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Faculty of Bioscience Engineering Academic year 2015 – 2016 Viral-bacterial interactions: The effect of histo-blood group antigen (HBGA)-expressing bacteria and probiotics on Noroviruses Dries Loncke Promotor: Prof. Dr. Ir. Mieke Uyttendaele Tutor: Dr. Dan Li Master’s dissertation submitted in partial fulfillment of the requirements for the degree of Master of Science in Bioscience Engineering: Cell and Gene Biotechnology Faculty of Bioscience Engineering Academic year 2015 – 2016 Viral-bacterial interactions: The effect of histo-blood group antigen (HBGA)-expressing bacteria and probiotics on Noroviruses Dries Loncke Promotor: Prof. Dr. Ir. Mieke Uyttendaele Tutor: Dr. Dan Li Master’s dissertation submitted in partial fulfillment of the requirements for the degree of Master of Science in Bioscience Engineering: Cell and Gene Biotechnology Acknowledgements While writing this thesis, it struck me how fast these past five years of studying Bioengineering went by. Looking back at this period really convinces me that I made the right decision choosing this path. I specialized in Cell and Gene Biotechnology, making the choice for a thesis concerning bacteria and viruses a rather logic decision. However, this experience would not have been possible without the help and support of certain people. In the first place, I would like to thank Prof. Dr. Ir. Mieke Uyttendaele for offering me the chance to work on the subject of viral-bacterial interactions as a master’s thesis as well as giving me advice on how to improve this thesis. Special thanks go out to my tutor Dan. She guided me along this journey, answered all my questions, read and corrected this thesis report, taught me experimental techniques and procedures, helped me explain the obtained results, and made me a better bioengineer. I really appreciate everything you have done for me! I enjoyed working at the Department of Food Safety and Food Quality of Ghent University. Thank you to Ann, Danny, and all the others in and around the lab for creating a nice working atmosphere. To my fellow thesis students, it was fun having you guys around! Last but not least, I want to express my gratitude towards my parents, brother and friends for always supporting me. They have been there for me during the fun and a bit less fun moments, not only this last year, but throughout my whole academic adventure. i ii Permission for use of content The author and the promotor give the permission to use this thesis for consultation and to copy parts of it for personal use. Every other use is subjected to the copyright laws, more specifically the source must be extensively specified when using results from this thesis. Ghent, June 2016 Toelating tot bruikleen De auteur en promotor geven de toelating deze scriptie voor consultatie beschikbaar te stellen en delen ervan te kopiëren voor persoonlijk gebruik. Elk ander gebruik valt onder de beperkingen van het auteursrecht, in het bijzonder met betrekking tot de verplichting de bron te vermelden bij het aanhalen van resultaten uit deze scriptie. Gent, juni 2016 The author, The promotor, De auteur, De promotor, Dries Loncke Prof. dr. ir. Mieke Uyttendaele iii iv Table of Contents Acknowledgements .................................................................................................................................. i Permission for use of content .................................................................................................................iii Toelating tot bruikleen ............................................................................................................................iii Table of Contents .....................................................................................................................................v List of Abbreviations ............................................................................................................................... vii Abstract ................................................................................................................................................... ix Samenvatting........................................................................................................................................... xi Chapter 1: Introduction ........................................................................................................................... 1 Chapter 2: Literature study ..................................................................................................................... 3 2.1 Introduction to Noroviruses ........................................................................................................ 3 2.1.1 Classification and taxonomy ................................................................................................. 3 2.1.2 Structure ............................................................................................................................... 4 2.1.3 Clinical symptoms and treatment ........................................................................................ 6 2.1.4 Transmission routes and prevalence .................................................................................... 7 2.1.5 Pathogenesis......................................................................................................................... 9 2.1.6 Non-cultivability and surrogates ........................................................................................ 10 2.2 Interactions between bacteria and viruses ............................................................................... 12 2.2.1 Types of interactions .......................................................................................................... 12 2.2.2 Histo-blood group antigen expressing bacteria ................................................................. 14 2.2.3 Probiotics ............................................................................................................................ 15 Chapter 3: Material and Methods ......................................................................................................... 21 3.1 Interaction between Norovirus and HBGA expressing bacteria ................................................ 21 3.1.1 Bacteria ............................................................................................................................... 22 3.1.2 Virus-like particles (VLPs) and antibodies........................................................................... 22 3.1.3 Monitoring of the bacterial growth .................................................................................... 22 3.1.4 HBGA expression test ......................................................................................................... 23 3.1.5 Direct ELISA ......................................................................................................................... 24 3.1.6 Mucin-binding ELISA ........................................................................................................... 24 3.2 Interaction between Norovirus and probiotic bacteria............................................................. 25 3.2.1 Bacteria and cell lines ......................................................................................................... 25 3.2.2 Viruses ................................................................................................................................ 26 3.2.3 Monitoring of the bacterial growth .................................................................................... 26 3.2.4 Culturing of the macrophage cell line RAW 264.7.............................................................. 26 v 3.2.5 Plaque assay ....................................................................................................................... 26 3.3 Statistical analysis ...................................................................................................................... 27 Chapter 4: Results ................................................................................................................................. 29 4.1 Interaction between Norovirus and HBGA expressing bacteria ................................................ 29 4.1.1 Optimisation of the HBGA expression test ......................................................................... 29 4.1.1.1 Concentration of antibodies ........................................................................................ 29 4.1.1.2 Concentration of blocking reagent .............................................................................. 30 4.1.1.3 Cell numbers of the tested bacteria ............................................................................ 30 4.1.2 Influence of gas atmosphere of bacterial growth on their HBGA expression and viral protective effects .......................................................................................................................... 31 4.1.2.1 Growth of E. coli LMG 8223 at different atmospheres ............................................... 31 4.1.2.2 HBGA expression of E. coli LMG 8223 and LFMFP 861 grown at different atmospheres ................................................................................................................................................... 33 4.1.2.3 Viral protective effects of E. coli LMG 8223 and LFMFP 861 grown at different atmospheres .............................................................................................................................. 33 A) Direct ELISA ..................................................................................................................... 33 B) Mucin-binding ELISA ........................................................................................................ 34 4.2 Interaction between Norovirus and probiotic bacteria............................................................. 35 4.2.1 B. longum growth ............................................................................................................... 35 4.2.1.1 Growth in bacteria culture medium TSB ..................................................................... 35 4.2.1.2 Growth in a food matrix .............................................................................................. 36 4.2.2 Effect of B. longum on NoVs ............................................................................................... 