in capillary tubes. The receptor binding ability of the heat treated or untreated virus samples were separated by
PGM-MB binding assay on the bead and the viral RNA was extracted. RT-qPCR was used to quantify RNA only from viral particles capable of binding to the PGM-MBs. The results of TV are shown in Figure 11. At 62°C (Fig. 11a), 72°C (Fig. 11b), and 80°C
(Fig. 11c), the reduction in the RNA copies of TV was less than 1 log10 RNA copies/ml.
86
Figure 11. TV heat inactivation at 62, 72, and 80˚C by using binding assay and RT- qPCR. TV was heated at 62, 72 and 80°C the receptor binding ability of the heat treated
or untreated virus samples were separated by PGM-MB RNA copies were (A) at 62°C,
(B) 72°C, and (C) 80°C.
87 Treatment at 72°C for 1 min resulted in a 0.7 log10 reduction of TV RNA copies, while no change in the RNA copies was notice after heat treatment at 62°C for 36 min.
Increasing the temperature to 80°C (Fig. 11c) resulted in 2.5 log reduction of RNA copies after 10 s.
For HuNoV GII.4, heat treatment at 62°C (Fig. 12a) resulted in 0.7 log10 reduction in the viral RNA copies, while treatment at 80°C lead to an approximate 1 log reduction of the
RNA copies (Fig. 12b). By comparing plaque assay and PGM-MB assay for TV, it was concluded that PGM-MB assay overestimated the survival of TV and it is not an appropriate assay to evaluate the survival of HuNoV treated by heat.
88
Figure 12. HuNoV heat inactivation at 62, and 72˚C by using binding assay and RT- qPCR. HuNoV was heated at 62 and 72°C the receptor binding ability of the heat treated or untreated virus samples were separated by PGM-MB RNA copies were (A) at 62°C, and (B) 72°C.
2.4.6. Effect of heat treatment on TV RNA level. The fact that TV RNA was not significantly reduced when TV was subjected to a lethal heat treatment (80oC for 10 s)
89 suggests that heat treatment did not physically damage viral genomic RNA. Since real- time RT-PCR only detected a small fragment from genomic RNA, it remains possible that genomic RNA was degraded upon a lethal dose of heat. To rule out this possibility, the full-length VP1 gene was amplified by One-step RT-PCR. As shown in figure 13, there was no loss in abundance of the VP1 gene even following 5 min of treatment at
80°C, indicating that the heat treatment did not degrade the viral RNA.
Figure 13. The effect of heat treatment at 80˚C on the capsid gene (VP1) of TV: The
VP1 of TV untreated or treated at 80˚C for 10s or 5 min was amplified by one-step RT-
PCR and visualized on 1% agarose gel electrophoresis.
90 2.4.7. The effect of thermal treatment on purified TV. To investigate the effect of heat treatment on viral capsid integrity, purified TV particles suspended in TNC buffer were heated at 80˚C for 5 s, 10 s, and 5 min. The treated and untreated particles were negatively stained by 1% uranyl acetate and visualized under TEM. At the shortest treatment time (5s) there was a clear change in the virion appearance compared to the untreated virus, including a loss of the normal round structure with rough edges observed on the particle and an apparent loss of genomic material from the center of the particle
(Figure 14). More severe damages were observed following 10 s and 5 min treatment times (Figure 14). These results indicate that the TV capsid lost its integrity following 5 s of the thermal treatment at 80˚C.
Figure 14. - Visualize the damaged TV particles after heat treatment at 80˚C for 5s,
10s, and 5 min. TV untreated or treated at 80˚C for 5 s, 10 s or 5 min were visualized by
EM after negatively stained by 1% ammonium molybdate. 91 2.5. Discussion
Approximately 7.6-14.5 million illnesses in the US are attributed to the consumption of contaminated seafood each year, and enteric viruses were responsible for more than 50% of these cases (Butt et al., 2004). Overall, epidemiological evidence has shown that more than 11 shellfish-borne viral outbreaks occur each year worldwide
(Bellou et al., 2013). Human enteric viruses are rather ubiquitous in harvested shellfish.
Keller et al. (2013) showed that 100% of shellfish samples collected from Vitória Bay,
Espírito Santo, Brazil were positive for RV and adenovirus. However, only 80% of the growing water samples were positive for these pathogens. The viral titers were 400 times higher in the shellfish samples compared to the levels in the growing water, indicating the high level of natural bioaccumulation that occurs (Keller et al., 2013). In the Galician
Rias area, the largest shellfish production area in the European Union, 55% of mussels, clams, and cockle samples were contaminated by HuNoV GI and GII and HAV (Polo et al., 2015). Thus, understanding the ecology and persistence of enteric viruses in shellfish is needed to help prevent future outbreaks.
In this study, we explored the localization, the pattern of the bioaccumulation, and the heat inactivation parameters of the enteric viruses most frequently associated with shellfish outbreaks. Data available for the heat inactivation of enteric viruses differs among research studies due to differences in the thermal treatment conditions such as temperature, holding time, sample matrix, data modeling, and virus type (Arthur &
Gibson, 2015). Therefore, comprehensive studies investigating the thermal inactivation of viruses, the natural bioaccumulation patterns of the viruses in shellfish, and thermal
92 inactivation of viruses in shellfish tissues is useful for industry and for regulatory purposes to improve seafood safety.
2.5.1. Virus bio-accumulation and distribution in oysters. Previous work has evaluated the distribution of bacterial pathogens in shellfish, however little is known about the bioaccumulation and distribution of human enteric viruses in shellfish. This study aimed to directly assess the bioaccumulation of the most important enteric viruses that are frequently associated with shellfish outbreaks including HuNoV, HAV, and RV.
Our results demonstrate that oysters can efficiently bioaccumulate all enteric viruses to a high titer from the growing water and can be detected in all oyster tissues within 72 h.
Although all the examined viruses were detected in different oyster tissues, the pattern of viral localization varied based on the exposure time and the virus. For instance,
TV, MNV-1, and HAV were detected in the digestive gland at a high titer in the early after viral exposure (24 h), while RV more likely to be detected in the gills and muscles.
Specific and nonspecific binding may be responsible for the virus’ localization within the oyster tissues. For instance, Maalouf et al. (2010 and 2011) showed that different oyster and clam tissues express carbohydrate ligands similar to the histo-blood group antigens
(HBGAs), the cellular receptor of HuNoVs. The expression of these ligands varies based on the season and the organ (Maalouf et al., 2010, Maalouf et al., 2011). The finding that many members of Caliciviridae, such as HuNoV and TV, are highly accumulated in the digestive tissues of oysters within 24 h suggests that specific interaction of HuNoV and
TV with the HGBA-like carbohydrates expressed in these oyster tissues may play a role in virus bioaccumulations.
93 It has been shown that the viral genotype and also the season have an impact on virus localization. For instance, Maalouf et al. (2010) compared the bioaccumulation of three HuNoV strains belonging to genotypes GI.1, GII.4, and GIII.3 in oyster
(Crassostrea gigas) tissues. The GI.1 strain had the highest bioaccumulation level compared to the GII.4 strain in the digestive organs. This effect was increased in the winter season where the HBGA-like carbohydrate ligand is expressed at higher levels in the oyster digestive organs. In addition to the digestive tissues, the GII.4 strain was detected in all other oyster tissues. This result is thought to be due to the fact that the
GII.4 strain can recognize a sialic acid-containing ligand as an alternative receptor, which allowed for the bioaccumulation in other shellfish tissues. GIII.3 was accumulated most in the gills and mantle and was subsequently concentrated in digestive gland (Maalouf et al., 2010). Drouaz et al. (2015) compared the distribution of HuNoV GI, TV, and mengovirus (MgV) in oyster tissues after 24 h of oyster cultivation in contaminated seawater. The results showed that HuNoV GI and TV were more accumulated in digestive glands, while MgV was more concentrated in gills and the three viruses persisted in oyster tissues for more than 50 days (Drouaz et al., 2015). These data, together with our findings on HuNoV GII.4, RV, MNV-1, HAV, and RV, exemplifies that different enteric viruses have different distribution patterns in oyster organs based on the virus, virus genotype, shellfish type, and the season.
2.5.2. Thermal inactivation of viruses. To date, there is no standard method that has been established to study the thermal stability of enteric viruses. In this study, we systemically compared stability of four viruses upon heat treatment and found that the
94 thermal stability can be raked from the more heat stable to the less stable as the following: HAV>RV>TV>MNV-1.In general it has been shown that increasing the treatment time increases viral inactivation by thermal treatment. For example, treatment temperatures less than 60˚C required longer holding times to achieve high viral inactivation. At higher temperature (for example 80˚C), most human enteric viruses are rapidly inactivated. However, HAV and RV showed more heat resistance compared to other enteric viruses. In our study we found that MNV-1 was the most sensitive virus to heat treatment compared to the other viruses examined. Previously it was shown that
MNV-1 was reduced by 1 log at 63°C in <25 sec and at 72°C in <10 sec (Cannon et al.,
2006). Similarly we found the D62°C value for MNV-1 to range between 0.55-1.81 min and the D72°C value between 1.13-7.29 s. In another study the thermal stability of MNV-1 and TV was compared and both viruses were inactivated beyond the detection limit
(approximate 6 log reduction) at temperatures of 70°C and 75°C in cell culture medium
(Hirneisen et al., 2013). At lower temperatures, the thermal inactivation kinetics of both
MNV-1 and TV was similar indicating that both viruses are sensitive to heat treatment
(Hirneisen et al., 2013). We found that both MNV-1 and TV had very similar D-values at temperatures of 62, 72, and 80°C, suggesting both viruses are equally sensitive to heat treatment.
We found that viral thermal inactivation does not follow a linear logarithmic model. In a linear logarithmic model the slope of the inactivation curve can be used to calculate the decimal reduction value (D- value), the time required to reach one log reduction of the virus at a specific temperature (Chick, 1908; Tuladhar, et al., 2012).
Several other models have been used to describe virus inactivation by heat. For example,
95 the biphasic reduction model is used to calculate the time to the first log10 reduction
(TFL-value), instead of calculating D-value, from the inactivation curve based on two different rates of inactivation (de Roda Husman, et al., 2009; Tuladhar, et al., 2012; van
Boekel, 2002; Bozkurt et al., 2014). The Weibull model Cwas described by van Boekel
(2002) and it has been shown as the best fit to express virus inactivation in many studies
(van Boekel, 2002). In the Weibull model the inactivation parameters, α and β, and the scale and shape of the probability density function are used to describe the time required for a desired amount of inactivation at a specific temperature (Bozkurt et al., 2014). In our study, we found that at temperatures of 62 and 72°C the biphasic reduction model was the best fit to the data. This is due to the shouldering and tailing effect observed in the curve. However, at a treatment temperature of 80°C (culture medium or oyster), the
Weibull model was the best fit for the data. This is likely due to the increased linearity of the inactivation curve due to the increase temperature. These results demonstrate the necessity of applying appropriate models to accurately determine the D-values.
The complexity of the sample matrix has been shown to affect the efficacy of heat to inactivate enteric viruses. In our study, we found that the oyster provided a protective effect to the virus during heat treatment at 80°C. A treatment time of 3 min was required to inactivate TV (4.8 log reduction) and MNV-1 (4.7 log reduction) in oyster tissues, while 4 min was required to completely inactivate RV (3.1 log reduction) in oysters.
Even after 6 min of treatment at 80°C, there was no significant reduction in the titer of
HAV in oyster tissues. Previously, Croci et al. (1999) showed that heat treatment of HAV at 60 ˚C for 30 min and for 10 min at 80 ˚C was not sufficient to eliminate 5 logs of the virus from contaminated mussel tissues. In fact, a treatment at 100˚C was required to
96 inactivate HAV in the mussels (Croci et al., 1999). HAV was also more resistant to thermal inactivation in the dried mussels compared to culture medium (Park and Ha,
2015). Only 3.16 and 4.38 log reduction in HAV titer was achieved at 60 and 85˚C after
30 and 10 min of heat treatment, respectively. The D-values at 60, 85, and 100˚C were
6.3, 0.98, and 0.2 min, respectively, in culture medium; whereas, in dried muscles the D- values were increased to 7.93, 3.05, and 0.85 min at 60, 85, and 100˚C, respectively (Park and Ha, 2015). Soft-shell clams were artificially contaminated with either HAV or
MNV-1 and the efficacy of thermal treatment on viral inactivation was evaluated. It was found that in shucked clams a treatment at 90°C for 180 seconds was required to completely inactivate the viruses (Sow et al., 2011).
2.5.3. Insight into the mechanism underlying thermal inactivation of viruses.
Currently, nucleic acid based detection methods are used to test for HuNoV in foods.