36 Chapter 5: Discussion ............................................................................................................................ 41 5.1 Interaction between Norovirus and HBGA expressing bacteria................................................ 41 5.2 Interaction between Norovirus and probiotic bacteria............................................................. 43 Chapter 6: General Conclusions ............................................................................................................ 47 References ............................................................................................................................................. 49 vi List of Abbreviations ABO Blood group system ATCC American Type Culture Collection BCCM Belgian Coordinated Collection of Microorganisms BCS Bacterial cell suspension BGMT Bacterial growth medium filtrate BSA Bovine serum albumin CFU Colony forming units DMEM Dulbecco’s modified Eagle’s medium DMSO Dimethyl sulfoxide ELISA Enzyme-linked immunosorbent assay EMEM Eagle’s minimum essential medium EPS Extracellular polymeric substances FAO Food and Agricultural Organization FBS Fetal bovine serum FCV Feline calicivirus GI/II Genogroup I / II HBGA Histo-blood group antigen HRP Horseradish peroxidase HSV Herpes simplex virus IFN Interferon IgG Immunoglobulin G IL Interleukin INSERM Institut National de la Santé et de la Recherche Médicale LAB Lactic acid bacteria LFMFP Laboratory of Food Microbiology and Food Preservation LMG Laboratory of Microbiology at the University of Ghent LPS Lipopolysaccharide MMTV Mouse mammary tumour virus MNV Murine norovirus vii NoV Norovirus NS Non-structural protein NTPase Nucleoside triphosphate hydrolase NV Norwalk virus OD Optical density ORF Open reading frame PBS Phosphate buffered saline PCR Polymerase chain reaction PFU Plaque forming units PPS Peptone physiological salt PVR Poliovirus receptor RdRp RNA-dependent RNA polymerase RGD Arginyl-glycyl-aspartic acid RNA Ribonucleic acid RT-PCR Reverse transcriptase – polymerase chain reaction SRSV Small round structured viruses TMB Tetramethylbenzidine TSA Tryptic soy agar TSB Tryptic soy broth TV Tulane virus VLP Virus-like particle VP Viral protein WHO World Health Organization viii Abstract It is known that noroviruses (NoVs) show affinity for histo-blood group antigen (HBGA) receptors. The first part of this study aimed to investigate whether the gaseous atmosphere wherein HBGA-expressing E. coli grow influences the receptor expression as well as the protective role these bacteria have on NoV GII.4 virus-like particles (VLPs) from acute heat stress. E. coli LMG 8223 and E. coli LFMFP 861 were included in this experiment. HBGA expression was identified by a method based on the principle of ELISA. The VLP antigen integrity after heat treatment (2 min at 90°C) was detected by a direct ELISA test. The receptor binding ability of the VLPs, after suffering from the same heat stress, was evaluated by a porcine gastric mucin-binding assay. For E. coli LMG 8223, anaerobic growth led to more HBGA expression and aerobic growth resulted in the highest receptor binding ability. No difference between aerobic and anaerobic growth was seen for the VLP antigen integrity. For E. coli LFMFP 861, aerobic growth favoured HBGA expression. Anaerobic growth maintained the VLP antigen integrity the most and no difference was detected for the receptor binding ability. The impact of probiotic bacteria on viral infections has recently gained a lot of interest. In the second part of this study, the effect of Bifidobacterium longum against NoVs was investigated. Murine norovirus type 1 (MNV-1), used as a human NoV surrogate, was detected by means of cell plaque assays with murine macrophage RAW 264.7 cells. The direct effect of the probiotics was found to be limited. Within all the countable results, the number of viruses decreased the most when MNV-1 was co-incubated with cell-free bacteria culture filtrate for 48 h at 37°C: a 2.4 log reduction in plaque forming units per mL (PFU/mL). A reduction greater than 5 log PFU/mL was observed when the virus was co-incubated with a high number of bacterial cells in a skim milk-sucrose matrix. However, it is possible that this reduction is not only caused by the antiviral effect of B. longum, but the viral inhibiting effect of certain milk components might have played a role as well. Results were also obtained that support the idea that the presence of B. longum could inhibit the multiplication of MNV1 on RAW 264.7 cells. ix x Samenvatting Het is reeds geweten dat norovirussen (NoVs) affiniteit vertonen voor een binding met histo bloedgroep antigen (HBGA) receptoren. Daarnaast zijn er ook al bacteriën geïdentificeerd die deze HBGA’s produceren en op hun oppervlak tot expressie brengen. Het eerste deel van deze studie had als doel te onderzoeken indien de HBGA-expressie van dergelijke bacteriën, alsook de beschermende rol die de bacteriën uitoefenen tegenover NoV GII.4 virus-like particles (VLPs) tijdens acute hittestress, beïnvloed wordt door de bacteriën ofwel aeroob ofwel anaeroob op te groeien. E. coli LMG 8223 en E. coli LFMFP 861 werden gebruikt in deze test. De meting van de HBGA-expressie was gebaseerd op de ELISA-techniek. De integriteit van VLP-antigenen na hittebehandeling (2 min bij 90°C) werd gedetecteerd met behulp van een directe ELISA test. De mate waarin VLPs nog in staat zijn om aan hun receptor te binden na hittebehandeling werd geëvalueerd aan de hand van een zogenaamde ‘porcin gastric mucin-binding’ assay. Bij E. coli LMG 8223 werd HBGA-expressie bevoordeeld bij anaerobe groei, maar aerobe groei van de bacteriën leverde dan weer de hoogste receptorbinding op. Er werd geen verschil waargenomen tussen aerobe of anaerobe groei inzake de integriteit van VLP-antigenen. Bij E. coli LFMFP 861 resulteerde aerobe groei in een hogere HBGA-expressie. Anaeroob opgegroeide bacteriën konden de integriteit van VLP-antigenen het best bewaren, maar qua binding aan de receptoren werd geen verschil waargenomen. Het potentiële nut van probiotica in de strijd tegen virale infecties geniet steeds meer interesse van wetenschappers. In het tweede deel van deze studie werd het effect van Bifidobacterium longum tegen NoVs onderzocht. Murine norovirus type 1 (MNV-1) werd gebruikt als surrogaat voor het menselijke NoV. De detectie van het virus gebeurde via cell plaque assays met RAW 264.7 macrofaagcellen van de muis. Het directe effect van B. longum op het virus bleek eerder beperkt te zijn. Uit alle telbare resultaten bleek dat het virusaantal het meest daalde wanneer MNV-1 48 uur bij 37 °C geïncubeerd werd samen met celvrij filtraat van een B. longum cultuur: een reductie van 2,4 log plaque vormende eenheden per mL (PVE/mL). Een daling van meer dan 5 log PVE/mL werd bekomen door het virus te incuberen samen met een hoog aantal bacteriecellen in een medium van magere melk met sucrose. Het is echter mogelijk dat deze daling niet enkel veroorzaakt werd door de probiotische bacteriën, maar ook door welbepaalde melkproteïnen die de virale infectie van cellen kunnen voorkomen. Daarnaast werden ook resultaten bekomen die de idee steunen dat de aanwezigheid van B. longum de vermeerdering van MNV-1 in RAW 264.7 cellen inhibeert. xi xii Chapter 1: Introduction Norovirus (NoV) is the most important cause of foodborne acute gastroenteritis worldwide. Annually, the estimated number of people that get infected with NoVs lies around 20 million in the USA and 15 million in Europe (CDC a, n.d.; WHO, 2015). The most common clinical symptoms include vomiting, non-bloody diarrhoea, abdominal cramps, nausea, fever and dehydration. NoV infections are mildly virulent, transient, and self-limiting within 2-3 days. However, sometimes hospital care is required or death occurs in infants, elderly, chronically ill or immunocompromised patients. Up to now, no vaccine or therapeutic treatment is available for this viral disease. The genetic heterogeneity among the different NoV strains and the inability to cultivate human NoVs make it very difficult to perform direct studies on this virus. Within the complicated transmission routes, evidences have been found that interactions occur between NoVs and bacteria in the environment. For instance, it was shown that bacteria which express histo-blood group antigen (HBGA) like structures have affinity to NoVs (Hutson et al., 2002; Li et al., 2015; Lindesmith et al., 2003). Other studies indicated that probiotic bacterial strains can have antiviral activities against NoVs (Aboubakr et al., 2014; Rubio-del-Campo et al., 2014). The main goal of this thesis was to look deeper into the above mentioned viral-bacterial interactions and discover the importance of specific bacterial strains when it comes to either protecting or fighting against the NoV. The first part of this thesis focused on the protective effect that might occur when human NoVs bind to a selection of HBGA expressing bacterial strains. Different growth conditions of the bacteria were tested for their HBGA expression. The protective effect of aerobically and anaerobically grown E. coli against heat treatment was investigated by looking into the integrity and functionality of NoVs in presence or absence of bacterial suspensions. Respectively a direct enzyme-linked immunosorbent assay (ELISA) and a porcine gastric mucin-binding ELISA were used for this. The second part of this thesis was devoted to the influence of probiotics on NoVs. Viruses and probiotic bacteria were combined at different ratios and incubated at the agreeable conditions for bacterial growth (culture medium, temperature, atmosphere, etc.) for different time intervals. A food matrix was attempted to be introduced in the treatment system in order to investigate the possibility of applying this treatment in food industries. First of all, the virucidal effect of probiotics on virus particles was investigated. This refers to the direct effect of the probiotics. Secondly, the inhibiting effect of probiotic bacteria on the viral infection and replication on a tissue culture of macrophage cells was examined. In both cases, cell plaque assays were carried out. 1 2 Chapter 2: Literature study 2.1 Introduction to Noroviruses 2.1.1 Classification and taxonomy In 1968, an elementary school in Norwalk, Ohio (USA) suffered from an outbreak of ‘winter vomiting disease’. At that time there was no consensus on what the exact agent was that caused the acute gastroenteritis. It took four years before Kapikian et al. (1972) discovered, with the use of immune electron microscopy (Figure 1) on infectious stool filtrates, that a virus was the wrongdoer of the outbreak in Norwalk. The virus got named after the place where it was first discovered: ‘Norwalk Virus’ (NV). Later on also other viruses with a similar morphology were identified and grouped together as ‘small round structured viruses’ (SRSVs). In the 1990’s, further typing and classification of the SRSVs was possible with the help of molecular techniques. Because of this breakthrough, SRSVs were shown to be members of the Caliciviridae family. Figure 1: Immune electron microscopy image of Norwalk Virus from an infected stool filtrate. (Kapikian et al., 1972) The Caliciviridae family is subdivided into four genera: Vesivirus, Lagovirus, Norovirus (formerly known as Norwalk-like viruses) and Sapovirus (the former Sapporo-like viruses) (Baert et al., 2000). Molecular technology, more specifically reverse transcriptase polymerase chain reaction (RT-PCR), has allowed the cloning and sequencing of many NoV strains. Within the genus of NoVs, at least five genetic groups are distinguishable based on the sequence similarities across highly conserved regions in the genome (e.g. the RNA-dependent RNA polymerase [RdRp] and the shell domain of the VP1 capsid protein). 3 Such genetic groups are also termed ‘genogroups’. Genogroups I and II (GI and GII) are the ones containing the majority of NoVs that can cause disease in humans. Human NoVs within one genogroup share at least 60% amino acid sequence identity of the major capsid protein VP1. A genogroup can still be subdivided into so-called ‘genetic clusters’. For example, GII.4 resembles NoV strains from genetic cluster 4 of genogroup II. NoV strains within a genetic cluster share at least 80% VP1 amino acid sequence identity with the cluster’s reference strain (Hutson et al., 2004). So NoVs are genetically and antigenically diverse. An overview of all the genogroups and genetic clusters is presented in Figure 2. Although GII.4 is currently considered the most predominant NoV strain causing viral gastroenteritis worldwide, recent data from China suggest that NoV GII.17 could be the next big upcoming genotype (Lu et al., 2015). Figure 2: Phylogenetic relationship of the five known genogroups (GI to GV) within the NoV genus based on the complete capsid amino acid sequences of NoV strains. (Tan & Jiang, 2011) 2.1.2 Structure NoVs are small icosahedral viruses, with a diameter between 27-30 nm. A positive-stranded RNA genome of 7.7 kilobases is linked with a VPg protein at the 5’-end and polyadenylated at the 3’-end (Baert et al., 2000). The genome is protected from the environment by a protein capsid without envelope. The viral genome is organized into three open reading frames (ORF 1-3), but murine NoVs have a fourth ORF. This extra ORF overlaps ORF 2 but has an alternate reading frame (Karst et al., 2014). The first ORF, at the 5’ proximal end, encodes for a large polyprotein of seven non-structural protein products. Cleaving of this polyprotein is done by a viral protease called NS6 or Pro, which is 4 also one of the seven mature protein products. The other six non-structural products are: NS1, NS2 (p48), NS3 (NTPase), NS4 (p22), NS5 (VPg) and NS7 (RdRp). The functions of all these and other viral proteins are given in Table 1. A visual representation of the NoV genome and the proteins it encodes is shown in Figure 3. Figure 3: The NoV genome. The 5’ proximal ORF 1 (shown in orange) encodes a non-structural polyprotein, which is cleaved into mature products by the virally encoded protease (NS6 or Pro). Other non-structural proteins include NS1/2 (also referred to as p48), NS3 (NTPase), NS4 (p22), NS5 (VPg), and NS7 (the RNAdependent RNA polymerase; RdRp). ORF 2 encodes the capsid protein referred to as VP1; this protein can be divided into shell (S; shown in yellow) and protruding (P) domains. The P domain is further subdivided into the P1 stalk domain (shown in blue) and the hypervariable P2 domain comprising the tips of the arches (shown in red). ORF 3 encodes the minor structural protein VP2, and ORF 4 of murine NoV genomes encodes virulence factor 1, or VF1. NoV genomes are covalently linked to VPg at their 5’ ends and polyadenylated at their 3’ ends. (Karst et al., 2014) Table 1: Names and functions of the proteins that are encoded by a NoV genome. (Karst, 2010) Protein name Function NS1 Unknown 1 NS2 / p48 Inhibits trafficking of host proteins to the cell surface Involved in replication complex formation? Has NTPase activity 1 Inhibits secretion of host proteins Involved in replication complex formation? Primes viral RNA replication following its uridylylation Recruits host translation initiation factors Mediates cleavage of ORF1 polyprotein 1 Replicates the viral genome Generates uridylylated VPg Major structural role 1 Increases expression of VP1 and stabilizes particles Recruits genomes into progeny virions? 3 NS3 / NTPase NS4 / p22 NS5 / VPg NS6 / Pro NS7 / Pol VP1 VP2 Encoded by ORF 1 1 1 2 5 The capsid of NoVs is mainly composed of one major structural protein: viral protein 1 (VP1), which is encoded by ORF 2. This VP1-protein consists of two parts: a well-conserved shell domain (S) and a protruding domain (P). The P domain can be further subdivided into subdomains P1 and P2. Next to VP1, the capsid also contains a few copies of a minor structural protein called VP2. ORF 3 is responsible for encoding this small protein. VP1 and VP2 are translated from subgenomic RNA. With the help of cryo-electron microscopy and x-ray crystallography, Prasad et al. (1999) were able to determine the capsid structure of NoV virus-like particles (VLPs) as shown in Figure 4. Figure 4: The NoV VLP structure. NoV VLPs have 90 dimers of capsid protein (left, ribbon diagram) assembled in T = 3 icosahedral symmetries. Each monomeric capsid protein (right, ribbon diagram) is divided into an Nterminal arm region (green) facing the interior of the VLP, a shell domain (S-domain, yellow) that forms the continuous surface of the VLP, and a protruding domain (P-domain) that emanates from the S-domain surface. The P-domain is further divided into subdomains, P1 and P2 (red and blue, respectively) with the P2subdomain at the most distal surface of the VLPs. (Hutson et al., 2004) 2.1.3 Clinical symptoms and treatment People from all ages are prone to NoV infections. Infections can occur in settings like day care centres, schools, hospitals, hotels, recreational camps, and so on. NoV outbreaks are more likely to occur in cooler winter months, although people can get infected at any time during the year. In addition, there can be an increase of 50% in the number of NoV illnesses in years when there is a new strain of the virus circulating (CDC a, n.d.). Moreover, the infectious dose of NoV is very low: it is estimated that 10100 viral particles may already be sufficient to infect an individual (ECDC, n.d.). 6 The incubation period of a NoV infection can take 12 to 48 hours (CDC b, n.d.). The most common clinical symptoms include vomiting, non-bloody diarrhoea, abdominal cramps, nausea, fever and dehydration. NoV infections are generally mildly virulent, transient, and self-limiting within 2-3 days. Although most of those infected fully recover, sometimes hospital care may be required or the infection might lead to death in infants, elderly, chronically ill or immunocompromised patients (Baert et al., 2000). There is no specific medicine or cure available to treat or prevent a human NoV infection. An oral rehydration providing electrolyte replacement and sugar (glucose or sucrose) is commonly used as the clinical treatment (Thistlethwaith, 2015). Sometimes patients can suffer from severe dehydration and are unable to tolerate oral fluids. In these cases, intravenous replacement of electrolytes is the method of choice. Since NoV infection has a viral origin, an antibiotic therapy will not be effective. In 2011, Siddiq et al. suggested nitazoxanide, a broad-spectrum antimicrobial agent, as a potential therapeutic treatment for NoV infections because it was successfully used to resolve NoV gastroenteritis of an immunocompromised patient. 2.1.4 Transmission routes and prevalence Worldwide, NoV is the leading cause of acute gastroenteritis (inflammation of the stomach or intestines or both). The virus has been identified as the cause of between 73% to more than 95% of global epidemic nonbacterial gastroenteritis outbreaks and approximately half of all gastroenteritis outbreaks (Atmar & Estes, 2006). In the USA e.g. more than 20 million people per year get infected by this virus, leading to 570-8000 deaths (CDC a, n.d.). WHO estimates that NoV is the most common cause of foodborne illness in the European region with close to 15 million cases each year, causing more than 400 deaths (WHO, 2015). An estimated 200,000 deaths occur annually among young children in developing countries (Patel et al., 2008). Apart from being highly contagious, constantly evolving, and evoking only limited immunity, another attribute that makes NoVs an ideal infectious agent is the fact that multiple transmission routes exist. Factors such as the high stability of the viruses in the environment and their high prevalence facilitate the transmission (Hutson et al., 2004). Furthermore, it is also possible that an infected person can still shed infectious virus particles after recovery for up to three weeks after exposure to NoVs (Rockx et al., 2002). NoVs are foodborne viruses and their primary transmission route is a person-to-person spread through the faecal-oral route. They can also be transmitted through food, water, the environment, contaminated surfaces or aerosols. A schematic overview of the most important NoV transmission routes is presented in Figure 5. 7 Figure 5: Main transmission routes of NoVs. (Leutenez, 2016) The most efficient way of transmitting NoVs is via direct person-to-person contact or via aerosols that are produced while vomiting. Food handlers that are infected with NoV play an important role in spreading the virus, because they can easily transmit viruses to mainly ready-to-eat food (Bidawid et al., 2000). Two other categories concerning the transmission of NoVs through food are fresh produce and shellfish. Fresh produce poses a threat because it is mostly consumed without first undergoing a treatment that could effectively destroy the virus (Ethelberg et al., 2010). Bivalve molluscan shellfish, such as mussels or oysters, are filter-feeders. They take up surrounding water and filter out nutritive components. However, when human pathogens are present in the water, they can also be filtered out and accumulate in the shellfish. Contaminated water is not only an infection source for shellfish (Westrell et al., 2010), but it can also directly infect humans if it is used as drinking water or recreation water (Hoebe et al., 2004). As was noted by FAO and WHO (2008), large-scale outbreaks are often a result of a combination of several transmission routes. For example, the virus can be introduced in a sensitive population by contaminated food, water or an asymptomatic shedder. Direct person-to-person contact or a contaminated environment may lead to an efficient spread of the virus through the population. 