These methods do not discriminate between RNA detected from an infectious or non- infectious viral particle, and therefore positive results may not accurately reflect the risk of disease development upon consumption. Recently, porcine gastric mucin conjugated to magnetic beads (PGM-MBs) has been developed to capture HuNoV particles capable of binding to their cellular receptor, the histo-blood group antigens (HBGAs) (Dancho et al., 2012; Tian et al., 2005; Tian et al., 2008; Tian et al., 2010). HuNoV bound to PGM-
MBs can then be collected, excluding particles which have lost receptor binding ability, and theoretically allow for RNA detection from only intact HuNoV particles. The PGM-
MBs have been used to detect HuNoV inactivation by HPP and UV (Dancho et al., 2012) and subsequently for treatments with chlorine, chlorine dioxide, peroxyacetic acid,
97 hydrogen peroxide, and trisodium phosphate (Kingsley et al., 2014). TV also utilizes
HBGAs as cellular receptors (T. Farkas et al., 2010), so there is the potential to use this cultivatable virus as a side-by-side control to determine the ability of the PGM-MB binding assay to discriminate between infectious and non-infectious virus particles.
In this study, we attempt to utilize PGM-MB assay to discriminate infectious and noninfectious HuNoV GII.4 strain by heat treatment. TV was used as a critical control as it is cultivable and utilizes same receptors as HuNoV does. Unfortunately, the log reduction of TV obtained from PGM-MB assay was not consistent with the survival data generated from plaque assay. Thus, this assay is not ideal to estimate the survival of
HuNoV or TV by heat. For example, TV RNA copies were not significantly reduced when TV was treated by a lethal dose of heat (80 oC for 10s). PGM-MB assay is a receptor binding based assay which requires the technologies to efficiently inactivate the receptor binding capability. Thus, the applicability of PGM-MB will depend on the mechanism of the inactivation induced by a particular thermal or non-thermal processing technology. Perhaps, other technologies (such as HPP, UV, and chlorine) may more efficiently damage the receptor binding than heat treatment. Based on the heat treatment of VLP, we found that receptor binding of HuNoV was severely damaged only at higher temperature (100˚C for 5 s) or longer holding time (80˚C for 5 min). Perhaps, heat treatment first damages the interaction between capsid and genomic RNA which is lethal to virus. However, the receptor binding capability of capsid is still intact, thus can efficiently bind to PGM-MB. Furthermore, the genomic RNA is still binding to the capsid, thus can be detected by real-time RT-PCR. Using purified TV, we found that the integrity of viral capsid was disrupted by a lethal heat dose. TV infectivity was lost
98 quickly (<10 s) during treatment 80°C, which correlates with the change in the particles observed by TEM. However, it appears that the viral particles still efficiently bind to the
PGM-MBs and that the viral RNA was minimally impacted by thermal treatment. No significant RNA reduction was observed at a lethal dose of heat. Using the PGM-MB binding assay followed by RT-qPCR to detect HuNoV inactivation by thermal treatment at 80°C for 1 min a maximum 1 log reduction was detected. Similarly, TV treated at
80°C for 1 min had a maximum 2.5 log reduction. Direct RT-PCR detection of the TV
VP1 gene also confirmed that viral RNA remained intact following the most severe thermal treatments. Perhaps, severe heat treatment did damage the receptor binding activity. HuNoV GII.4 VLPs retained the ability to bind to PGM-MBs following 80°C treatment as detected by SDS-PAGE analysis. In fact, a treatment of 100°C for 5 s was required to inhibit HuNoV VLP binding to PGM-MBs. Taken together these results indicate that disruption of the integrity of viral capsid and denaturation of viral protein, but not degradation of viral RNA is the primary mechamsim of viral inactivation by heat.
The fact that a lethal dose of heat was not sufficient to damage viral receptor binding activity suggests heat may disrupt the capsid-genome interaction which is lethal to virus infectivity.
Overall this study demonstrate, (i) enteric viruses can be efficiently bioaccumulated in oyster tissues within 72 hours; (ii) differences between the major bioaccumulation site in oysters can observed between different viruses early after virus exposure (24 hours); (iii) enteric virus resistance to heat treatment can be ranked as HAV
(most resistant)>RV>TV>MNV-1(most sensitive); (iv) oyster tissue provides a protective effect to viruses during heat treatment; and (v) thermal treatment 80oC for more than 6
99 min is capable of eliminating major foodborne viruses from oysters . These results can be used by industry professionals to design and implement prevention and control measure to limit virus-shellfish associated outbreaks.
100
CHAPTER 3
High-pressure Inactivation of Rotaviruses: The Role of Treatment Temperature and Strain Diversity in Virus Inactivation
3.1. Abstract
Rotavirus (RV) is the major etiological agent of acute gastroenteritis in infants worldwide. Although high pressure processing (HPP) is a popular method to inactivate enteric pathogens in food, the sensitivity of different virus strains within same species and serotype to HPP is variable. This study aimed to compare the baro-sensitivity of seven RV strains derived from four serotypes (G1: Wa, Ku, and K8, G2: S2, G3: SA-11 and YO, and
G4:ST3) following high pressure treatment. RV strains showed varying responses to HPP based on the initial temperature and had different inactivation profiles. Ku, K8, S2, SA-11,
YO, and ST3 showed enhanced inactivation at 4°C compared to 20°C. In contrast, Wa strain was not significantly impacted by the initial treatment temperature. Within serotype
G1, Wa strain was significantly (p<0.05) more resistant to HPP compared to Ku and K8.
Overall, the resistance of the human RV strains to HPP at 4°C can be ranked as
Wa>Ku=K8>S2>YO>ST3 and in terms of serotype G1>G2>G3>G4. In addition, pressure treatment of 400 MPa for 2 min was sufficient to eliminate the Wa strain, the most pressure resistant RV, from oyster tissues. HPP disrupted virion structure, but did not degrade viral protein or RNA, providing insight into the mechanism of viral inactivation by HPP. In
101 conclusion, HPP is capable of inactivating RV at commercially acceptable pressures and the efficacy of inactivation is strain dependent.
3.2. Introduction
Rotavirus (RV) is the major etiological agent of acute gastroenteritis in infants worldwide (Desselberger, 2014; Parashar, 2006). RVs are estimated to cause nearly
500,000 deaths annually among children (FAO/WHO, 20089, CDC, 2013). The virus is transmitted by the fecal-oral route, and contaminated water and food are common vehicles for infections (FAo/WHO, 2008, CDC, 2013). RV belongs to the genus
Rotavirus, subfamily Sedoreovirinae, and family Reoviridae. There are eight species
(groups) of rotavirus, referred to as A, B, C, D, E, F, G, and H. Humans are primarily infected by species A, B and C, most commonly by species A. Rotavirus species A can be further divided into different serotypes. RV is a segmented double-stranded RNA virus with a triple-layered icosahedral capsid. The outer capsid glycoprotein (VP7) and the spike protein (VP4) differentiate RVs into 14 G (Glycoprotein) serotypes and 27 different P (Protease sensitivity) genotypes, respectively (Desselberger, 2014; Kollaritsch et al., 2015; Mameli, 2012). Currently, five serotypes (G1–G4, and G9) are the predominant circulating viruses, accounting for almost 95% of strains worldwide
(Desselberger, 2014). Recently, commercial RV vaccines have been used in children to provide immunity against the most commonly circulating strains (Kollaritsch et al.,
2015). Despite major efforts, RV outbreaks still occur worldwide due to the high genetic diversity of RVs and lack of cross-protection (Mwenda 2010, Bost et al., 2000, Kittigul, et al., 2014). Therefore, alternative strategies for the prevention of RV infection must be established. 102 Enteric viruses are a leading cause of foodborne illnesses. Within foodborne viruses, human norovirus (NoV), rotaviruses (RV), hepatitis A virus (HAV), and human sapovirus are the most prevalent viral pathogens associated with foodborne outbreaks
(CDC, 2013; FAO/WHO, 2008; Jaykus and Escudero-Abarca, 2010; Koopmans and
Duizer, 2004). In recent years, epidemiological evidence indicates that viruses cause the majority of outbreaks associated with bivalve shellfish (Lees, 2000; Potasman and Odeh,
2002). RV has frequently been detected in both fresh and marine water sources (Lodder and de Roda Husman, 2005; Prevost et al., 2015). As a consequence, RVs are often found to contaminate bivalve shellfish (Gabrieli et al., 2007; Leguyader et al., 1994;
Loisy et al., 2005; Quiroz-Santiago et al., 2014). In a survey of 300 shellfish (including oysters, muscles, and cockles) harvested in growing waters off the coast of Thailand, RV was detected in 8% of the samples (Kittigul et al., 2015). In a survey of oysters in
Mexico City (n=30), 26.9% were found to contain RV (Quiroz-Santiago et al., 2014).
Although outbreaks of RV are rare in adults and infections are typically asymptomatic, infected adults may unknowingly expose infants, the elderly, and the immunocompromised to the virus (Desselberger, 2014). Therefore, there is an urgent need to develop technologies that can inactivate RV in foods while maintaining the sensory and nutritional qualities of the product.
High pressure processing (HPP) is a promising non-thermal technology that inactivates foodborne viruses while maintaining the organoleptic properties of processed foods (Cruz-Romero et al., 2004; He, 2002; Hoover, 1989; Lou et al., 2015; Rastogi et al., 2007). The technology applies hydrostatic pressure instantaneously and uniformly throughout foods regardless of size, shape, and geometry, thus inactivating both surface
103 and internalized pathogens (Hoover, 1989; Lou et al., 2015). HPP is an increasingly popular method used by the shellfish industry to inactivate Vibrio parahaemolyticus, enteric viruses, and other pathogens (Abdel Karim, 2011; Lou et al., 2015; Rastogi et al.,
2007). HPP levels up to 600 MPa for several minutes is sufficient to inactivate most pathogenic microorganisms, such as bacteria and viruses (Lou et al., 2015). In addition to ensuring the safety of the shellfish, HPP treatment between 100-600 MPa separates the meat of the shellfish from their shell, which minimizes labor costs (Cruz-Romero, 2004;
Lopez-Caballero et al., 2000; Lou et al., 2015). High pressure treated oysters are more voluminous, juicy, and higher in moisture content compared to the untreated oysters
(Kingsley, 2014). It was reported that pressure treated oysters are also more desirable based on sensory evaluations (Lou et al., 2015; Ye et al., 2015).
To effectively inactivate pathogens, it is critical to optimize the conditions for pressure treatment. The effectiveness of HPP is influenced by many factors including both processing parameters (pressure, temperature, and holding time) and non-processing parameters (virus structure itself, food matrix, pH, and aw of foods). In general, it has been established that increasing both the treatment pressure and treatment time increases viral inactivation. However, one interesting observation for HPP inactivation of viruses is that different viruses are variable in their susceptibility to high pressure. For example, enveloped viruses are less stable to environmental stresses than non-enveloped viruses.
However, some enveloped viruses (e.g. vesicular stomatitis virus, VSV) are much more stable than non-enveloped viruses during HPP treatment. In addition, viruses are more stable in cold environments compared to room temperature. However, many viruses (e.g.
104 human NoV, murine norovirus, and Tulane virus) are more easily inactivated by HPP at cold temperature (e.g. 4˚C) than at ambient temperature.
According to the International Committee on Taxonomy of Viruses (ICTV), viral classification starts at the level of order and continues as follows: family, subfamily, genus, species, and serotype. To date, only a few studies have shown that different viruses from the same family or genus have variable high pressure susceptibilities.
However, it is not known whether different viruses from the same species or serotype have different pressure susceptibilities. The genus Rotavirus has substantial genetic diversity. For instance, the amino acid homology of the capsid proteins of species A rotavirus strains can range from 70-95%. This high genetic diversity makes rotavirus an ideal model to study the role of strain diversity in pressure sensitivity.
This study aims to compare the baro-sensitivity of different RV strains derived from four serotypes to HPP and to gain a better understanding of the correlation between strain diversity and pressure resistance. Understanding this fundamental question will help to optimize the conditions for pressure inactivation of RVs and facilitate the development of technologies to eliminate RVs from high risk foods.
3.3. Materials and Methods
3.3.1. Viruses and cell culture. Viruses and cell culture. Seven RV strains were used in this study. These strains include serotype G1 (Wa, Ku, and K8 human strains), G2 (S2 human strain), G3 (SA-11 simian strain and YO human strain), and G4 (ST3 human strain).
For tissue culture adaption, RV strains S2 and ST3 were first propagated in grivet monkey kidney (BGM-70) cells before being adapted to rhesus monkey kidney cells (MA-104) cell
105 line. All other strains were grown in MA-104 cells cultured in Eagle's minimum essential medium (MEM). Briefly, confluent MA-104 cells in T75 flasks were infected by each RV strain at a multiplicity of infection (MOI) of 1.0. After 1 h incubation, 10 ml of MEM containing 6 µg/ml of trypsin (Invitrogen) were added. At 48-96 h post-infection, viruses were harvested by three freeze-thaw cycles and centrifugation at 1,500 × g for 15 min. The virus titer was determined by plaque assay and virus stocks were stored a -80 °C.
3.3.2. RV plaque assay. RV plaque assay was performed as described previously (Lou et al., 2011). Briefly, monolayers of MA-104 cells were grown in six-well plates (Corning
Life Sciences, Wilkes-Barre, PA) at a density of 2 × 106 cells per well. 400 µl of a 10-fold dilution series of RV was added to each well and the plates were incubated for 1 h at 37°C with agitation every 10-15 min. The plates were overlaid with 1% agarose MEM in the presence of 2.5 µl/ml trypsin, 1% sodium bicarbonate, 0.1 mg of kanamycin/ml, 0.05 mg of gentamicin/ml, 15 mM HEPES (pH 7.7), and 2 mM L-glutamine. Plates were incubated at 37°C and 5% CO2 for 72 h. A 10% formaldehyde solution was used to fix the plates for
2 h, and plaques were visualized by crystal violet staining. Viral titer was expressed as log10 plaque forming unit (PFU)/ml.