8 2.1.5 Pathogenesis The possibility of a NoV infection is not only dependent on the viral dose and strain one is exposed to. Research has shown that the specific human susceptibility plays an important role as well. In 2002, Hutson et al. were the first to report that the risk of a Norwalk virus infection and disease was associated with someone’s ABO blood type. In following research, volunteer studies (Lindesmith et al., 2003) and outbreak investigations (Rockx et al., 2005) delivered increasing evidence of HBGAs being receptors of NoV infection. HBGAs are complex carbohydrates present on the surfaces of red blood cells and mucosal epithelium of the respiratory, digestive and urogenital tracts. They can also be found as free oligosaccharides in biologic fluids such as saliva, intestinal contents, milk and blood (Tan & Jiang, 2005). As reviewed by Hutson et al. (2004), these carbohydrates are synthetically related triand tetrasaccharide moieties that are located at the distal ends of carbohydrate chains on cellular glycolipids and glycoproteins found on the exterior cell surface. The variety of expressed carbohydrates is determined by the presence or absence of specific glycotransferase enzymes as a result of a person’s genetics. The human HBGA system is controlled by multiple gene families, that contain silent alleles (Marionneau et al., 2001). The recognition of human HBGAs by NoVs occurs through a typical protein-carbohydrate interaction, in which the protruding domain of the viral capsid protein forms an interface with the oligosaccharide side-chains of the antigens. The P2 subdomain forms the most exterior surface of the capsid (Figure 4). A plausible binding pocket in the P2 subdomain was identified by sequence homology modelling based on the receptor binding patterns of different NoV strains (Tan et al., 2003). The pocket consists of a RGD-like motif and three scattered, binding pattern-specific amino acids. Nearby this pocket, another binding site that is composed of three amino acid residues on the NoV capsid was identified (Chakravarty et al., 2005). Thus, the two binding sites could belong to one binding interface and act cooperatively with respective sugar side-chains of the HBGAs. Figure 6 displays a close-up starting from a NoV particle up to the HBGA binding site. Although a lot of efforts have already been made to investigate the human NoV-HBGA interaction and since the human NoVs are still non-cultivable, the complete picture of virus-host interaction could be more complex. 9 Figure 6: The four panels show structures of NoVs at different levels: (from left to right) an electron microscopy image of NoVs, a single VLP, a P dimer with indication of the carbohydrate-binding interface (coloured region), and the crystal structure of the HBGA-binding interface. (Tan & Jiang, 2010) 2.1.6 Non-cultivability and surrogates Numerous efforts have been made to establish an in vitro cultivation technique of human NoVs. In 2004, Duizer and his colleagues tried to grow NoV on multiple routinely cultured cell lines (Caco-2, HeLa, and many more). Even with the addition of bioactive digestive enzymes, substances to induce cell differentiation (insulin, DMSO or butyric acid) or using different inoculation methods, successful virus propagation could not be achieved. Three years later, Straub et al. (2007) were the first to report that NoVs can infect and replicate in a 3-dimensional (3-D), organoid model of human small intestinal epithelium. The cells were grown on porous collagen-I coated microcarrier beads under conditions of physiological fluid shear in rotating wall vessel bioreactors. The NoV infection was proven using microscopy, PCR and fluorescent in situ hybridisation. However, no follow-up experiments were found until the same research group studied human NoV infectivity using a 3-D model of large intestinal epithelium (Straub et al., 2011). Culturing human NoVs was made possible to a certain extent using this system, although other laboratories have not yet been able to verify and repeat the findings of Straub. In an attempt to develop an in vitro infection model of human NoVs, Jones et al. (2014) identified B cells as a cellular target of the virus. A BJAB human B cell line was inoculated with the Sydney human NoV GII.4-positive stool sample. The viral genome copy numbers increased 10 to 25 times at respectively three and five days after infection. Notably, the replication in B cells required the presence of HBGA-expressing enteric bacteria. Nowadays, a lot of attention is paid on this topic and further research is being done to confirm the model of Jones. The study of human NoVs faces some difficulties due to the inability to distinguish infectious human NoVs from the non-infectious ones, and the unavailability of large amounts of infectious viruses. To counteract these problems, surrogates are used to study the stability and inactivation of human NoVs. 10 The surrogates should share pathological and/or biological features with the human NoVs and they should be able to propagate in cell culture. Much of the current knowledge on NoV biology was acquired by studies with surrogates. As an example, studies with murine NoVs have led to the discovery of the important role of the STAT-1 molecule in the resistance of a NoV disease in vivo and the control of virus growth in vitro (Mumphrey et al., 2007). Feline calicivirus (FCV) shows similarities with human NoVs in primary sequence and genomic organisation. In addition, the virus grows well in vitro. These characteristics made FCV a frequently used surrogate virus in previous studies (e.g. Aboubakr et al., 2014; Lee et al., 2012; Sosnovtsev et al., 2005). The main disadvantages of this cat virus are: it does not cause gastro-intestinal diseases but respiratory ones, and it does not bind to HBGAs but to sialic acid (reviewed by Vashist et al., 2011). Murine norovirus type 1 (MNV-1) was firstly identified in 2003 by Karst et al. Reasons why MNV-1 nowadays is often the model of choice for human NoV research include its close relatedness to human NoVs, the natural host is relatively cost-effective and genetically well-characterised, and it is the only NoV that can be efficiently propagated in vitro because it has a tropism for dendritic cells and macrophages (Vashist et al., 2011). This last feature allows the propagation of MNV-1 in RAW 264.7 murine macrophage cells (Wobus et al., 2004). MNV-1 and human NoV not only share a similar size and shape, but also the three big ORFs are characteristic (Wobus et al., 2006). However, the use of MNV-1 as a surrogate for human NoVs has some limitations. First of all, MNV-1 uses sialic acid as a receptor for attachment to specific cell lines instead of HBGAs. Secondly, MNV-1 does not cause gastroenteritis in its host, but some of the occurring symptoms include encephalitis, meningitis or vasculitis. Another attractive surrogate model could be a virus that infects a non-human primate. An example in this category is the Tulane virus, a calicivirus discovered in rhesus monkeys (Farkas et al., 2008). Although this virus can recognize HBGAs as receptors, it does not belong to the NoV genus and it does not cause gastroenteritis. The use of bacteriophages as surrogates can be considered because of their non-pathogenic properties and easy detection and cultivation (Li, 2012). Coliphage φX174 is an examples of a bacteriophage surrogate for human NoV that has already been used for experimental means (Li et al., 2011). The use of recombinant VLPs has been a valuable method to investigate the virus-host interaction. The discovery of NoVs being able to bind to HBGAs, as described earlier, was achieved by performing experiments with human NoV VLPs (e.g. Harrington et al., 2002 & 2004). VLPs are generated by the expression of the viral major capsid protein (VP1) with or without the minor capsid protein (VP2). The produced proteins then self-assemble to form VLPs, having the same morphological characteristics of 11 the original human NoV virion (Vashist et al., 2011). Expressing the viral capsid proteins in insect cells using baculovirus expression systems is a frequently used strategy (Belliot et al., 2001; Caddy et al., 2014; Li et al., 2015). In some cases, only the P domain of the NoV capsid is considered for experimental use. The domain forms a 24-mer subviral particle, called P particle. P particles can easily be produced in E. coli, are very stable and also highly immunogenic, making them an interesting platform for vaccine development (Tan & Jiang, 2012). 2.2 Interactions between bacteria and viruses 2.2.1 Types of interactions There is an estimation of 1031 viruses on earth (Breitbart & Rohwer, 2005). Scientists have estimated the total number of bacteria on earth to be 5 x 1030 (University of Georgia, 1988). The human body is colonized by an immense population of microorganisms. A widely cited statement says that bacteria outnumber the amount of human cells by a ratio 10:1 in humans. However, a recent study rejects this theory, saying that a “reference man” has approximately the same number of bacteria and human cells in his body (Sender et al., 2016). Considering that the mean weight of a cell equals one nanogram, a person of 70 kg would consequently harbour about 70 trillion bacteria. As humans also often get infected by viruses, it would not be a surprise if an interaction would occur between viruses and bacteria inside the body. Since this thesis mainly focuses on NoVs, this section will focus on interactions between enteric bacteria and NoVs. A first category of mechanisms involves the direct facilitation of viral infection, including bacterial stabilization of viral particles and the facilitation of viral attachment to host target cells (Karst, 2016). Binding of enteric viruses to bacterial surface polysaccharides enhances virion stability towards a heat treatment or inactivation by chlorine bleach (Robinson et al., 2014). The same research group also discovered that bacterial polysaccharides enhance poliovirus cell attachment by increasing binding to the viral receptor PVR. An example of increased cell invasion is the stimulation of B cell infection by NoVs in presence of HBGA expressing bacteria (Jones et al., 2014). A graphical presentation of these examples is given in Figure 7. 12 Figure 7: (A) The binding of poliovirus to lipopolysaccharide (LPS) leads to an increase in viral thermostability and resistance to inactivation by dilute chlorine bleach. (B) Poliovirus associated with LPS binds to the poliovirus receptor (PVR) more efficiently on the surface of target cells. Several lines of evidence show that LPS enhancement is conferred by facilitating viral binding to the known PVR: pre-treatment of permissive cells with PVR-specific antibody inhibits viral binding in both the presence and absence of LPS; virus bound to LPS does not gain competency to infect non-PVR-expressing cells; and virus incubated with LPS has increased binding to soluble PVR compared with virus alone. (C) Human NoV infection of B cells is facilitated by commensal bacteria that express the appropriate HBGA. The first indication that commensal bacteria stimulate human NoV infection of B cells was provided by the observation that the filtration of virus-positive stool to remove commensal bacteria reduced viral infectivity. Providing direct evidence, the supplementation of filtered stool with HBGA-expressing bacteria fully restored infectivity. (Karst, 2016) 13 A second category concerns mechanisms that indirectly influence the antiviral immune response in a way that promotes viral infection (Karst, 2016). The strategy mouse mammary tumour virus (MMTV) uses to evade the host’s immune responses has been described by Kane et al. (2011). The binding of MMTV to bacterial lipopolysaccharide triggered Toll-like receptor 4 on macrophages or dendritic cells. These cells subsequently started producing interleukin-6 (IL-6), leading to the induction of the inhibitory cytokine IL-10 by B cells. An immunosuppressive microenvironment was created by this means. As a consequence, MMTV could persist and a tolerance for their antigens was established. In 2014, Kernbauer, Ding, & Cadwell found that commensal bacteria possess the ability to suppress the type III interferon (IFN) response by diminishing the production of antiviral IFNλ. If IFNλ is produced, which is the case in microorganism-depleted mice, it will activate the type III interferon receptor (IFNλR) in enterocytes or in other bystander cells to indirectly inhibit norovirus persistence. So suppressing IFNλ production facilitates MNV persistence. In mice lacking functional type III IFN signaling pathways, interactions with commensal bacteria are not necessary to gain persistence because the bystander cells are impaired in their ability to respond to IFNλ (Karst, 2016). 2.2.2 Histo-blood group antigen expressing bacteria HBGAs play an important role in multiple viral infections. It has been mentioned before that human NoVs contain binding pockets in their P2 domain, through which they are able to bind carbohydrate structures on the surfaces of red blood cells and mucosal epithelium of the respiratory, digestive and urogenital tracts (Tan et al., 2003). These receptors were identified as HBGAs. NoVs are not the only viruses that can bind HBGAs. In 2012, Huang et al. discovered that human rotaviruses, the most important cause of severe gastroenteritis in children, also recognize HBGAs. The spike protein VP8* of the virus is responsible for this interaction. A number of gram-negative bacteria show blood group expression as well (Springer et al., 1961). The discovery that human NoVs are capable of binding to these HBGA expressing bacteria has only recently been confirmed. Transmission electron microscopy demonstrated that VLPs of human NoVs mainly bound to extracellular polymeric substances (EPS) of an enteric bacterium strain (SENG-6), closely related to Enterobacter cloacae (Miura et al., 2013). This is also the location bearing HBGA-like molecules. NoV VLPs of a GI.1 wild-type strain and a GII.6 strain that can recognize HBGA type A were shown by ELISA to bind to the EPS extracted from Enterobacter sp. SENG-6. The study of Jones et al. (2014) identified B cells as a cellular target of both murine and human NoVs. The presence of enteric HBGA expressing Enterobacter cloacae bacteria was essential to facilitate viral attachment to, and infection of, B cells. M12 and WEHI-231 mouse B cell lines were infected with MNV14 1 or MNV-3 to check whether MNVs infect B cells in culture. Both virus strains replicated efficiently in B cells. In vivo experiments confirmed that B cells are indeed targets of the NoVs: significant reductions in the number of viruses were found in B cell-deficient mice compared to normal mice. Antibiotic depletion of normal intestinal flora in mice led to a significant drop in MNV titers, demonstrating that enteric bacteria play an important role during the process of NoV infection. For human NoVs, a BJAB human B cell line was inoculated with the Sydney human NoV GII.4-positive stool sample. The viral genome copy number increased 10 to 25 times at respectively three and five days after infection. Filtering the infected stool sample over a 0.2 µm membrane reduced genome replication. Adding Enterobacter cloacae to the filtered stool before inoculation of BJAB B cells restored infectivity of the virus in a dose-dependent way. Enterobacter cloacae is an HBGA type H expressing bacterial strain to which the GII.4 Sydney human NoV strain can bind. The presence of synthetic H antigen together with the filtered stool was sufficient to restore the attachment of the virus to B cells. In 2015, a study was done by the Laboratory of Food Microbiology and Food Preservation (LFMFP) at Ghent University on the effect of HBGA expressing bacteria on human NoV (Li et al., 2015). Both E. coli strains LMG 8223 and LFMFP 861 were found to express HBGAs and bind to human NoV VLP GI.1 and GII.4. E. coli ATCC 8739 served as a negative control, since it shows no HBGA expression and no VLP binding. VLPs pre-incubated with HBGA expressing or non-HBGA expressing bacteria were heated at 90°C for 2 min. Afterwards, the antigenicity of the VLPs was measured by a direct ELISA test. A porcine gastric mucin-binding ELISA test examined the receptor binding ability of VLPs after the heat treatment. As a result, the presence of HBGA expressing E. coli always secured higher antigen integrity as well as mucin-binding ability of the VLPs after exposure to heat stress, indicating a protective effect of HBGA expressing E. coli on human NoVs from acute heat treatment. 2.2.3 Probiotics The consumption of antibiotics to kill harmful bacterial infections is huge: from 2000 to 2010, the global antibiotic consumption grew from approximately 30 billion to 50 billion standard units, based on data gathered from 71 countries (Van Boeckel et al., 2014). Having the wrong bacteria in the wrong place can cause health problems. However, the right bacteria in the right place can lead to benefits. This is the part where probiotics have their importance. Probiotics are defined as “living micro-organisms, which upon ingestion in certain numbers, exert health benefits beyond inherent basic nutrition” (Gionchetti et al., 2012). One of the largest groups of probiotic organisms are the lactic acid bacteria (LAB), containing e.g. Lactobacillus sp, Bifidobacterium 15 sp and Enterococcus sp (Ljungh & Wadstrom, 2006). The presence of living bacteria inside the human body is not abnormal. There are up to 1013 - 1014 bacteria found in the gastrointestinal tract, divided over more than 400 bacteria species (FAO & WHO, 2001). In order for those bacteria to be probiotic, they should be able to survive both the stomach and bile acids, sufficient quantities must arrive in the intestines and they must show some beneficial effects on human health. A few examples of the beneficial health effects probiotics promote, include alleviation of lactose intolerance, prevention and treatment of diarrhoea, maintenance of normal intestinal flora, antagonism against pathogens, stimulation of the immune system, anticarcinogenic activity, and reduction of serum cholesterol levels (BC Dairy, 2016). Effects on the human or animal host may be caused directly by preventing the infection and fighting against the causative agent of an intestinal disorder, or indirectly by balancing the disrupted equilibrium of the enteric flora and raising the host’s immune response (Maragkoudakis et al., 2010). Probiotics are not only found in the human body. Other sources are certain food products (e.g. fermented dairy products or soybean foods) or dietary supplements. Figure 8 gives an overview of the main mechanisms of action of probiotics. Figure 8: Mechanisms of action of probiotics. The probiotics are coloured in yellow. IEC: epithelial cells, DC: dendritic cells, T: T cells, Tn: naïve T cells, Th: helper T cells, Treg: regulatory T cells, B: B cells, IL: interleukin, TGF: transforming growth factor. (Custom Probiotics Inc., n.d.) 16 LAB strains generally produce antimicrobial substances with a narrow or broad spectrum against homologous gram-positive bacteria. They additionally often produce microbicidal substances with dual functions: showing an effect against gastrointestinal pathogens and other microbes, or competing for cell surface and mucin binding sites (Ljungh & Wadstrom, 2006). The LAB metabolites causing antimicrobial activity include organic acids (lactic and acetic acid), hydrogen peroxide, ethanol, diacetyl, acetaldehyde, carbon dioxide, reuterin, and bacteriocins (Suskovic et al., 2010). A few studies demonstrated the possibility of probiotics or the substances they naturally produce to not only possess antimicrobial characteristics, but also enhance the antiviral effect on viruses. For instance, by blocking binding sites on epithelial cells, some Bifidobacterium species can inhibit the growth of viral pathogens (Colbère-Garapin et al., 2007). Bacteriocins could help in reducing the viral coat protein synthesis (Wachsman et al., 1999), decreasing the infectious virus yield (Serkedjieva et al., 2000), or blocking the host cell receptor required for viral attachment (Berkhout et al., 1997). The use of probiotics as a possible method to control and treat viral infections is an interesting topic for investigation. In the case of human NoVs for example, several key challenges remain in assessing the efficacy of vaccines and antiviral drugs. There is for instance no robust cell culture system, limiting the direct study of these viruses. Also the genetic heterogeneity among NoV strains makes it difficult to find a suitable vaccine. If probiotics have antiviral activity against NoV, this will offer a promising alternative to finding a cure for NoV infections. A lot of researches have already been done to examine the influence of probiotics on viruses. The studies listed in Table 2 aimed at examining possible antiviral activities of probiotic LAB on viruses. Frequently used techniques to detect the viruses included cell plaque assays, RT-qPCR and ELISA. In most experiments, the presence of LAB probiotics reduced the infectivity of viruses. However, this reduction was very variable. For instance, a 6 to 7 log infectivity reduction was measured when FCV was co-incubated with either bacterial growth medium filtrate or bacterial cells of the Lactococcus lactis strain (Aboubakr et al., 2014). On the other hand, no direct effect was found on MNV-1 and Tulane virus (TV) when they were co-incubated with cell-free supernatant of a commercial probiotic mixture (Shearer et al., 2014). It is difficult to draw general conclusions. A lot of different LAB strains and viruses were used in the listed experiments, and there is still a lack of enough thorough studies on this topic. 