3.3.3. Pressure inactivation of different RV strains. RV strains were treated by HPP at levels from 200 to 450 MPa with a holding time of 2 min at an initial temperature of 4°C or 20°C. Briefly, 400 µl of each RV strain was suspended in MEM medium and were double packaged and sealed in sterile polyethylene stomacher pouches (Fisher Scientific
International, Ontario, Canada). Samples were subjected to HPP treatment using a lab
106 scale HPP unit (model Avure PT-1; Avure Technologies, Kent, WA) with water as the hydrostatic fluid. The 2 min holding time did not include the pressure come-up and release time (22 MPa/s, and 4 seconds, respectively). The virus survival was determined by plaque assay and expressed as log10 PFU/ml.
3.3.4. Purification of RV. Purification of RV Wa and SA-11 strains was performed using the protocol described previously ((Lou, Neetoo, Li, & Chen, 2011). Approximately 180 ml of RV Wa or SA-11 strain stock (1.5 × 106 PFU/ml) was centrifuged at 82,000 × g through a 40% (wt/vol) sucrose cushion at 4°C for 5h, in a Ty50.2 rotor (Beckman Coulter,
Fullerton, CA). The virus pellet was re-suspended in 300 µl of TNC buffer (100 mM NaCl,
10 mM Tris, 1 mM CaCl2) on ice overnight. The virus suspension contained a mixture of double-layered particles (DLPs) and triple-layered particles (TLPs) of RV. A CsCl isopycnic gradient (1.37 g/ml) was used to separate the TLPs and DLPs in a SW50.1 rotor
(Beckman Coulter) centrifuging at 115,000 × g at 4°C for 18 h. The upper and lower bands containing TLPs and DLPs, respectively, were separately collected and re-suspended in
TNC buffer (0.05 M Tris-HCl, 0.15 M NaCl, 15 mM CaCl2, pH 6.5). The TLPs and DLPs were further purified by ultra-centrifugation at 68,000 × g for 2 h at 4°C, and the final TLP or DLP pellets were re-suspended in 300 μl of TNC buffer.
3.3.5. Reverse transcriptase polymerase chain reaction (RT-PCR). Reverse transcription polymerase chain reaction (RT-PCR) was used to determine whether the VP7 gene, the RV viral capsid gene, was degraded by HPP. Total viral RNA of Wa or SA-11 strains was extracted from RV treated at 400 or 550 MPa and untreated virus using the
107 RNeasy Mini Kit (Qiagen, Valencia, CA). Two primers were used to amplify the VP7 gene
(5’-GAGAGAATTTCCGTCTGGCTAA-3’ and 5’-CTTGCCACCACTTTTTCCAAT-3’) for Wa strain and (5’GGTCACATCATACAATTCTAACC-3’ and
5’GGCTTTAAAAAGAGAGAATTTCC) for SA-11. DNA bands were visualized using
1% gel electrophoresis.
In addition, the RNA of RV (Wa) was extracted from the virus particle. Viral RNA was either directly subjected to HPP treatment or 20µl of viral RNA was treated with 1µl
RNase-out (diluted 1:10) prior to HPP treatment. Viral RNA without RNase-out or with
RNase-out was then amplified by RT-PCR. This method was used to establish whether
HPP treatment directly degraded viral RNA.
3.3.6. Transmission electron microscopy. 10µl of purified TLPs and DLPs of Wa or SA-
11 strains were treated at 200, 400 and 600 MPa for 2 min at an initial temperature of 4°C, control samples were untreated. The samples were then fixed to copper grids (Electron
Microscopy Sciences, Inc.) and subjected to negative staining using uranyl acetate. Virus particles were then visualized by a FEI Tecnai G2 Spirit Transmission Microscope at the
Microscopy and Imagining Facility at the Ohio State University.
3.3.7. Analysis of RV proteins by SDS-PAGE. 10 µl of purified treated (200, 400, 600
MPa) or untreated TLPs and DLPs of Wa and SA-11 strains were examined by SDS-
PAGE. Each sample was mixed (1:4 V/V) with SDS-PAGE loading buffer which consists of 1% SDS, 2.5% β-mercaptoethanol, 6.25 mM Tris-HCl (pH 6.8), and 5% glycerol. The
108 mixture was boiled for 5 min and loaded to a 15% polyacrylamide gel. The protein bands were visualized by Coomassie blue staining.
3.3.8. Bioaccumulation of RV in oyster tissues. To mimic the natural bioaccumalition of
RV in oysters, twenty five live oysters (Kroger Co.) were cultivated in 4 L of salt water containing 1 × 106 PFU/ml of Wa for 24 h, under aeration conditions with phytoplankton feed. Following bioaccumulation, the contaminated oysters were shucked and the meat was packaged and sealed in sterile polyethylene stomacher pouches. The oyster meat was treated at pressure levels ranging from 200-500 MPa for 2 min at an initial temperature of
4°C or 20 °C. Following HPP treatment, the oyster meat was homogenized in 5ml HBSS using a mortar and pestle. Virus survival was determined by plaque assay.
3.3.9. Statistical analysis. All experiments were performed in triplicate. Virus titer was expressed as mean log PFU/ml ± 1 standard deviation. Statistical analysis was performed by one-way multiple comparisons using SPSS 8.0 statistical analysis software (SPSS Inc.,
Chicago, IL). A P value of <0.05 was considered statistically significant.
3.4. Results
3.4.1. The effect of HPP initial temperature on different RV strains. The initial temperature of HPP is a critical parameter influencing the effectiveness of virus inactivation: working either synergistically or antagonistically with pressure. Chilling
(4°C) and ambient (20°C) temperatures were selected to avoid the thermal factor
(combination of heat and pressure) in comparing HPP inactivation of different RV strains.
109 To determine the role of temperature on HPP inactivation of RV strains, seven RV strains of G1 (Wa, Ku, and K8 human strains), G2 (S2 human strain), G3 (SA-11 simian strain and YO human strain), and G4 (ST3 human strain) were treated by different levels of HPP ranging from 200-450 MPa at an initial temperature of either 4 or 20°C for a holding time of 2 min. Plaque assay was used to determine virus survival.
The results for the G1 strains are shown in figure 15. In general, increasing the pressure level enhanced virus inactivation. At the initial temperature of 20°C a shoulder effect was observed at low HPP levels (200 MPa) and a tailing effect was observed at high HPP levels
(400 and 450 MPa) for all three G1 strains (Fig. 15). There was no significant difference in the inactivation kinetics of Wa strain at either initial temperature, 4 or 20°C (P>0.05). A pressure level of 450 MPa for 2 min was not sufficient to completely inactivate the Wa strain (5.9 log10 PFU) suspended in culture medium and 1.4 and 1.8 log10 PFU of Wa survived following 450 MPa treatment at either 4 or 20°C, respectively (Fig. 15a). Ku was more susceptible to HPP at the low temperature (4°C) compared to an initial temperature of 20°C (Fig. 15b).
110
Figure 15. Effect of temperature on inactivation of RV serotype G1 strains. RV stock
(106PFU/ml) in cell culture medium (MEM) was processed under pressures ranging from
200 MPa to 450 MPa held for 2 min at either 4°C or 20°C. Data are the means of three replicates. Error bars represent standard deviations. (A) The effect of temperature on Wa strain; (B) The effect temperature on Ku strain; and (C) The effect of temperature on K8
strain. 111 At 4 °C, 400 MPa treatment reduced the Ku titer to below the detection limit (Fig.
15b). However, 1.8 log10 PFU of Ku was still detected at an initial temperature of 20°C after 400 and 450 MPa treatments (Fig. 15b). Similarly, K8 strain showed the same high baro-sensitivity as Ku at low initial temperature (4 °C) (Fig. 15c). A 5-log reduction of K8 was achieved at 400 MPa at 4°C (Fig. 15c). Conversely, approximately 1 log10 PFU of K8 was detected at 450 MPa at 20°C (Fig. 15c).
The HPP inactivation profile of the G2 virus, S2 strain is presented in Figure 16.
S2 was more susceptible to HPP at a low initial temperature compared to a high initial temperature. Treatment of 450 MPa at 4 °C for 2 min reduced the S2 titer below the detection limit (Fig. 16). In contrast, 1.5 log10 PFU of S2 remained after the treatment of
450 MPa at 20 °C for 2 min (Fig. 16).
Viruses in genotype G3 (SA-11 and YO) had differing inactivation patterns in response to high pressure treatment depending on initial treatment temperature (Fig. 17).
YO was more susceptible to pressure treatment at the low initial temperature. At 4°C, a
5.3-log reduction was achieved at 400 MPa (below detection limit); however at 20°C, a tailing effect was observed with 1 log10 PFU detected following 450 MPa HPP treatment
(Fig. 17b). On the other hand, there was no significant effect of the initial temperature on
SA-11 inactivation. Approximately 1 log10 PFU of SA-11 was still viable after 450 MPa of pressure treatment at both 4°C and 20°C (Fig. 17a).
112
Figure 16. Effect of temperature on inactivation of RV serotype G2 strain S2. RV
stock (106PFU/ml) in cell culture medium (MEM) was processed under pressures ranging from 200 MPa to 450 MPa held for 2 min at either 4°C or 20°C. The surviving viruses were determined by plaque assay. Data are the means of three replicates. Error bars represent standard deviations. Viral reduction was significantly different between
the initial temperature of 4°C and 20°C (P<0.05).
113
Figure 17. Effect of temperature on inactivation of RV serotype G3 strains. 7.5
log10 PFU/ml of RV strain SA-11 and 5.5 log10 PFU/ml of YO strain in cell culture medium (MEM) were processed under pressures ranging from 200 MPa to 450 MPa held
for 2 min at either 4°C or 20°C. Data are the means of three replicates. Error bars
represent standard deviations. (A) The effect of temperature on SA-11 strain (B) The effect temperature on YO strain. Viral reduction was significantly different between the
initial temperature of 4°C and 20°C (P<0.05).
114 For the G4 strain, ST3, decreasing the initial temperature to 4°C resulted in a significant enhancement in viral inactivation compared to 20°C. Treatment of 400 MPa at
4°C reduced the level of ST3 below the detection limit (Fig. 18). At 20°C, > 1 log10 PFU remained after treatment of 450 MPa (Fig. 18).
Figure 18. Effect of temperature on inactivation of RV serotype G4 strain ST3. RV
stock (106PFU/ml) in cell culture medium (MEM) was processed under pressures
ranging from 200 MPa to 450 MPa held for 2 min at either 4°C or 20°C. The surviving
viruses were determined by plaque assay. Data are the means of three replicates. Error
bars represent standard deviations. Viral reduction was significantly different between
the initial temperature of 4°C and 20°C (P<0.05).
115 3.4.2. Comparing the baro-sensitivity of different human RV strains. Since all RV strains tested were more susceptible to HPP at 4°C, we directly compared the sensitivity of all RV strains at this temperature. As shown in figure 19, RV strains showed different inactivation profiles due to HPP treatment, even within strains of the same serotype.
Figure 19. Direct comparison of pressure inactivation of six human RVs derived
from four serotypes. RV stock (106 PFU/ml) in cell culture medium was processed
under pressures ranging from 200 MPa to 450 MPa held for 2 min at 4°C. Data are the
means of three replicates. Error bars represent standard deviations. Wa strain was significantly (p<0.05) more resistant to HPP inactivation compared to Ku and K8. S2 was
more significantly (p<0.05) inactivated by HPP compared to G1 strains (Wa, Ku, and
K8). YO had significantly (p<0.05) increased inactivation by HPP compared to the G2
and G1 strains. ST3 was the most sensitive strain (p<0.05).
116 G1 Wa strain was significantly (p<0.05) more resistant to HPP inactivation compared to Ku and K8. Overall, the level of resistance of G1 strains to HPP at an initial temperature of 4°C can be ranked as Wa>Ku=K8. The G2 strain, S2, was more significantly (p<0.05) inactivated by HPP compared to G1 strains (Wa, Ku, and K8) at the low initial temperature (4°C). The G3 strain, YO, had significantly (p<0.05) increased inactivation by HPP compared to the G2 and G1 strains at an initial temperature of 4°C.
ST3, the G4 strain, was the most sensitive strain to HPP treatment at an initial temperature of 4°C. Overall, the resistance of the six strains to HPP treatment at an initial temperature of 4°C can be ranked as Wa>Ku=K8>S2>YO>ST3 and in terms of serotypes
G1>G2>G3>G4.
3.4.3. The effect of HPP on RV capsid. To understand the mechanism of RV inactivation by HPP, we determined the effects of HPP on the viral capsid, proteins, and genomic RNA using two RV strains, a human strain (Wa) and a simian strain (SA-11). Briefly, purified
TLPs and DLPs of Wa and SA-11 strain suspended in TNC buffer were pressure treated at levels between 200 and 600 MPa at 4˚C for a 2 min holding time. The pressured particles were negatively stained with ammonium molybdate and examined by TEM. At the lowest pressure level applied, 200 MPa, the TLPs were intact and the appearance was similar to untreated samples (Fig. 20). At the same pressure level, the DLPs were damaged and high levels of debris were observed. By increasing the pressure level to 600 MPa, the TLPs and
DLPs were completely disrupted and no intact particles were observed (Fig.20).
117
Figure 20. HPP disrupts the integrity of RV particles. Purified TLPs and DLPs of RV
Wa and SA-11 strain were treated by HPP at 200, 400, 600 MPa at 4°C for 2 min. HPP- treated and untreated samples were negatively stained by 1% ammonium molybdate and
visualized by transmission electron microscopy.
These results indicate that the RV capsid can be completely disrupted at 600 MPa for 2 min at 4°C. Also, it was observed that TLPs (complete viral particle) were more
118 resistant to HPP treatment than DLPs, which lack the outermost layer of the complete virion, which is comprised of VP4 and VP7.
3.4.4. HPP effect on the viral proteins. To investigate the effect of HPP on viral proteins of different RV strains, we compared the most HPP resistant strain (Wa) and the simian strain (SA-11). 10µl of purified TLPs and DLPs of Wa or SA-11 were treated at 200, 400, and 600 MPa for 2 min at 4°C, and viral proteins were analyzed by SDS-PAGE. For both
Wa and SA-11 strains, six structural proteins (VP1, VP2, VP3, VP4, VP6, and VP7) were observed in untreated and treated TLPs. HPP treatment of 200, 400, and 600 MPa did not alter the abundance of viral proteins (Fig. 21a and 21b). Similarly, the density of the proteins of DLPs was not significantly altered by HPP treatment for both Wa and SA-11 strains (Fig.21c and 21d). The data suggests that the primary structure of the viral proteins remained intact although the virion structure was completely disrupted at a pressure level of 600MPa.
119
Figure 21. The effect of HPP on RV proteins. Purified RV TLPs and DLPs were
pressurized at 200, 400, and 600 MPa at 4°C for 2 min. The structural proteins of
untreated and treated RV were analyzed by 15% SDS-PAGE followed by Coomassie blue staining. (A) SDS-PAGE of 2 μg of total RV Wa TLPs. (B) SDS-PAGE of 2 μg of total RV SA-11 TLPs. (C) SDS-PAGE of 1.0 μg of total RV Wa DLPs. (D) SDS-PAGE
of 1.0 μg of total RV SA-11 DLPs.
120 3.4.5. The effect of HPP on viral RNA. To evaluate the effect of HPP on the virus genomic RNA, purified RV Wa strain with or without RNase inhibitor was treated by HPP ranging from 200 to 600 MPa at 4 °C for 2 min. After treatment, total RNA was extracted and subjected to RT-PCR targeting the outer capsid gene VP7, and the amplified DNA bands were visualized on 1% agarose gel electrophoresis. For RV without RNase inhibitor, the density for VP7 gene decreased as the pressure increased (Fig.22a). This suggests that exogenous RNase degraded the genomic RNA after the viral capsid was disrupted by HPP.
However, no significant decrease in the density of VP7 gene was observed between the pressurized particle and the control at any pressure level in an RNase free environment
(Fig.22b). To determine whether HPP directly degraded viral RNA, total viral RNA was extracted from RV Wa strain (without HPP), subjected to HPP treatment, and the VP7 gene was amplified by RT-PCR. As shown in figure 22c, there was no significant difference in
VP7 gene detected by RT-PCR between treated and untreated samples. This suggests that
HPP does not directly degrade RNA. This observation is consistent with the fact that HPP does not break covalent bonds.
121
Figure 22. The effect of HPP on viral genomic RNA. (A) The effect of HPP on RV genomic RNA in the absence of RNase inhibitor. Purified RV Wa strain were treated at
400 and 600 MPa for 2 min at 4˚C. After treatment, total viral RNA was extracted, and
the VP7 gene of RV was amplified by one-step RT-PCR and visualized on 1% agarose
gel electrophoresis. (B) The effect of HPP on RV genomic RNA in the presence of
RNase inhibitor. 1 unit of RNase inhibitor (RNase-out, Invitrogen) was added to the
purified RV Wa strain. The samples were treated at 400 and 600 MPa for 2 min at 4˚C.
After treatment, the VP7 gene of RV was amplified by one-step RT-PCR. (C) The effect of HPP on naked RV genomic RNA. Total viral RNA was extracted from RV Wa strain, treated at 400 and 600 MPa for 2 min at 4˚C, and the VP7 gene of RV was amplified by
one-step RT-PCR.
122 3.4.6. Inactivation of RV in oyster tissues. Finally, we determined whether HPP is capable of inactivating RV Wa strain in oysters, one of the high risk foods often contaminated by foodborne viruses. To mimic the natural contamination route, Wa strain was bio-accumulated in oysters by adding virus to feed water during oyster growth. After bioaccumulation, oysters were harvested and treated by HPP at pressure levels ranging between 200-500 MPa at either 4 or 20 ˚C with a holding time of 2 min. After HPP treatment, oysters were homogenized and virus survival was determined by plaque assay.
In oysters, the inactivation curve showed a shoulder effect at low pressure levels (200 and
300 MPa) (Fig. 23), which was similar to the effect observed when the virus was suspended in culture medium. Following treatment of oysters at 400 MPa, a 4.5 log10 PFU virus reduction was achieved and virus levels were below the detection limit at either initial temperature used (Fig. 23). There was no significant difference (p>0.05) of initial temperature on the virus inactivation in oyster tissues, which is consistent with the results obtained for Wa strain HPP inactivation in culture medium.
123
Figure 23. Inactivation of RV Wa strain in oyster tissues by HPP. 106 PFU of RV Wa
strain was added to the feed water during oyster growth. At day 3 post-inoculation,
oysters were harvested and treated by HPP at 200, 300, 400, and 500 MPa at 4°C for 2 min. After treatment, five oysters in each treatment were homogenized, and the surviving
RV was quantified by plaque assay. Data are the means of three replicates. Error bars
represent standard deviations.
3.5. Discussion
RV is a major cause of infant gastroenteritis and death worldwide (Desselberger, 2014;
Parashar et al., 2006). Although outbreaks in the US are less frequent due to vaccination,
RV remains a major public health concern in the developing world. The genus Rotavirus, is highly diverse and vaccination is insufficient to protect against all strains of RV
124 (Bishop,1996, Ghazanfar et al., 2014, Kittigul et al., 2014). In addition, many susceptible populations such as immunocompromised individuals cannot receive this vaccine. RV outbreaks have been associated with the consumption of contaminated foods and water, therefore the development of control measures to eliminate RV from food and water sources is critical to prevent the outbreaks.
HPP is a non-thermal process that can be used to eliminate foodborne pathogens while maintaining the organoleptic and nutritional properties of foods. To date, the HPP sensitivity of different virus strains in the same genus or family has not been well studied.
RVs are genetically diverse which makes it a good model to study the role of strain diversity in HPP inactivation of viruses (Kittigul, 2014). In this study, we found that different RV strains in same genus and genotype responded to HPP differently. Within same genus, the sensitivity of RV strains to HPP can be ranked G1>G2>G3>G4.
Interestingly, HPP inactivation of the Wa strain was not dramatically different in culture medium at 4°C and 20° C. Likewise, temperature differences were not apparent for Wa strain inactivation in oyster tissue. In contrast, all other RV strains were more easily inactivated at 4˚C compared to 20˚C. Within same genotype G1, Wa strain was significantly more stable than Ku and K8 strains. The different sensitivities of different
RV strains to HPP also raise the possibility that these strains may have different stabilities in the environment and in response to other treatments. It is possible that the difference in the degree of resistance to environmental stress may correlate to the difference in prevalence between those serotypes although direct evidence is lacking.
125 3.5.1. Role of initial temperature on the inactivation of seven RV strains.
Temperature is a critical factor influencing virus inactivation by HPP. The optimal initial temperature during HPP varies greatly between different viruses (Chen et al., 2005;
Cromeans et al., 2014; Grove, 2008; Guan et al., 2006; Kingsley et al., 2004; Li et al.,
2013; Lou et al., 2011). In this study, we choose to compare two different initial temperatures 4°C and 20°C for their influence on HPP inactivation of the seven RV strains. These temperatures were selected in order to avoid thermal effects caused by the combination of high temperature and HPP. RV strains Ku, K8, S2, SA-11, YO, and ST3 showed enhanced inactivation at an initial temperature of 4°C compared to 20°C.
Interestingly, HPP inactivation of the Wa strain was not significantly impacted by the initial temperature during treatment in culture medium or oyster tissues. This is consistent with our previous observation that approximately a 5-log virus reduction for the Wa strain was observed under 400 MPa at either 4 or 20°C (Lou et al., 2011). Overall, our results suggest that a low initial temperature increases the inactivation of a majority of the
RV strains by HPP. Also, it was previously reported that treatment of RV Wa strain with
300 MPa for 2 min at 25°C inactivated approximately 8 log10 TCID50/ml of the virus, although different temperature conditions were not compared in this study (Khadre &
Yousef, 2002). This is a much more dramatic reduction in the titer of Wa strain than what we observed in this study. It is possible that the initial viral titer used for treatment and/or the quantification methods (PFU vs TCID50) contributed to this difference. Collectively, these data demonstrate that HPP is capable of effectively inactivating RV.
The mechanism behind the increase in the inactivation at the lower initial temperature of the RV strains is not clear, however this effect has also been observed for
126 many other viruses. For example, viruses in the family Caliciviridae, including human
NoV, feline calicivirus (FCV), Tulane virus (TV), and murine norovirus (MNV), were all found to be more sensitive to HPP at low treatment temperature than at higher initial temperatures (Chen et al., 2005; Cromeans et al., 2014; Dancho & Kingsley, 2012;
Kingsley & Chen, 2009; Li & Chen, 2015; Li et al., 2013; Lou et al., 2011). In contrast, pressure inactivation of HAV, a non-enveloped virus in the family Picornaviridae, is enhanced at ambient and above temperatures compared to lower temperatures (Kingsley
& Chen, 2009; Kingsley et al., 2006; Kingsley et al., 2013). It has been suggested that at lower initial temperatures the viscosity of the hydrostatic fluid is altered, leading to increases inactivation of non-enveloped viruses (Balny, 2002, Kingsley, et al., 2008).
High pressure effects on biological macromolecules: from structural changes to alteration of cellular processes. However, this does not explain the increased sensitivity of HAV to
HPP at elevated temperatures. Also, a decrease in temperature in combination of HPP may also influence the stability of viral capsid, making the particle more easily disrupted at low temperatures compared to higher temperatures (Kingsley et al., 2009). More research is needed in order to establish the mechanism behind this phenomenon.
3.5.2. Role of strain diversity on HPP inactivation. It has been established that different viruses have differing optimal conditions for inactivation by HPP (Grove, 2008;
Kingsley & Chen, 2008, and 2009; Kingsley et al., 2005; Kingsley et al., 2006; Kingsley et al., 2007; Kingsley et al., 2002; Li et al., 2013; Lou et al., 2011a; Lou et al., 2011b;
Tang et al., 2010). However, the role of strain diversity within the same genus, species, or serotype on inactivation by HPP is still poorly understood. In this study, we examined
127 six human RV strains that belong to the most prevalent genotypes (G1-4) for sensitivity to HPP. We found that RV strains showed different inactivation profiles even within the same genotype. For instance, G1 strains Ku and K8 were more susceptible to HPP than the Wa strain. Overall, the ranking of the stability of RV genotypes to HPP is
G1>G2>G3>G4. These results indicate that the response of different RV strains to HPP is widely different even though they are closely genetically related and have a similar capsid composition. Sequence analysis found that the amino acid homology of capsid proteins among the species A RV strains can range from 70-95%. Perhaps, the nucleotide and amino acid diversity could impact both protein-protein and RNA-protein stabilities, which contribute to the differences in stability under pressure treatment. Also, trypsin is required for rotavirus infectivity. It is possible that trypsin can bind to virions and can be co-purified with the virions which may affect the stability of virus during HPP treatment.
These results suggest that there are some distinct molecular or biologic differences between the strains that led to the differences in stability to HPP.
Previously, it has been documented that viruses within the same family or genus are highly diverse in inactivation profiles by HPP treatment. Coxsackie B5 (Species
Enterovirus B, Genus Enterovirus) and poliovirus (Species Enterovirus B, Genus
Enterovirus), which belong to the family Picornaviridae, are extremely stable at high levels of HPP. Both Coxsackie B5 and poliovirus had less than a 1 log reduction in viral titer after 600 MPa treatments for 5 min at room temperature (Kingsley et al., 2004). In contrast, HAV, a virus in species Enterovirus C in genus Enterovirus, was found to be highly susceptible to HPP compared to Coxsackie B5 and poliovirus (Kingsley et al.,
2004, Grove et al., 2008). A 7 log reduction in HAV was achieved after 450 MPa of
128 treatment at room temperature for 5 min in culture medium (Kingsley et al., 2006).