17 Table 2: Antiviral studies of probiotics. Bacteria Virus Cell line Effect Reference Lactobacillus delbrueckii subsp. bulgaricus 1043 Lactobacillus paracasei/ rhamnosus Q85, Lb. paracasei A14 and F19, Bifidobacterium longum Q46 Lb. paracasei, Lactobacillus reuteri SD2112, Lb. rhamnosus GG, Lb. paracasei NCC 2461, Lactobacillus johnsonii NCC 533, Streptococcus thermophilus NCC 2496 Lb. rhamnosus GG, Lb. casei Shirota Influenza virus A chick embryo fibroblast cells Serkedjieva et al. (2000) Vesicular stomatitis virus pig alveolar macrophage cells Rhesus rotavirus MA104 cells (mouse) No inactivation effect on extracellular virus Viral reproduction considerably reduced Lactobacilli and bifidobacteria decrease viral infection by establishing the antiviral state in macrophages, by production of NO and inflammatory cytokines Lactobacillus rhamnosus strain GG had the strongest influence in reducing prevalence, duration and severity of diarrhoea Rotavirus, Transmissible gastroenteritis virus Human immunodeficiency virus type 1 CLAB porcine epithelial cells Maragkoudakis et al. (2010) Herpes simplex virus (HSV) type 1 Vero African green monkey kidney cells and murine monocyte/macrophage cell line J774 cells RAW 264.7 and CrandellReese feline kidney cells Co-incubation of the CLAB cell line with the bacteria resulted in increased survival percentages, up to 80% Greatest virus inhibition with Lactobacillus curvatus VM25 (55.5%), Lactobacillus fermentum VMA (52.5%), and Pediococcus pentosaceous VM95 (49.0%) Lb. rhamnosus significantly increased cell viability (47.6 – 49 %) of the HSV-1 infected Vero cells and it reduced plaque forming (15 – 18 %). Similar effect for E. coli. At the end of the fermentation (20 days), the population of FCV and MNV decreased about 4.12 and 1.47 log units, respectively Pre-treatment of FCV with BGMF or BCS: 1 to 2 log reduction Co-treatment of FCV with BGMF or BCS: 6 to 7 log reduction The intracellular level of Coxsackievirus B3 RNA was inhibited by 50% Kim et al. (2014) 38 bacterial strains (15 different species), isolated from breastmilk of healthy women Lb. rhamnosus PTCC 1637, Escherichia coli PTCC 25923 TZM-bI cells LAB species in general after Dongchimi fermentation Murine norovirus, Feline calicivirus Lactococcus lactis subsp. lactis LM0230 (bacterial growth medium filtrate (BGMT) or bacterial cell suspension (BCS)) Bifidobacterium adolescentis SPM 1605 Feline calicivirus Crandell-Reese feline kidney cells Coxsackievirus B3 HeLa cells Ivec et al. (2007) Pant et al. (2007) Martín et al. (2010) Khani et al. (2012) Lee et al. (2012) Aboubakr et al. (2014) 18 E. coli Nissle 1917, Lactobacillus casei BL23 Norovirus P particles HT-29 cells Commercial mixture of Lactobacillus acidophilus, Lb. rhamnosus, Bifidobacterium bifidum, Lactobacillus salivarius, and S. thermophilus LAB species in general after oyster fermentation Murine norovirus 1, Tulane virus RAW 264.7 and LLCMK2 cells Murine norovirus, Feline calicivirus RAW 264.7 and CrandellReese feline kidney cells Lactobacillus bulgaricus 761N Hepatitis C virus Human liver hepatocellular carcinoma (HepG2) cells, Lactobacillus ruminis SPM 0211, B. longum SPM 1205 and SPM 1206 Human rotavirus Caco-2 and Vero cells, In vivo in mice E. coli Nissle, Lb. rhamnosus GG Human rotavirus In vivo in neonatal gnotobiotic piglets B. adolescentis Murine norovirus 1, Human norovirus GI.1 VLP RAW 264.7, Caco-2 and HT-29 cells Total inhibition when co-incubating E. coli with the virus particles and the cells. The effect was limited with Lb. casei A lack of direct effect of cell free supernatant on the virions Rubio-del-Campo et al. (2014) Fermentation with 5 % NaCl for 15 days: MNV and FCV decreased with 1.60 and 3.01 log units, the number of LAB increased from 3.62 to 8.77 log CFU/g Pre-treatment (adding bacteria to the cells before incubation with HCV): 70.9 % reduction in viral load Post-treatment (adding bacteria after incubation): 88.7 % reduction Similar results for Caco-2 and Vero cells: Lb. ruminis SPM 0211 and B. longum SPM 1206 decreased plaque formation up to 45 % (dependent on the bacterial dose). In vivo: Lb. ruminis SPM0211 caused a decrease of 56 % in viral gene expression, B. longum SPM 1205 and SPM 1206 respectively 39 % and 47 %. Mean peak virus shedding titers were significantly lower in the E. coli- colonized compared with Lb. rhamnosus-colonized or uncolonized piglets. Presence of B. adolescentis inhibits multiplication of MNV-1 on RAW 264.7 cells within 48 h of coincubation and it also decreased the binding of human NoV GI.1 VLPs to both Caco-2 and HT-29 cells. Seo et al. (2014) Shearer et al. (2014) El-Adawi et al. (2015) Kang et al. (2015) Kandasamy et al. (2016) Li et al. (unpublished) 19 20 Chapter 3: Material and Methods 3.1 Interaction between Norovirus and HBGA expressing bacteria The objective of this study is to investigate whether the gaseous atmosphere, in which HBGA expressing bacteria grow, could affect the protective effect of these bacteria on NoVs from heat stress. Due to a limitation of the facility availability, flow cytometry was not used like previously (Li et al., 2015). Alternatively, an ELISA based method was firstly established to measure HBGA expression of the bacteria. Optimisation was performed to determine the most favourable antibody concentration, blocking reagent concentration and starting bacterial cell numbers. The investigation of the influence of the gaseous atmosphere started by following the growth of HBGA expressing E. coli aerobically and anaerobically. Subsequently, the effect of bacterial growth atmosphere on the viral protection capability was explored by three different tests. The pre-optimised HBGA expression test was employed to compare the HBGA expressing levels produced by the bacteria. The direct ELISA method was employed to measure the antigenicity of NoVs after they suffered a heat treatment in the presence of (an)aerobically grown HBGA expressing bacteria. The mucin-binding ELISA test was employed to study the receptor binding ability of the viruses after heat stress in the presence of (an)aerobically grown HBGA expressing bacteria. These three tests were performed with the use of two different HBGA expressing E. coli strains, in order to observe the consistency between strains. Figure 9 displays a flowchart of the procedures described above. Optimisation HBGA measuring test • Concentration antibodies • Concentration blocking reagent • Bacterial cell numbers E. coli growth: aerobic vs anaerobic atmosphere E. coli LMG 8223: aerobic vs anaerobic growth E. coli LFMFP 861: aerobic vs anaerobic growth • HBGA expression • Direct ELISA • Mucin-binding ELISA • HBGA expression • Direct ELISA • Mucin-binding ELISA Figure 9: Flowchart showing the different steps that were performed to study the protective effect of HBGA expressing bacteria on NoVs when the bacteria were grown in different gaseous conditions. 21 3.1.1 Bacteria An overview of the bacterial strains that were used to study the interactions of NoVs and HBGA expressing bacteria, is presented in Table 3. Escherichia coli LMG 8223 was selected because it was found by Li et al. (2015) to express type A HBGAs and it is also capable to bind to human NoV VLP GII.4. This strain was obtained from the Belgian Coordinated Collection of Microorganisms (BCCM/LMG). E. coli LFMFP 861 expresses type B HBGAs and can also bind human NoV VLP GII.4. This strain was isolated at the Laboratory of Food Microbiology and Food Preservation (LFMFP), Ghent University. E. coli ATCC 8793 was obtained from the American Type Culture Collection (ATCC). This strain was used as a negative control because it does not express HBGAs. All bacterial strains were cultured in tryptic soy broth (TSB, Oxoid, Thermo; Hampshire, UK) at 37°C. Table 3: Characteristics of the E. coli strains that were used for the experiments with HBGAs. The percentage of positively stained cells determined by flow cytometry analysis were grouped as in the following. < 1%: - ; 1-5%: + ; 5-10%: ++ ; > 10%: +++. (Dan Li et al., 2015) Bacteria Strain Biological origin Escherichia coli E. coli E. coli LMG 8223 Clinical isolate ATCC 8739 LFMFP 861 Feces Thick whey products HBGA expression Type A Type B +++ ++ - +++ VLP GII.4 binding +++ +++ 3.1.2 Virus-like particles (VLPs) and antibodies VLPs were used as an alternative for human NoV, due to the non-cultivability of the latter. VLPs of human NoV GII.4, based on the NoV isolated in Dijon in 1996, were obtained from the Institut National de la Santé et de la Recherche Médicale (INSERM, Paris, France). Anti-HBGA antibodies (anti-A #21 and anti-B #49) and anti-VLP antibodies (lp132) were also supplied by INSERM. Horseradish peroxidase (HRP) congjugated anti-mouse IgG and HRP-conjugated anti-rabbit IgG antibodies were purchased from Promega (Madison, WI, USA). 