Human NoV can potentially serve as a good model to study the role of genotype differences in pressure sensitivity because it is highly diverse both genetically and antigenically. However, it has been a challenge to study human NoV as it cannot be grown in cell culture. Recently, a surrogate assay (viral receptor binding assay) was developed to discriminate noninfectious and infectious human NoV particles. Using this assay, it was found that human NoV GI.1 strain was more resistant to the HPP than the
GII.4 strain at 450 MPa at 1°C (Li et al., 2013). In addition, recent evidence suggests that different species in the genus Norovirus within the family Caliciviridae have different pressure sensitivities. Pressure treatment at 400 MPa at 6°C for 5 min completely inactivated murine norovirus, a member of genogroup V within the genus Norovirus.
However, this pressure condition was insufficient to prevent Norwalk virus [genogroup 1 genotype 1 (G.1.1.) within genus Norovirus] infection and shedding in human subjects.
Interestingly, HPP at 600 MPa and 6°C for 5 min was required to completely inactivate
Norwalk virus in seeded oysters, based on the lack of infection and virus shedding in the challenged volunteers (Leon et al., 2011). Overall, the response of different viruses to
HPP appears not to be correlated to virus size, shape, presence of an envelope, family, genera, or genotypes. The large disparity in the resistance of the different viruses may be attributed to the nature of the virus itself, the size and shape of the virus particle, its high thermodynamic stability, differences in viral receptor binding properties, or differences in protein structure, amino acid composition, and isoelectric point. Understanding the continuity or variability of inactivation associated with different viral strains following
129 HPP can aide in the design of appropriate treatment parameters to inactivate diverse viral populations.
3.5.3. Mechanism underlying HPP inactivation of viruses. In this study, we found that disruption of the integrity of the viral capsid but not degradation of viral protein or genomic RNA is the primary mechanism of viral inactivation by HPP. The size of the complete TLP of RV is between 70 to 80 nm, with a round shape. The TLP is composed of three structural proteins; VP7 the outer-capsid protein, VP6 the medial capsid protein, and VP2 the inner capsid protein. VP4, the outer-spike protein, is cleaved by proteases into VP5 and VP8. The DLPs lack VP7 and VP4, and only consist of VP6 and VP2
(Desselberger, 2014). We found that at a low pressure level (200 MPa) the DLPs of both
Wa and SA-11 strains were disrupted, whereas the TLPs remained intact. This indicates that the outer-capsid proteins (VP7 and VP4) stabilize the capsid structure of RV upon
HPP treatment. Elevating the pressure level to 600 MPa resulted in disruption of DLPs and TLPs of both strains and no intact particles were observed.
In general, HPP does not break covalent bonds at the level applied for food processing (up to 800 MPa) (Lou et al., 2011a; Tang et al., 2010). Consistent with this, we found that the structural proteins of both TLPs and DLPs were intact after HPP when they were analyzed by SDS-PAGE. In addition, viral genomic RNA was not physically degraded in RNase free conditions when purified RV was treated by a lethal level of
HPP. However, the abundance of viral RNA was significantly decreased when RV was not treated with an RNase inhibitor. Based on these observations, a model for a mechanism of viral inactivation by HPP is illustrated in figure 24.
130
Figure 24. Models of viral inactivation by HPP. Rotavirus is a small round particle
approximately 60-80 nm in diameter. The native infectious rotavirus virions, termed
triple-layered particles (TLPs), are composed of three concentric layers of proteins and
11 segments of double-stranded RNA. The outermost layer of the virion is comprised of
two proteins, VP4 and VP7. VP4 forms dimeric spikes that project from the surface of
the virion. The middle layer is comprised of VP6. The innermost layer is composed of
three proteins, VP1, VP2, and VP3. After pressure treatment, the structure of the viral
capsids was disrupted whereas the viral capsid proteins and genomic RNA were not
degraded by HPP. The naked viral RNA genome was released from the capsid and
subsequently degraded by exogenous RNase present in food and the environment. 131
After pressure treatment, the structure of the viral capsid was disrupted and the naked viral RNA genome was released from the capsid and subsequently degraded by exogenous RNase present in the food and the environment.
Previously, using human NoV virus-like particles (VLPs) as a model, we found that HPP disrupted the binding capability to histo-blood group antigens (HBGAs), the functional receptor for human NoV (Lou et al., 2012). Collectively, HPP-induced virus inactivation may include disruption of the integrity of the viral capsid and viral receptor binding activity.
3.5.4. Inactivation of RV (Wa strain) in oyster tissues. The consumption of raw oysters is associated with a large number of foodborne virus outbreaks. Shellfish are filter feeders, for instance oysters can circulate around 16 gallon of water per oyster/day and the viral and bacterial concentration in shellfish meat can reach 400 times higher than its levels in the growing water (Keller et al., 2013). HPP can be used to eliminate foodborne pathogens, such as bacteria and viruses, from oysters to enhance safety, reduce labor costs of shucking, and increase shelf life (He et al, 2002, Kingsley et al., 2014, Cruz-
Romero, 2004). Here, we found that HPP treatment at 400 MPa at an initial temperature of either 4 or 20°C was capable of eliminating 4.5-log 10 of RV Wa strain (the most stable RV strain) from the oyster tissues. The initial temperature during treatment did not have a significant impact on the inactivation of the Wa strain in oyster tissues, which was similar to the results obtained when the virus was suspended in medium. This observation, coupled with the fact that both HAV and a human NoV G1.1 strain can be
132 effectively inactivated by HPP at commercially acceptable pressure levels (≤600MPa), suggests that HPP is a highly promising technology to ensure the safety of oysters and other seafood.
The complexity of food matrix has been shown to provide a baroprotective effect to viruses during HPP (Kingsley & Chen, 2008, and 2009; Kingsley et al., 2005; Lou et al., 2011). Specifically, carbohydrates, fats, salts, proteins, ions, and other food constituents can protect viruses from inactivation (Baert et al., 2009; Balny et al., 2002;
Gross & Jaenicke, 1994; Kingsley & Chen, 2008 and 2009). The pressure-treatment of bovine enterovirus (a surrogate for hepatitis A) and feline calicivirus (a surrogate for human norovirus) in shellfish, seawater, and culture medium were found to be most resistant when treated in oysters and mussels (Murchie et al., 2007). Similarly, HAV was more resistant to HPP in oyster homogenates than in 0.3% NaCl solutions at a similar pH.
(Kingsley & Chen, 2009). In this study, it was found that a 4.9-log reduction of the Wa strain was achieved in aqueous medium following 400 MPa treatment for 2 min whereas only a 4.2 log reduction was achieved at the same pressure level in oyster tissues. At 300
MPa, approximately 3.0 log reduction of Wa strain in medium was observed whereas only 2.0 log reduction was achieved in oysters. This result suggests that the food matrix confers protection to RV during HPP treatment. Also, it should be noted that the Wa strain in oysters was either completely inactivated at 400 MPa, at 4° and 20° C or else some natural inhibitors to the infectious Wa strain were present/released from the oyster meat matrix after 400 MPa treatment. Thus, it is necessary to optimize the processing parameters for each product since the efficiency of viral inactivation varies with the food matrix.
133 In conclusion, we demonstrated that (i) RV inactivation by HPP is favored at 4°C compared to 20°C with the exception of Wa strain; (ii) RV strains in different genotypes have different susceptibility to high pressure; and (iii) HPP treatment disrupted the RV virion structure, but did not degrade viral protein or RNA.
134
CHAPTER 4
Resistance of Tulane virus, an enteric primate calicivirus, to thermal and high
pressure processing
4.1. Abstract
In general, microorganisms are capable of acquiring resistance under stress.
Enteric caliciviruses cause acute gastroenteritis in humans or animals. Whether enteric caliciviruses can develop resistance during thermal and nonthermal processing is not known. This study aims to utilize Tulane virus (TV), an enteric primate calicivirus, as a surrogate for human norovirus (HuNoV), to screen for heat and high pressure (HPP) resistant strains. Wild type (WT) TV was exposed to either heat treatment at 70°C for 6 s in culture medium for 5 passages or HPP at 300 MPa for 2 min at 20°C for three passages. Viral plaques were purified after each heat or HPP treatment and were subsequently re-treated or re-pressured and the change in log reduction after each treatment cycle was determined by plaque assay. The results showed that TV was able to develop heat resistance after 5 rounds of heat treatment. The log reduction of WT and the most heat resistant strains was 2.3 and 0.5 log10 PFU/ml, respectively. TV was able to develop HPP resistance after three passages. The log reduction of pressure resistant isolates ranged from 1.7 to 2.0 whereas 3.6 log reductions were achieved for the WT TV after treatment at 300 MPa for 2 min at 20°C. Interestingly, these heat and pressure resistant TVs had smaller plaque size, delayed replication kinetics and cytopathic effects,
135 indicating they were attenuated in cell culture. Subsequently, the full-length genome of these resistant TV mutants was sequenced. Several mutations were observed in the VP1 gene, the virus capsid gene, at T593I and F300S in heat resistant isolates and at S82A,
K163R, and A278G in HPP resistant isolates. In conclusion, this is the first work to characterize a panel of heat and HPP resistant caliciviruses. TV was able to develop heat and HPP resistance, and mutations in VP1 may be associated with the resistance. The finding that enteric virus can develop resistance highlights the need to optimize processing parameters to completely inactivate virus in foods.
4.2. Introduction
It is well known that microorganism can develop resistance in response to environmental stress and therapeutic agents. For example, the use of antibiotics and drugs has led to the prevalence of antimicrobial resistancant bacteria. In the food industry, bacterial pathogens that are resistant to thermal and nonthermal processing technologies have been widely reported. Viruses are obligate intracellular parasites that must utilize a host cell to synthesize their own genetic material and proteins. Whether enteric viruses can develop resistance to processing technologies is relatively less understood. However, virus inactivation kinetics under most mild treatments (such as heat, high pressure processing, etc.) does not follow the log-linear reduction model. Rather, target viral pathogens show tailing effects at the end of the curve (Kingsley et al. 2006; Li et al.
2014; Bozkurt et al., 2015), suggesting that some resistant subgroups in the virus quasi- species may survive the treatment. For example, reoviruses and vesicular stomatitis virus
(VSV), have exhibited the ability to adapt to different types of stress and develop
136 resistance to survive heat treatment and human serum neutralization via mutations to either the outer capsid protein or the envelop glycoprotein (Agosto et al., 2007; Hwang and Schaffer, 2013). Hwang and Schaffer (2013) showed that VSV developed several mutations in the G protein (positions K66T, T368A and E380K) after six heat cycles and additional mutations at S162T, T230N and T368A that enhanced the virus resistance to the human serum (Hwang and Schaffer, 2013). Reovirus subjected to heat treatment showed intragenic pseudoreversions of a double mutation in the M2 gene encoding the outer-capsid protein, µ1. A mutation at D371A was shared in all the mutant particles, and comparing the thermal stability of the D371A mutant to wild type virus revealed that the mutation provides the heat resistance for the mutant viruses. However, this mutation resulted in virus attenuation, resistance to μ1 protein rearrangement in vitro, and loss of the ability to lyse red blood cells. For the mutant particles to restore fitness for growth, another set of mutations was required including 9 different positions in the μ1protein
(Agosto et al., 2007). Moreover, in a survey conducted by Negovetich and Webster
(2010), the thermal stability of 7 isolates of influenza virus (H2N3) was investigated after heat treatment at 37 and 57°C. The results showed that the viral population consisted of two groups, one of which were thermal stable and another that were thermal sensitive.
Increasing the temperature to 57°C resulted in two different inactivation phases, the thermal stable group had inactivation time 14 times higher than the sensitive population
(Negovetich and Webster, 2010).
Structural changes to the viral capsid during thermal inactivation have been linked to functional entry-associated rearrangements in many viruses. For instance, poliovirus converted to the 135S entry intermediate following heat treatment and exposure to
137 hypotonic buffer (Epand and Epand, 2002; Middleton et al., 2007). Epand (2002) studied the effect of heat and pH on untreated influenza virus (X-31 strain) and bromelain-treated virus. The differential scanning calorimetry (DSC) and SDS/PAGE analysis of treated and untreated virus showed that the main effect of the heat on the virus was denaturation of the haemagglutinin (HA) protein. At neutral pH, heat treatment of the virus caused a loss of the HA protein trimer and virus aggregation occurred at temperatures ranging from 55 to 70°C (Epand and Epand, 2002). It is not yet clear whether thermal and non- thermal resistant of viruses occur naturally in viral quasi-species or if it is built up as a response to the selective environment of the treatment.
Although HuNoV and other members (such as human sapovirus) in the family
Caliciviridae cause more than 50% of foodborne outbreaks; there is still limited knowledge about their ability to develop resistance to heat and high pressure processing
(HPP) and the mutations behind the conferred resistance. HuNoV causes severe gastroenteritis characterized by vomiting, diarrhea, and stomach cramps (Atmar et al.,
2008). In the US, it is estimated that HuNoV accounts for more than 60% of foodborne illnesses and it is the second leading cause of gastroenteritis-related mortality, causing
797 deaths annually (Scallan, Hoekstra et al., 2011; Hall et al., 2013). It has been a challenge to study HuNoV because it cannot be grown in a cell culture system and lacks a small animal model. Therefore, cultivable viral surrogates that are closely related to
HuNoV, such as murine norovirus (MNV) and Tulane virus (TV) have been used to study the survival of HuNoVs in foods and the environment (Li et al., 2013; Predmore et al., 2015). Tulane virus (TV) serves as a good model to study the stability of calicivirus capsid under thermal and non-thermal processing. TV is a cultivable primate calicivirus
138 that was isolated from the stool of rhesus macaques with symptoms of gastroenteritis in
2008 (Farkas et al., 2008). TV is a single-stranded, positive-sense RNA virus, with a genome size of 6.714 kb. Similar to HuNoV, the genome of TV contains three open reading frames (ORFs). ORF1 encodes the nonstructural (NS) polyprotein, while ORF2 encodes the major capsid protein, and ORF3 encodes the minor capsid protein (Farkas et al., 2008). TV is a promising surrogate for HuNoV for several reasons. First, TV is an enteric virus similar to HuNoV and it recognizes the authentic receptor for HuNoV, the histo-blood group antigens (HBGAs) (Farkas et al., 2008). In addition, Cryo-Electron
Microscopy (Cryo-EM) results show that the structure and the organization of the TV major capsid protein is similar to HuNoV (Yu et al., 2013).
In our knowledge, this is the first attempt to study the HPP and heat resistance of caliciviruses. The goal of this research is to utilize TV, as a surrogate for HuNoV, to screen for heat and HPP resistant strains; investigate the mutations behind the resistance; and to determine the effect of the mutations on the viral infectivity and fitness for growth.
4.3. Materials and Methods
4.3.1. Cell culture and virus propagation. TV was provided by Dr. Xi Jiang at
Cincinnati Children's Hospital Medical Center, OH and propagated as described by (Li et al., 2013). Briefly, TV was propagated in monkey kidney cells LLC-MK2 (ATCC). The cells were seeded in a T150 flask containing Opti-MEM Reduced Serum medium supplemented with L-glutamine and 2% heat-inactivated fetal bovine serum (FBS) and grown for 24 h at 37°C in presence of 5% CO2. Confluent cells were infected with TV at multiplicity of infection (MOI) of 0.1 and were incubated at 37°C in 5% CO2 atmosphere
139 1 h with agitation, each 10 to 15 min. Eighteen milliliter of Opti-MEM Reduced Serum medium supplemented with L-glutamine and 2% heat-inactivated fetal bovine serum was added to the infected cells and incubated for 48 to 72 h post-infection at 37°C in 5% CO2 atmosphere. The virus was harvested by three freeze-thaw cycles followed by low-speed centrifugation at 1,500 × g for 15 min and kept at -80°C.
4.3.2. TV plaque assays. TV plaque assay was performed as described by (Predmore et al., 2015). In short, after the LLC-MK2 cells were seeded into six-well plates and incubated for 24 h, the cells were washed with Hank’s Balanced Salt Solution (HBSS).
400 μl of the viral inoculum in a 10-fold dilution series were inoculated to the cells and incubated for 1 h at 37°C with agitation every 10 to 15 min. The cells were overlaid with
2.5 ml of 2× EME solution containing 5% FBS, 1% agarose, 1.5 % sodium bicarbonate,
0.1 mg of kanamycin/ml, 0.05 mg of gentamicin/ml, 15 mM HEPES (pH 7.7), and 2 mM
L-glutamine. The six-well plates were incubated for 48 h at 37°C and in 5% CO2 atmosphere. The plates were fixed in 10% formaldehyde, and the plaques were visualized by staining with 0.05% (wt/vol) crystal violet.
4.3.3. Screening for heat resistant isolates. To isolate subpopulations of TV that varied in thermostability, the viral quasi-species of wild type TV in culture medium was heated in capillary tubes (Kimble Chase, 1.5-1.8 × 100 mm) and sealed with a vinyl plastic cover
(Leica Critoseal) at 70 °C for 2, 4, 6, 8, and 10s in a circulating thermostatically controlled water bath. The level of virus survival was determined by standard plaque assay. From the death curve (data not shown), it was found that 6 s lead to a 2.3 log10
140 reduction in TV. These conditions were used to screen for heat resistant genotypes in the
TV quasi-species. Briefly, TV was heat treated at 70°C for 6 s. The surviving viruses were quantified by standard plaque assay. Before removing the overlay media, individual plaques were randomly selected and picked up by filter tip pipette. The selected plaques were classified based on the plaque diameter into four groups and labeled as A, B, C, and
D (A is the smallest and D is largest plaque in diameter). Each selected plaque was passaged in a T25 flask seeded with LLC-MK2 cells for 48 h at 37°C in 5% CO2 atmosphere. The viruses were harvested as described previously. The plaque purified viruses were re-heated (70°C for 6 s) and re-purified for five passages. The survival of each isolate was quantified after each passage by standard plaque assay, and the log reduction was calculated to monitor the development of heat resistance.
Log reduction= Log10 N0 – Log10 N
Where N0 is the initial titer before the heat treatment, N is the viral titer after the heat treatment (Log10 PFU/ml).
4.3.4. Screen for HPP resistant isolates. . Previous studies (Li et al., 2013), have concluded that HPP treatment at a level of 300 MPa at 20°C in neutral pH for 2 min resulted in 3.6 log10 reduction of TV (Li et al., 2013). Thus, 400 µl of TV (WT) was suspended in Opti-MEM medium and double packaged and sealed in sterile polyethylene stomacher pouches (Fisher Scientific International, Ontario, Canada). Samples were subjected to HPP treatment at 300 MPa for 2 min at 20°C using a lab scale HPP unit
(model Avure PT-1; Avure Technologies, Kent, WA) with water as the hydrostatic fluid.
The two minute holding time did not include the come-up and release time (22 MPa/s,
141 and 4 seconds, respectively). The level of survived virus was determined by plaque assay and expressed as log10 PFU/ml. Random plaques were selected and propagated and retreated at the same conditions for three passages as described in Section 4.3.3. The infectious virus particles after each passage were detected by plaque assay using standard plaque assay to screen for HPP resistant strains.
4.3.5. TV plaque purification. After each passage of heat or HPP treatment standard plaque assays were conducted in six-well culture plates of confluent LLC-MK2 cells.
Four plaques were picked from wells and dissolved in 500 μl medium and propagated in a new T25 Flask to start a new passage.
4.3.6. Plaque size determination. The plaque sizes of different TV (WT) isolates after heat or HPP treatment were scanned using a flatbed scanner, and the plaque sizes were detected using GIMP 2.8 software, (http://www.gimp.org).
4.3.7. Single-cycle growth curves of heat and HPP isolates. After five cycles of heat treatment at 70°C for 6 s and three cycles of HPP at 300 MPa for 2 min at 20°C, two heat resistant isolates, three HPP resistant isolates, and TV wide type (WT) were selected to study the kinetics of growth. Confluent LLC-MK2 cells in 6-well plates were infected with individual isolates of TV (WT), heat, or HPP resistant isolates at a MOI of 0.01.
After 1 h of adsorption, the inoculums were removed; the cells were washed twice with
HBSS, and fresh Opti-MEM Reduced Serum medium supplemented with 2% FBS was added the infected plates. The plates were incubated at 37°C in presence of 5% CO2.
142 50µl of the aliquots of the cell culture fluid of each well was withdrawn at different time points, and the virus titer was determined by standard plaque assay.
4.3.8. Determination of the mutations in the heat and HPP treated isolates. To determine any mutations in the viral genome, the genome of TV isolates were amplified by RT-PCR and sequenced. 100 µl of each TV heat or HPP resistant isolate were used to extract the RNA using the RNeasy minikit (Qiagen, Valencia, CA) according to the manufacturer’s manual. One Step Reverse Transcription-PCR (RT-PCR) was used to amplify the genome. The genome of TV was amplified by five overlapping fragments
(Table 10). Based on published nucleotide sequence of TV serial of oligonucleotide primers were used:
143 Table 10: List of primers used for TV sequencing
Name Position Polarity Sequence (5’-3’)
P1-F 0-31 + GGGTGACTAGAGCTATGGATACGTC
P2-R 455-480 - GTATGCCAGGAAAAAGCTACATGGG
P3-F 781-805 + TTTTTAAGAAATCCCTGAATGCTTT
P4-F 1561-1587 + GGGGTTGCAACGGACCAAAGCCATAC
P4-F 2341-2365 + ATGAGTGGTGGGATCCGGATGGTGA
P4-F 3121-3145 + CAAATCCAGGAAGAGTGAACCAAGC
P5-F 3901-3930 + ATGATGAGGTCATCAACTGTCCCTG
P6-F 4781-4706 + TGGCAGGCAATGCCTTCTCTGCTGG
P7-F 6241-6265 + TCTTATGTCTAAGTCCCTCTCTTCA
P8-R 3195-3220 - GCGGACAAATCTATTCGTGTCTTCAC
P9-R 5974-5992 - GCCAGCCATTATCTAAAGA
P10-R 6680-6714 - GTATGCCAGGAAAAAGCTACATGGG
The amplified DNA bands were visualized by 1% agarose gel electrophoresis and purified using QIAquick PCR Purification Kit according to the manufacturer’s manual.
The purified RT-PCR products of the heat and HPP resistant TVs were sequenced at
Plant-Microbe Genomics Facility at The Ohio State University.
4.3.9. Sequence Alignment. Sequencing of TV from GenBank sequence (GenBank:
ACB38132.1) was aligned with the TV heat and HPP resistant isolates using DNASTAR
Lasergene 11 (multiple sequence alignment program).
144 4.3.10. Statistical analysis. All experiments were performed in triplicate. Virus titer was expressed as mean log PFU/ml ± 1 standard deviation. Statistical analysis was performed by one-way multiple comparisons using SPSS 8.0 statistical analysis software (SPSS Inc.,
Chicago, IL). A P value of <0.05 was considered statistically significant.
4.4. Results
4.4.1. Screening for heat resistant isolates. To isolate heat resistant genotypes, TV
(WT) was heated in capillary tubes at 70°C for 6 s in a circulating thermostatically controlled water bath. The level of survived viruses was determined by standard plaque assay. Based on the plaque size, random plaques were selected. The selected plaques were classified based on the plaque diameter into four groups and labeled as A, B, C, and
D (A is the smallest and D is largest plaque in diameter). The heat inactivation of TV
(WT) at 70°C for 6 s showed a 2.3 log10 PFU/ml reduction of the virus titer (Fig.1). This treatment was called P0. Each isolated plaque from P0 was cultivated in LLC-MK2 cells and survived viruses were reheated for five passages at 70°C for 6 s (Fig.25).
At P1 to P4 (from the first to forth cycle of heat treatmen) there were no significant differences (P>0.05) in the log reduction among or within the four isolates (A,
B, C, and D). The average of log reduction ranged between 2.2 to 1.8 logs. At P5, the four isolates showed different log reduction levels. The isolates with the small plaque sizes, A and B, were significantly more heat resistant (lower log reduction) compared to the WT and the isolates of larger plaque sizes, C and D (P<0.05).
145
Figure 25. Plaque sizes and log reduction of WT TV after heat treatment at 70˚C for
6 s. (A) Four plague sizes were selected to screen for heat resistant strains labeled as A
the smallest and D is the largest in sized. (B) Log reduction of WT TV was determined
by plaque assay and by subtracting the number of surviving particles (log10 PFU/ml)
after heat treatment from the number of untreated. The error bar is 1 times the standard
deviation.
The log reduction of the four isolates were 1.16, 0.97, 1.92, and 1.44 logs for A,
B, C, and D isolates, respectively. The results may indicate that some of the virus population had developed heat resistance (Figure 26). To further investigate heat resistant genotypes the four isolates further heated at 70°C for 6 s and random plaques of each isolate were selected, grown, and heat treated for another cycle.
146
Figure 26. Log reduction (log10 PFU/ml) of TV heat-resistant variants after heat treatment at 70˚C for 6s for 5 passages. Log reduction was determined by subtracting the number of surviving particles (log10 PFU/ml) after heat treatment from the number of
untreated. The error bar is 1 times the standard deviation.
The survived viruses were quantified by plaque assay and the log reduction was calculated (Fig. 27). The average log reduction of the isolates ranged between 0.53 to 2.3 log10 PFU/ml. Two isolates (named B5-2 and C5-2) were selected for further analysis by studying the growth kinetics and sequencing.
147
Figure 27. Log reduction (log10 PFU/ml) of TV heat-resistant variants from passage
5 after heat treatment at 70˚C for 6 s. Log reduction was determined by subtracting the
number of surviving particles (log10 PFU/ml) after heat treatment from the number of
untreated. The error bar is 1 times the standard deviation.
4.4.2. Screen for HPP resistant isolates. To isolate HPP resistant phenotypes, TV (WT) was treated under HPP at 300 MPa for 2 min at 20°C. The survived viruses were determined by standard plaque assay. Based on the plaque size, random plaques were selected following treatment. The selected plaques were classified based on the plaque diameter into four groups and labeled as A, B, C, and D (A is the smallest and D is 148 largest plaque in diameter). The HPP inactivation of TV (WT) at 300 MPa for 2 min at
20°C showed a 3.6 log10 PFU/ml reduction in virus titer (Fig. 28). This treatment was called P0.