3.1.3 Monitoring of the bacterial growth Bacteria culture was made by dilution in peptone physiological salt solution (PPS; 1 g of neutralised bacteriological peptone [Oxoid, Basingstoke, UK] and 8.5 g of NaCl [Sigma Aldrich, St. Louis, MO, USA] per liter) and inoculation into fresh TSB (30 g of tryptic soy broth powder [Oxoid] per liter) from an estimated concentration of 105 to 106 CFU/ml. Aerobic culture was made by incubation at 37°C 22 overnight while shaking at a speed of 200 rpm (IKA®-Werke GmbH & Co. KG, Staufen, Germany). Anaerobic culture was made by incubation at 37°C overnight with the use of ANAEROGENTM COMPACT (Oxoid, Thermo). Before each experiment, a first passage was made aerobically as described above from the defrozen culture to activate the bacteria. The defrozen culture was stored at 4°C for maximally one month. Experimentally applied bacteria were always prepared from the activated first passage either aerobically or anaerobically as described above (the second passage). In order to follow the bacterial growth, five tubes were prepared from the first passage in parallel and incubated under the conditions defined above for 0, 3, 6, 12 and 24h respectively. For the bacteria from each tube, the optical density at 600 nm (OD600) and viable bacteria numbers were measured. The optical density was measured by using a VersaMax ELISA Microplate Reader (Molecular Devices, Sunnyvale, CA, USA). In order to obtain viable bacteria numbers, a tenfold dilution series in PPS was made for each tube, and 100 µL of each dilution was plated onto tryptic soy agar (TSA; 40 g of tryptic soy agar powder [Oxoid] per liter). After overnight incubation at 37°C, the colonies that were formed on the spread plates were counted. The following formula was used to calculate the number of bacteria in the original second passage tubes: X= A∗V I X equals the number of CFUs per mL in the second passage tube, A represents the number of counted colonies, V is the dilution factor and I the inoculation volume in mL. 3.1.4 HBGA expression test Bacterial cells (concentration to be determined by preliminary test) were collected by centrifuging for 5 min at 6,000 rpm (Eppendorf Micro Centrifuge 5415C). The pellet of each sample was washed with 100 µL phosphate buffered saline (PBS, pH 7.4 [Lonza, Verviers, Belgium]) by centrifuging, resuspended with 100 µL of bovine serum albumin (BSA; Sigma Aldrich; concentration to be determined by preliminary test) in PBS and incubated at 37°C for 1 h to block the nonspecific binding. After centrifugation, 100 µL of mouse anti-HBGA monoclonal antibodies (anti-A #21 for E. coli LMG 8223, anti-B #49 for E. coli LFMFP 861) that were diluted 250 times in 0.1% BSA-PBS were added to the pellet. The mixtures of bacteria and antibodies got vortexed and then incubated at 37°C for 1 h. Subsequently, two washing steps by centrifugation with PBS were performed. One hundred µL secondary antibodies, anti-mouse antibodies conjugated with the enzyme HRP at a 1:10,000 dilution 23 in 0.1% BSA-PBS, were added and incubated at 37°C for 1 h. Two washing steps by centrifugation were performed with PBS. Per sample, 100 µL 3,3′,5,5′-Tetramethylbenzidine (TMB) One Solution (Promega, Madison, WI, USA) was added as the substrate for HRP. After adding 50 µL 0.10 N phosphoric acid (H3PO4), the optical density was measured at 450 nm by a VersaMax ELISA Microplate Reader. 3.1.5 Direct ELISA E. coli cells were washed with PBS and mixed with NoV GII.4 VLPs (100 µL per sample, 1:100 diluted in PBS to get 5 µg/mL). These mixtures of VLPs and bacteria in 0.5 mL thin-walled PCR tubes were heated at 90°C for 2 min in a PCR cycler (Arktik™ Thermal Cycler, Thermo Fisher Scientific; Waltham, MA, USA) and cooled down immediately on ice. The treated VLP-bacteria mixtures were diluted at 1:10 in a carbonate buffer (CBS, 1.50 g Na2CO3 [Sigma Aldrich] and 1.92 g NaHCO3 [Merck KGaA, Darmstadt, Germany] per liter, pH 9.6) and then coated on a Nunc Maxisorp immunoplate (100 µL per well; Sigma Aldrich) overnight at 4°C. Three non-coated wells were included as blank controls. The plate was washed three times with PBS-T (0.05% Tween 20 [polyoxyethylenesorbitan monolaurate, C58H114O26, Sigma Aldrich] in PBS) and blocked with 5% skim milk (Difco TM; Becton, Dickinson and Company; Sparks, MD, USA) in PBS (‘5% milk-PBS’) for 1 h at 37°C. One hundred µL primary antibodies, anti-VLP rabbit polyclonal antibodies lp132 diluted at 1:1,000 in 5% milk-PBS, were added per well and incubated for 1 h at 37°C. After washing, 100 µL secondary antibodies, HRPconjugated anti-rabbit IgG antibodies, diluted 1:2,500 in 5% milk-PBS were added per well and incubated 1 h at 37°C. After washing, 100 µL TMB One Solution was added as the substrate of the enzyme, followed by adding of 50 µL phosphoric acid to stop the reaction. The optical density was measured at 450 nm (OD450). 3.1.6 Mucin-binding ELISA A Nunc Maxisorp immunoplate was pre-coated with porcine gastric mucin (Sigma-Aldrich, 2µg/well). After overnight incubation at 4°C, the wells were washed three times with PBS-T and then blocked with 5% milk-PBS for 1 h at 37°C. E. coli cells were washed with PBS and mixed with NoV GII.4 VLPs (100 µL per sample, 1:100 diluted in PBS to get 5 µg/mL). These mixtures, either untreated or treated at 90°C for 2 min in a PCR cycler and then cooled down on ice, were added to each well. Five per cent milk-PBS without VLPs was added to mucin-coated wells to include blank controls. The plates were incubated for 1 h at 37°C. The rest of the procedure – adding antibodies, TMB, H3PO4 and measuring the absorbance at 450 nm – was performed as was described for the direct ELISA. 24 3.2 Interaction between Norovirus and probiotic bacteria The second part of this thesis was dedicated to the influence of probiotic bacteria on NoVs. Bifidobacterium longum was chosen for this study based on the previous results generated from LFMFP, Ghent University (data not published). The growth of B. longum was followed both in bacteria culture medium and in skim milk, a representative model for food matrices being natural sources of probiotics. Murine Norovirus 1 (MNV-1) was used as a surrogate of human NoVs. Plaque assays were used to test the antiviral effect of B. longum on MNV-1. First of all, the survival of MNV-1 in different solutions wherein the treatment with B. longum would occur were tested. In the following, the effect of B. longum on MNV-1 was investigated with different bacterial cell numbers, treatment time intervals and food matrices. Figure 10 shows an overview of the experimental set-ups. Growth of B. longum in TSB Choice of a food matrix model Cell plaque assay without bacteria Cell plaque assay with bacteria • TSB • Milk + sucrose • Cell-free bacteria culture filtrate • 1010 bacterial cells for 1 h • Bacteria growing in TSB for 0 to 48 h • 1011 bacterial cells in TSB or milk + sucrose for 48 h Figure 10: Flowchart showing the different steps that were performed to study the effect of B. longum as a probiotic bacterium on a NoV infection. 3.2.1 Bacteria and cell lines Bifidobacterium longum (LMG 10502, biological origin: infant intestines) was obtained from the Belgian Coordinated Collection of Microorganisms (BCCM/LMG). B. longum was cultured in TSB at 37°C in an anaerobic atmosphere, generated with the use of ANAEROGENTM COMPACT. Cells of the murine macrophage cell line RAW 264.7 (ATCC TIB-71) were kindly provided by Prof. H. W. Virgin, Washington University School of Medicine, St. Louis, MO, USA. The RAW 264.7 cells were cultivated in complete DMEM medium and grown at 37°C under a 5 % CO2 atmosphere. The complete DMEM medium contained Dulbecco’s modified Eagle’s medium (DMEM; Lonza, Walkersville, MD, USA) containing 10 % low-endotoxin fetal bovine serum (FBS; HyClone, Logan, UT, USA), 100 U/mL penicillin and 100 µg/mL streptomycin (Lonza), 10 mM Hepes (Lonza), and 2 mM L-glutamine (Lonza). 25 3.2.2 Viruses MNV-1, which was used as a surrogate for human NoV, was kindly provided by Prof. H. W. Virgin, Washington University School of Medicine, St. Louis, MO, USA. The virus lysate was prepared by Dr. Dan Li from LFMFP as described previously (Li et al., 2011). 3.2.3 Monitoring of the bacterial growth The growth of B. longum was monitored as described in part 3.1.3 except that the first passage was grown for 48 h before making a second passage. Samples of the second passage were taken at 0, 12, 24 and 48 h. For the viable bacteria numeration, the inoculated plates were incubated at 37°C for 48 h before counting the colonies. 3.2.4 Culturing of the macrophage cell line RAW 264.7 RAW 264.7 cells were maintained in low-endotoxin medium (DMEM with 10 % low-endo FBS, 1 % Hepes, 1 % penicillin/streptomycin, 1 % L-glutamine). The cells were put in 20 mL of this medium and grown at 37°C under a 5 % CO2 atmosphere in 75 cm2 tissue culture flasks. The cells were splitted when a confluence of 70-75 % was reached. The old medium was removed and 10 mL fresh DMEM medium was added to the flask before scraping off the cells. A single-cell suspension was made by sucking up the medium with cells and then forcefully squeezing it through a pipette tip pressed onto the flask surface. This was repeated 5 times. Finally, 1 mL (1:10 dilution) or 2 mL (1:5 dilution) of the cell suspension was transferred into a new flask, already containing 19 mL respectively 18 mL new fresh complete DMEM before pursuing cell growth. Cell growth always occurred in a CO2 incubator C150 (Binder GmbH, Tuttlingen, Germany). All other steps were performed in a MSC-Advantage™ Class II Biological Safety Cabinet (Thermo Fisher Scientific). 3.2.5 Plaque assay The plaque assay for MNV-1 was always executed in a MSC-Advantage™ Class II Biological Safety Cabinet. On the first day, RAW 264.7 cells were seeded into 6-well plates (3.5 cm diameter) at a density of 2 x 106 viable cells per well. The plates were rocked and put overnight in a CO2 incubator C150 at 37°C and 5% CO2. On the next day, tenfold serial dilutions in antibiotics-free complete DMEM (low endo DMEM, 10% low endo FBS, 1% Glutamine and 1% Hepes) of a viral-bacterial mixture were prepared in 24-well plates. 26 The next step comprised rapidly pouring out the medium of the 6-well plates and adding 500 µL of the desired diluted viral-bacterial sample in duplicate wells. The 6-well plates were incubated at room temperature for 1 h, manually rocking the plates every 15 min. After aspirating the inoculum, the cells in each well were overlaid with 2 mL 1.5% SeaPlaque agarose (42°C; Cambrex, Rockland, ME, USA) in antibiotics-free complete Eagle’s minimum essential medium (37°C; complete EMEM; EMEM [Lonza], 10% low endo FBS, 2% L-glutamine and 1% Hepes) at a 1:1 ratio. The plates were placed in a tissue culture incubator at 37°C and 5 % CO2 for 2 days. Overlaying the cells in each well with 2 mL of a 1:1 mixture of 1.5% SeaKem agarose (50°C; Cambrex, Rockland, ME, USA) and antibiotics-free complete EMEM containing 1% neutral red (Sigma Aldrich), allowed for visualizing the plaques after incubation for 4 h at 37°C and 5% CO2. The following formula was used to calculate the number of viruses in the original virus-bacteriamedium mixture: X = log(A ∗ V) X equals the log number of PFU per mL in the original mixture, A represents the total number of counted plaques in the duplicate wells and V is the dilution factor. 