Figure 28. Log reduction in TV titer after HPP treatment at 300 MPa for 2 min at
20˚C. Reduction was determined by subtracting the number of surviving particles (log10
PFU/ml) after the HPP from the number of untreated. The error bar is 1 times the
standard deviation.
Each isolated plaque from P0 was cultivated in LLC-MK2 cells and survived viruses were treated by HPP at 300 MPa for 2 min at 20°C for three passages (Fig.29). At
149 P1 and P2, one of the isolates (with plaque size A) grew to a lower titer and was below the detection limit following pressure treatment. Therefore, we increased the initial inoculum titer and treated this strain again. At passage three (P3), a decrease in the log10 reduction was observed for the all three isolates. The average of log10 reduction ranged between 2.0 to 1.8 logs, which were significantly different compared to the log10 reduction for WT TV (P<0.05). The results may indicate that some of the virus population had developed a HPP resistance. Three plaques at P3 (Ap3, Bp3, and Cp3) were selected for further analysis.
Figure 29. Log reduction (log10 PFU/ml) of TV HPP-resistant variants after HPP at
300 MPa for 2 min at 20˚C for 3 passages. Log reduction was determined by subtracting the number of surviving particles (log10 PFU/ml) after HPP treatment at 3oo
MPa for 2 min at 20˚C for three passages. The error bar is 1 times the standard deviation. 150 4.4.3. Single-cycle growth curves of heat and HPP isolates. The replication kinetics of three HPP resistant isolates Ap3, Bp3, and Cp3, two heat resistant isolates B5-2 and C5-
2, and TV (WT) was examined in LLC-MK2 cells. Briefly, LLC-MK2 cells in 6-well plates were infected with either TV (WT), HPP resistant isolates, or heat resistant isolates a MOI of 0.01 and incubated for 1 h. The inoculum was removed and fresh Opti-MEM medium containing 2% FBS were added to each well. At the indicated time points, 50 µl of the supernatant of the infected cells was withdrawn and virus titer was determined by the plaque assay. The virus titer of each sample (Log10 PFU/ml) was plotted against the incubation time (fig. 30).
As it shown in figure 30. , B5-2 was significantly delayed in in the viral replication compared to the TV (WT) and the other isolates (P<0.05). After 48 h postinfection, B5-2 titer reached 5.1 log10 PFU/ml, which is 1.4 log10 lower than the titer of TV (WT). However, Ap3, Bp3, Cp3, and C5-2 isolates showed a delay in the early stage of the growth curve although they reached a peak titer similar to TV (WT) after 48 h postinfection.
151
Figure 30. Growth kinetics of TV wild type and five isolates of the most heat and
HPP resistant genotypes. HPP isolates are the high pressure resistant genotypes after 5
passages of HPP at 300 MPa for 2 min at 20˚C. Heat isolates are the heat resistant
genotypes after 5 passages of heat treatment at 70˚C for 6s. The growth curves was in
LLC-MK2 cells at a multiplicity of infection (MOI) of 0.01.
The progression of the cytopathic effect (CPE) was tested for all strains (Fig. 31).
Mild CPE in LLC-MK2 cells was observed within 24 h by TV (WT), and extensive CPE after 48 h of incubation. There was no clear CPE or change in LLC-MK2 cell
152 morphology caused by the B5-2 isolate, the heat resistant isolate, at 24 h post infection, while mild CPE was noticed after 48 h of infection.
Figure 31. The infectivity of TV (WT) and two isolates from heat resistant, and
three HPP resistant genotypes in LLC-MK2 cells at MOI 0.01. The images were
taken 0, 16, 24, and 36 h of post infection. WT is TV wild type, B5-2 is the most heat resistant strain, and C5-2 is a heat stable strain. Cp3, Bp3, and Ap3 is HPP stable strains.
The data confirm the lack of growth fitness for the heat stable strain and delay in growth
of some HPP resistant strains. 153 The B5-2 isolate replication was delayed by 24 h and infection was less severe after 48 h compared to WT. Ap3, Bp3, Cp3, and C5-2 showed less CPE at 24 h compared to the WT, whereas after 48 h these isolates showed the same extensive CPE as the WT.
The results indicate that the heat resistance of B5-2 strain resulted in this isolate being significantly attenuated in cell culture whereas all other TV mutants had a moderate delay in viral replication.
4.4.4. Plaque size determination. The plaque sizes of different TV isolates after heat or
HPP treatment were scanned using a flatbed scanner, and the plaque sizes were measured using GIMP 2.8 software. As shown in Figure 32, a mixed population of plaques was observed at P1.
154
Figure 32. Plaque size of TV (WT) and two isolates from heat resistant, and three
HPP resistant genotypes in LLC-MK2 cells after incubation for 48 h. WT is TV wild type, B5-2 is the most heat resistant strain, and C5-2 is a heat stable strain. Cp3, Bp3, and
Ap3 is HPP stable strains. The data confirm the lack of growth fitness for the heat stable
strain and delay in growth of some HPP resistant strains. The plates was fixed by 10%
formaldehyde and stained by0.05% crystal violet.
The plaque diameters were between 5 and 0.2 mm for P1. For heat resistant TVs at P6, the plaque sizes of different isolates ranged between 1 ± 0.2 mm and 2 ± 0.2 For
155 pressure resistant TVs at P3, the plaque sizes of different isolates ranged between 5 ± 0.2 and 1 ± 0.2 mm.
4.4.5. Localization of mutations in heat and pressure resistant TVs: To investigate the presence of mutations in the TV genome, RNA of two heat and two HPP resistant isolates were extracted and the whole genome was amplified by RT-PCR using five overlapping fragments, and sequenced. The amino acid sequence of the nonstructural protein, major capsid protein (VP1), and minor capsid protein (VP2) protein of each resistant isolates were aligned with WT TV and the amino acid changes of each isolate is shown in Table
11. The majority of the amino acid changes were observed in the VP1 and VP2 genes of the isolates.
In heat stable isolates (B5-2 and C5-2), mutations in the VP1 gene were observed at positions F300S and T593I, indicating these mutations were associated with the heat resistance. Interestingly, mutations at positions I451M and R452C in VP1 were present in both HPP resistant isolates and also in the heat stable isolates C5-2, which may mean they contribute to both heat and HPP stability. Some resistant isolates had additional mutations. For example, a mutation was observed in VP1 at E7D in heat stable isolate
B5-2, the HPP resistant isolate Bp3 had a mutation at S82A, and HPP resistnat Cp5 had two additional mutations at K163R and A278G.
All of the heat and HPP resistant isolates had the same mutations in the VP2 gene.
The position of these mutations were at E164G, T182A, L185S, and Y208H. This indicates that these mutations induce changes that enhance both the thermal and HPP resistance.In addition, some mutations were also found in N-terminal domain of the
156 nonstructural protein, protease, and polymerase protein. For instance, the mutations at position Y107H and S127P in N-terminal domain of the nonstructural protein was found in all heat and HPP resistant isolates, indicating that they may contrubute to both thermal and HPP stability. Mutations were also found in the RNA dependent RNA polymerase
(RdRp) gene at positions V446A and G569E for both thermal resistant isolates. In addition a mutation was found in the 3C protease gene at position S99C only for the thermal resistant isolates. These mutations may also contribute to the increased thermal stability (Table 11).
157 Table 11: Localization of mutations in genome of heat and pressure resistant
TVs
TV protein B5-2 C5-2 Bp3 Cp5
5’ NTR ND ND ND ND
N-terminal Y107H, S127P Y107H, S127P Y107H, S127P Y107H, S127P
NTPase ND ND ND ND
VPg ND ND ND ND
Pro S99C ND ND ND
Pol V446A, G569E V446A, G569E ND ND
K163R, E7D, F300S F300S, I451M, S82A, I451M, VP1 A278G, I451M, T593I R452C, T593I R452C R452C
E164G, E164G, T182A, E164G, T182A, E164G, T182A, VP2 T182A, L185S, L185S, Y208H L185S, Y208H L185S, Y208H Y208H
3’ NTR ND ND ND ND
4.6. Discussion:
HuNoV is the leading cause of gastroenteritis worldwide and it is responsible for more than 50% of food outbreaks (Scallan, Hoekstra et al., 2011; Verhoef et al., 2013).
Due to the inability to grow HuNoV in cell culture and the lack of a suitable animal model for HuNoV, surrogates such as murine norovirus (MNV) and TV are wildly used 158 to study HuNoV inactivation (Bozkurt et al., 2011; Bozkurt et al., 2015). HPP and heat pasteurization are the most common approaches to inactivate enteric viruses in foods
(Kingsley, 2013; Bozkurt et al., 2014). However, viruses have been shown to have dynamic responses due to thermal treatment and this likely due to the presence of resistant mutants present within the viral quasi-species (Agosto et al., 2007; Middleton et al., 2007). Each viral population contains subgroups/quasispecies that have different thermal resistant properties (Hwang and Schaffer, 2013). Providing insight into this phenomenon is essential in order to design effective thermal and non-thermal inactivation approaches.
The thermal stability and HPP resistance of bacterial species has been studied, however, the heat and HPP resistance of caliciviruses remains poorly understood. In this study, we used TV as a model to study the potential of caliciviruses to develop thermal and HPP resistance. The data showed that TV can develop resistance to both heat and
HPP treatments. It appears that TV develops resistance to HPP more quickly after exposure compared to heat treatment. TV developed significantly enhanced resistance to
HPP following three passages whereas significantly enhance resistance was developed following 5 passages of heat treatment.
4.6.1. Heat resistance: A mild treatment temperature of 70°C was applied to observe a significant reduction in virus titer while maintaining a nonlinear decrease in the virus population. The data suggested that the TV population contained genotypes that were heat stable and other heat sensitive genotypes. For example after five passages at 70°C for 6 s the TV isolates showed significant decreases in the log reduction (i.e. increase in
159 resistance) achieved compared with the wide type. The log reduction decreased from 2.3 at the first passage to 0.9 log10 PFU/ml at passage 5. Plaque purification of the resistant isolates from passage 5 prior to another round of heat treatment (passage 6) revealed variation in heat stability even for plaque purified viruses obtained from the same isolate from P5. For instance, B5-2 showed the highest heat stability (0.5 log10 reduction) compared to the wide type (2.3 log10 reduction). In general, it has been shown that the rapid replication/lack of RNA proofreading mechanisms leads to a mutation rate in RNA viruses and this produces a population of mixed viral genotypes (quasi-species). The presence of these quasi-species leads to viral populations with increased stability under environmental stress. Quasi-species also lead to a viral population in which there is variation between the growth kinetics of different mutants within the population
(Scholtissek, 1985; Comas et al., 2005; Negovetich and Webster, 2010). We observed variations in the heat resistance of the plaque purified viruses after 6 passages, with approximately 1.8 logs of variation between viruses. It is likely that more resistant populations could be selected for by increasing the magnitude of the stress or the number of passages under the stress (Brown et al., 2009; Negovetich and Webster, 2010). On the other hand, for the heat resistant viruses the mutations that confer heat resistance may also decrease the ability of the virus to grow well. This decrease in viral growth fitness may lead to an over-estimation of the viral inactivation by heat, leading to variation in the log reductions detected (Agosto et al., 2007).
4.6.2. HPP resistance: HPP is widely used in the food industry and for vaccine production to inactivate pathogenic bacteria and viruses (Hirneisen and Kniel, 2013; Ye
160 et al., 2014). However, limited data is currently available concerning the potential of viruses to develop HHP resistance. Caliciviruses, the enteric viruses most associated with foodborne outbreaks, have not been studied for the ability to develop HPP resistance.
This study aimed to identify HPP stable isolates using TV as a model. WT TV was exposed to pressure treatment at 300 MPa for 2 min at 20°C to achieve a significant reduction in virus titer and maintain a nonlinear decrease in the virus population. This treatment reduced the virus titer by 3.6 logs, while subgroups of the virus population survived (Li et al., 2013). This data suggests that the TV population contains isolates that are HPP stable within the virus population. For example, after 3 passages at 300 MPa for
2 min at 20°C the TV isolates showed a significant decrease in the log reduction compared with WT. The log reduction decreased from 3.6 in the first passage to 1.7, 1.8,
2.0 log10 PFU/ml at passage 3 for the D, B, A, and C isolates, respectively. Huang et al.
(2014) studied the effect of pressure cycling on MNV-1 inactivation inoculated in fruit puree samples at 300 MPa and 0°C. The results showed that there was no further reduction in the virus titer after the first cycle of HPP of the survived viruses (Huang et al., 2014). Ferreira et al. (2009) reported that HPP treatment enhanced the heat stability of the attenuated poliovirus vaccine at 37°C (Ferreira et al., 2009). In our study, the decrease in log reduction between the three isolates was not significantly different at passage 3. More differences may be observed by increasing the pressure level or the number of passages under the same pressure. Overall, our results demonstrated that TV can quickly develop resistance to HPP treatment.
161 4.6.3. Phenotype of heat and pressure-resistant TV mutants. The heat and pressure resistant TV mutants exhibited differing phenotypes compared to WT. In terms of heat resistance, the mutant B5-2 was the most attenuated strain. After 24 h, when 80-90% of the LLC-MK2 cells showed extensive CPE, the titer of B5-2 was approximately 3 log10
PFU/ml compared to 7 log10 PFU/ml for the WT virus. This may indicate the most heat resistant genotypes in the viral quasi-species are present at a level of 1 to 10,000 in the viral population. In addition, B5-2 virus had significantly smaller plaque size and delayed replication kinetics compared to WT. This result suggests that the mutations that conferred heat resistance also cause defects in the virus’s ability to replicate in cell culture.
The other HPP and heat resistant isolates grew to the same titer as WT at 24 and
48 h postinfection. However, they showed a delay in replication in the first 16 h of infection. This delay in viral replication may reduce the final titer these resistant isolates can grow to when competing in mixed population with the quick growing WT.
Negovetich et al. (2010) showed that a population of H2N3 influenza virus contained two different heat sensitive subgroups. The selection experiments and plaque purification revealed that 0.1% of the H2N3 influenza virus population was composed of heat stable genotypes (Negovetich and Webster, 2010). Enteric caliciviruses typically cause gastroenteritis in humans and animals. Currently, there is no vaccine available for these viruses. To our knowledge, an attenuated strain of an enteric calicivirus has not been reported. Identification of an attenuated calicivirus strain will facilitate the development of live attenuated vaccines to prevent enteric calicivirus infection. Given the fact that some of the heat and pressure resistant TVs (particularly B5-2) are highly defective in
162 replication in cell culture, it is likely that they will also be attenuated in primates, the natural host of TV. From this perspective, these heat and pressure resistant TVs may not only lead to the development of live attenuated vaccines but also will be highly useful to investigate the pathology, immunology, and biology of enteric caliciviruses.
4.6.4. Genetic changes of the heat and HPP stable strains. The enhanced thermal and
HPP resistance of the selected isolates of TV must be attributed to changes in the viral genetic material, the single-stranded positive-sense RNA genome. Genomic sequencing of the selected TV mutants identified multiple mutations in the viral genome including mutations located in the major capsid protein gene. Previously it has been shown that viruses with enhanced stress tolerance had mutations in the outer viral capsid or glycoprotein genes. For example, the mutation responsible for reovirus heat resistant phenotypes was located in the outer capsid proteins (Agosto et al., 2007, Hwang and
Schaffer, 2013). Viral mutants that escaped antibody neutralization had mutations that lead to changes in the viral surface glycoproteins (Hwang and Schaffer, 2013). In this study, it is not known whether a particular mutation of a combination of mutations lead to the enhanced tolerance of TV to heat and HPP treatment. Interestingly, the majority of the mutations observed in the resistant isolates were located in the major capsid protein
(VP1) and the minor capsid protein (VP2), although mutations were also found in polymerase and N-terminal domain of the nonstructural protein. A few mutations located in the N-terminal domain and VP2 was observed in both the heat and HPP resistant isolates. This suggests the same mutations contribute to both heat and HPP resistance. It is likely that many of these mutations will also be responsible for reducing the growth
163 fitness of the heat and HPP tolerant strains. The heat and HPP resistant isolates had a smaller plaque size and reduced infectivity in LLC-MK2 cells compared to WT. This may be due to changes in the virus capsid that affected the virus attachment and release from the cell. However, the change in plaque size may be independent from the virus stability to heat and HPP (Negovetich and Webster, 2010).
In conclusion, this is the first work to characterize a panel of enteric caliciviruses that are resistant to heat and HPP. TV was able to quickly develop heat and HPP resistance, and mutations in viral genome may be associated with that resistance.
Interestingly, two shared groups of mutations are responsible for two different types (heat and pressure) of resistance. Also, these heat and pressure resistant TVs are attenuated in cell culture. Further studies are needed to determine whether these TV mutants were also attenuated in nonhuman primates and to evaluate whether these mutants can be used as live attenuated vaccine candidates. In addition, future studies to evaluate the role of these mutations in conferring resistance are required to design critical parameters for thermal and non-thermal processing. Practically, complete inactivation of the virus in food is critical because it eliminates the possibility of developing resistance.
164
CHAPTER 5
Conclusions and future directions.
5.1. Conclusions
5.1.1. This study demonstrates that major enteric viruses including human norovirus
(HuNoV) and its surrogates (murine norovirus, MNV-1; and Tulane virus, TV), hepatitis
A virus (HAV), and human rotavirus (RV) can be efficiently bio-accumulated in all oyster tissues within 72 hours. However, the pattern of the bioaccumulation substantially varied for each individual virus. Caliciviruses and HAV were localized in the stomach at a high level within the first 24 h whereas RV was first bio-accumulated in gills and disseminated to other tissues.
5.1.2. This study demonstrates that enteric viruses have different heat inactivation profiles. Thus study directly compared the heat stability of HAV, RV, TV, and MNV-1 in cell culture medium and oyster tissues. The four viruses can be ranked from the most heat resistant to the least stable as the following: HAV>RV>TV>MNV-1. Heat treatment at
80°C for 12 s was sufficient to completely inactivate all four viruses in an aqueous medium. However, more than 6 min was required to inactivate HAV and 4 min was needed to inactivate RV in oysters, demonstrating that oyster tissues provide protective effects on virus inactivation.
165 5.1.3. The research demonstrates that the primary mechanism underlying heat inactivation of viruses is the disruption of the integrity of the viral capsid but not degradation of the viral genome. A dose of heat that was lethal to the virus was not sufficient to inhibit viral receptor binding ability, suggesting that heat may also disrupt the capsid-genome interaction prior to the damage of receptor binding.
5.1.4. This study demonstrates for the first time that virus strains derived from different serotypes with the same species have different baro-sensitivity to high pressure processing. Specifically, RV serotype G1 (Ku and K8 strains), G2 (S2 strain), G3 (SA-11 and YO strains) and G4 (ST3 strain) showed enhanced inactivation at 4°C compared to
20°C. In contrast, G1 serotype Wa strain was not significantly impacted by the initial treatment temperature. Within serotype G1, Wa stain was significantly (p<0.05) more resistant to HPP compared to Ku and K8. Overall, the resistance of the human RV strains to HPP at 4°C can be ranked as Wa>Ku=K8>S2>YO>ST3 and in terms of serotype
G1>G2>G3>G4.
5.1.5. This study demonstrates that disruption of the integrity of viral capsid but not degradation of viral genomic RNA is the primary mechanism of high pressure-induced virus inactivation.
5.1.6. This study demonstrates for the first time that TV, an enteric primate calicivirus, easily develops resistance under both heat and high pressure treatment. These heat and pressure resistant TVs showed 2-3 logs more resistance to heat or pressure compared to
166 wild type TV. Consistent with the resistance phenotype, mutations were identified in the genome of these resistant TV strains. However, mutations responsible for heat resistance are different with those responsible for pressure resistance.
5.1.7. This study found for the first time that heat and pressure resistant TVs were attenuated in cell culture. Specifically, these heat and pressure resistant TVs had delayed replication kinetics, delayed cytopathic effects, and had diminished plaque size in LLC-
MK2 cells compared to wild type TV.
5.1.8. This study demonstrates that thermal heat and high pressure processing are capable of effectively inactivating major foodborne viruses by optimizing processing parameters.
The finding that virus can develop resistance highlights the need to completely inactivate virus during food processing. Implementation of these optimized processing parameters in food industry will limit shellfish associated virus outbreaks and thus improve food safety and public health.
5.2. Future directions:
This study systemically determined the effectiveness of the two most common processing technologies, heat and HPP, in inactivating foodborne viruses in cell culture medium and oysters. At the optimized processing conditions, both technologies were capable of effectively eliminating viruses from oysters without significantly altering the organoleptic properties of oyster tissues. This study also determined the mechanisms of viral inactivation by heat and HPP and found that both processing technologies disrupted
167 the integrity of viral capsid. In addition, different viruses within the same species had significant differences in response to heat and high pressure. Finally, it was found that a primate calicivirus easily developed resistance to both heat and high pressure. The findings of this study will facilitate the development of effective technologies to eliminate virus hazards in oysters. However, there many questions remain answered.
5.2.1. Determine whether receptors are involved in the bioaccumulation of enteric viruses in oysters. In this study, we found that different viruses had different distribution patterns in oyster tissues. The mechanism behind this is not known. One possibility is that viral receptor binding activity plays a role in this difference. Enteric viruses either utilize carbohydrates and/or proteins as the receptors. Interestingly, these receptor-like molecules are highly abundant in oyster tissues. For example, HuNoV utilizes histo blood group antigens (HBGAs) as the receptors. It was found that HBGA-like molecules exist in oyster tissues, and HuNoV can specifically bind to these HBGA-like molecules. For another example, MNV-1 uses sialic acid as a receptor, a type of carbohydrate moiety that is also abundant in oysters. Future studies should aim to determine whether a virus- receptor interaction plays a role in virus bioaccumulation in oysters.
5.2.2. Determine the survival of human norovirus (HuNoV) in foods. It remains a challenge to determine the survival of HuNoV because it lacks a robust cell culture system. All viruses must bind to a cellular receptor to initiate an infection. This disruption of viral receptor binding activity will likely be lethal to the virus. Using this theory, the
PGM-MB assay, a receptor binding-based assay was developed to estimate the survival
168 of HuNoV. In fact, this PGM-MB assay worked very well for estimation of the survival of HuNoV treated by high pressure processing (HPP). Previously, our laboratory showed that pressure treatment of HuNoV-contaminated oysters at 350 MPa at 0° C for 2 min resulted in 4.0 log virus reduction estimated by PGM-MB assay (Lou et al., 2015). This treatment condition was sufficient to prevent gnotobiotic piglets from an infection (Lou et al., 2015). Thus, the log reduction obtained from PGM-MB assay was consistent with the in vivo infectivity assay in gnotobiotic piglets. Unfortunately, this study found that the
PGM-MB assay was not suitable for determining the survival of HuNoV treated by heat.
This is probably due to the difference in the mechanism of viral inactivation by heat and high pressure. High pressure efficiently disrupted the receptor binding activity whereas heat did not efficiently disrupt the receptor binding. In the gnotobiotic pig model can be used to verify the survival of HuNoV treated by heat.
5.2.3. Is human B cell culture system robust enough for determination of HuNoV survival in foods? Recently, Dr. Karst’s group at University of Florida found that HuNoV GII.4-
Sydney strain can replicate in human B cells in the presence of enteric bacteria
(Enterobacter cloacae) expressing histo-blood group antigens (HBGAs), the functional receptor for HuNoV. This paper was published in Science (Jones et al., 2014). They observed an increase of 1-2 log RNA copies of HuNoV in this culture system. However, no standard plaque assay or TCID50 assay was reported, raising the question whether this system is robust enough to determine the survival of HuNoV. To test this, HuNoV stocks will be treated by heat or other technologies, and will be inoculated onto human B cells in
169 the presence of HBGA-expressing enteric bacteria. The increase of genomic RNA copies may be used as an indicator for survival.
5.2.4. Determine the mechanism behind the differences in baro-sensitivity of RV strains.
This study found for the first time that the response of different RV strains from either same serotype or species to HPP is widely different even though they are closely genetically related and have a similar capsid composition. The mechanism underlying this observation is not known. There is broad thought that the nucleotide and amino acid diversity could impact both protein-protein and RNA-protein stabilities, which contribute the different stability under pressure treatment. Understanding this question will need to develop a reverse genetics system for RV that allows us to analyze the individual gene, segment, or mutation in the difference in pressure sensitivity. Unfortunately, the reverse genetics system for RV has not been successful although this system has been recently reported for another double stranded RNA virus, reovirus.
5.2.5. Determine the roles of individual mutations in developing heat and pressure resistance: This work showed that TV easily developed a heat and high pressure resistance. Consistent with this, a number of mutations were found in the genomes of these TV mutants. However, it is not known which amino acid substitutions are responsible for the resistant phenotype. One of future directions is to use a reverse genetics system to determine the role of individual amino acids in developing resistance.
Recently, a TV reverse genetics system, which allows for the recovery of viral mutants from an infectious cDNA clone, has been reported. Using this system, each amino acid
170 change identified in this study can be introduced into an infectious cDNA clone, and recombinant TV carrying a specific mutation can be recovered. Subsequently, we can analyze whether these TV mutants are resistant to heat or pressure. It will be highly interesting to determine the specific gene or specific amino acids that are responsible for the resistant phenotype.
5.2.6. Determine whether heat and pressure resistant TVs are attenuated in primates and can be used as a live attenuated vaccine for TV. This study found that heat and pressure resistant TV mutants were attenuated in cell culture. They were defective in viral replication kinetics, had delayed cytopathic effect, and had significantly smaller plaques.
It is not known whether these TV mutants were also attenuated in primates, the natural host of TV. If these TVs are attenuated in monkeys, they will be highly promising live attenuated vaccine candidates. It is known that TV causes diarrhea in primates. An attenuated TV mutant will not cause enteric infection, thus can be used as a vaccine. To our knowledge, an attenuated strain of calicivirus has not been reported. These heat and pressure TV strains will be highly useful for studying the pathology, immunology, and biology of caliciviruses.
171
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