3.3 Statistical analysis Statistical analyses of the results were performed by independent samples t-tests with SPSS Statistics 23 for Windows (SPSS Inc., Chicago, IL, USA). Significant differences were considered when p was smaller than 0.05. 27 28 Chapter 4: Results 4.1 Interaction between Norovirus and HBGA expressing bacteria 4.1.1 Optimisation of the HBGA expression test In the previous study of our research group (Li et al., 2015), the HBGA expression of bacteria was identified by flow cytometry analysis with the use of BDTM LSRII flow cytometer at INSERM, Nantes, France. In this study, due to the inaccessibility of this equipment, a substitutive method based on the principle of ELISA was attempted. Therefore, primary tests had to be performed in order to optimise the experimental conditions. 4.1.1.1 Concentration of antibodies Different concentrations of the primary and secondary antibodies were examined. For the primary antibodies (mouse anti-A antibodies #21), dilutions 1:250 and 1:1,000 were made in 0.1% BSA-PBS. Secondary anti-mouse antibodies were diluted to concentrations 1:2,500 and 1:10,000 in 0.1% BSAPBS. The effect of the four possible combinations of these primary and secondary antibody dilutions were compared in HBGA expression tests. Besides the detection of HBGA on E. coli LMG 8223 (previously confirmed as an HBGA expression positive strain), two types of negative controls were also included in parallel. Control 1: Detection of E. coli LMG 8223 without the addition of primary antibody, in order to control the non-specific binding of E. coli LMG 8223 with the secondary antibody; Control 2: Detection of E. coli ATCC 8739 (previously confirmed as an HBGA expression negative strain), in order to control the binding of non-HBGA materials with primary and/or secondary antibody. The results of the corresponding HBGA tests are given in Table 4. Combinations 1 and 3 gave the highest OD450 values for E. coli LMG 8223, but the negative controls also had a high OD value (higher than 1). This indicates that nonspecific binding took place. Combination 4, having the highest dilutions of both antibodies, gave a rather small difference between the positive value and the negative controls. Combination 2 had low values for the controls, the positive value was high and the difference was big enough to make a good differentiation. This makes combination 2, with primary antibodies diluted 250 times and secondary antibodies 10,000 times, the desired combination for further HBGA tests. 29 Table 4: OD450 values indicating HBGA expression with different antibody concentrations (primary antibody:secondary antibody). 1 = combination 1:250 and 1:2,500; 2 = combination 1:250 and 1:10,000; 3 = combination 1:1,000 and 1:2,500; 4 = combination 1:1,000 and 1:10,000. Control 1: Detection of E. coli LMG 8223 without the addition of primary antibody. Control 2: Detection of E. coli ATCC 8739. Combination 1 Combination 2 Combination 3 Combination 4 3.122 1.602 1.001 1.859 0.860 0.490 1.964 1.388 0.984 1.113 0.984 0.538 E. coli LMG 8223 Control 1 Control 2 4.1.1.2 Concentration of blocking reagent BSA was used in the HBGA expression test to prevent non-specific bindings. This test was performed to determine the appropriate concentration of BSA as a blocking reagent. Similar with section 4.1.1.1, two types of negative controls were included in parallel. Control 1: Detection of E. coli LMG 8223 without the addition of primary antibody; Control 2: Detection of E. coli ATCC 8739. The results are shown in Table 5. The group with the use of 0.1% BSA as blocking reagent induced a better differentiation between positive detection and negative controls than the use of 1% BSA. It could be possible that the higher amount of BSA also covered some of the HBGA receptors so that these carbohydrates were not recognisable for the primary antibodies. Therefore 0.1% BSA was chosen as blocking reagent concentration for further HBGA tests. Table 5: OD450 values indicating HBGA expression with different blocking reagent concentrations. Control 1: Detection of E. coli LMG 8223 without the addition of primary antibody. Control 2: Detection of E. coli ATCC 8739. E. coli LMG 8223 Control 1 Control 2 1% BSA 0.1% BSA 0.959 0.506 0.160 1.556 0.775 0.377 4.1.1.3 Cell numbers of the tested bacteria The amount of cells in a bacterial solution will influence the outcome of the HBGA expression test. To investigate this effect, cultures of E. coli LMG 8223 and E. coli LFMFP 861 (both previously confirmed as HBGA expression positive strains) were grown for 6 h and 24 h in aerobic atmosphere. For this HBGA test, only one type of control was carried out: Detection of E. coli LMG 8223 without the addition of primary antibody. The test results are presented in Table 6. Although the OD450 values were higher for the 24 h cultures, the difference between positive detection and the negative control was rather low. A bigger 30 differentiation was acquired with the 6 h samples. The optical density at 600 nm which indicates the bacterial cell mass is also given in Table 6. The 6 h cultures had an OD600 of around 0.3. So in the following tests, the bacteria used in the HBGA expression test were always normalised to an OD600 of 0.3, since this concentration is expected to give a good distinction between positive and negative results for the HBGA expression. Table 6: OD values at 450 nm (HBGA expression) and 600 nm (cell mass) of E. coli LMG 8223 and E. coli LFMFP 861 cultures. Bacteria were grown for 6 h and 24 h in aerobic conditions. Control: Detection of E. coli without the addition of primary antibody. Each time the mean and standard deviation of three independent tests are given. OD 450 nm (HBGA expression) E. coli LMG 8223 E. coli LFMFP 861 6h 6 h, control 24 h 24 h, control 1.45 ± 0.08 0.61 ± 0.08 1.71 ± 0.29 1.48 ± 0.10 0.81 ± 0.05 0.49 ± 0.04 0.96 ± 0.05 0.88 ± 0.04 0.32 ± 0.02 0.67 ± 0.09 0.27 ± 0.01 0.56 ± 0.01 OD 600 nm (cell mass) 6h 24 h 4.1.2 Influence of gas atmosphere of bacterial growth on their HBGA expression and viral protective effects E. coli are facultative anaerobe bacteria. This means that they are capable of growing in both aerobic or anaerobic conditions. In this section, the goal is to audit if the amount of HBGA expression and the protective effect on viruses can be linked to the gas environment wherein the bacteria grow. 4.1.2.1 Growth of E. coli LMG 8223 at different atmospheres First of all, growth of E. coli LMG 8223 was followed in aerobic (shaking at 200 rpm, 37°C) and anaerobic (ANAEROGENTM COMPACT, 37°C) gaseous atmospheres. The number of viable bacteria (colony forming units per mL, CFU/mL) and the cell mass (OD600) were measured at different points in time: after 0, 3, 6, 12 and 24 h. The data are shown in Figure 11 for aerobic growth and Figure 12 for anaerobic growth. In both cases, considering the viable bacteria numbers, a plateau phase was reached after 6 h of growth. The total number of living bacteria stayed approximately constant from this point onward and was similar for aerobic and anaerobic growth. Even though both atmospheres showed a similar number of alive cells, the OD600 is much higher in the aerobic case. This difference can be explained by assuming that E. coli cells die faster in aerobic conditions and that these dead cells help increase the turbidity of the solution. 31 For further experiments on the influence of the gas atmosphere, a similar starting number of bacteria was pursued so that it would be easier to compare results. Based on the two figures, the choice was made to let the bacteria grow aerobically for 6 h or anaerobically for 24 h. In these conditions, a similar OD600 (0.3) and number of CFU/mL (9 log) was reached. 10 0,8 9 0,6 8 0,4 7 0,2 6 5 OD 600 nm Concentration (log[CFU/mL]) Aerobic growth E. coli LMG8223 0 0 5 10 15 20 25 Time (h) log(CFU/mL) OD 600 nm Figure 11: Growth of E. coli LMG 8223 in TSB over 24 h in aerobic atmosphere (shaking at 200 rpm) at 37°C, visualizing the log(CFU/mL) with circles and OD600 with triangles. 10 0,8 0,7 0,6 0,5 0,4 0,3 0,2 0,1 0 9 8 7 6 5 0 5 10 15 20 25 OD 600 nm Concentration (log [CFU/mL]) Anaerobic growth E. coli LMG8223 30 Time (h) log(CFU/mL) OD 600 nm Figure 12: Growth of E. coli LMG 8223 in TSB over 24 h in anaerobic atmosphere (ANAEROGENTM bags) at 37°C, visualizing the log(CFU/mL) with circles and OD600 with triangles. 32 4.1.2.2 HBGA expression of E. coli LMG 8223 and LFMFP 861 grown at different atmospheres In a next step in investigating the influence of gas, HBGA expression tests were performed with E. coli LMG 8223 and LFMFP 861 that were cultured for 6 h in aerobic conditions and 24 h in anaerobic conditions. The control comprises the detection of E. coli without addition of primary antibodies. For both growth conditions, the OD600 values were around 0.3 (Table 7). Assuming this means a comparable number of cells was present, HBGA tests were performed. In the same table, also the results of the HBGA expression test are given. Comparing the two different gas atmospheres, E. coli LMG 8223 gave a significantly higher HBGA expression for the 24 h anaerobe samples (p value 0.009), however E. coli LFMFP 861 showed a significantly higher HBGA expression for the 6 h aerobe samples (p value 0.034). No consistent conclusions on the effect of the different atmosphere could be drawn from these results for both strains. Table 7: OD values at 450 nm (HBGA expression) and 600 nm (cell mass) of E. coli LMG 8223 and E. coli LFMFP 861 cultures. Bacteria were grown for 6 h in aerobic atmosphere or 24 h in anaerobic atmosphere. Control: Detection of E. coli without the addition of primary antibody. (*) indicates a significantly higher result between the aerobic and anaerobic samples. Each time the mean and standard deviation of three independent tests are given. OD 450 nm (HBGA expression) E. coli LMG 8223 E. coli LFMFP 861 6 h aerobic 6 h aerobic, control 24 h anaerobic 24 h anaerobic, control 1.45 ± 0.08 0.61 ± 0.08 1.87 ± 0.13 (*) 0.57 ± 0.07 0.81 ± 0.05 (*) 0.49 ± 0.04 0.64 ± 0.07 0.43 ± 0.02 0.32 ± 0.02 0.30 ± 0.04 0.27 ± 0.01 0.26 ± 0.01 OD 600 nm (cell mass) 6 h aerobic 24 h anaerobic 4.1.2.3 Viral protective effects of E. coli LMG 8223 and LFMFP 861 grown at different atmospheres A) Direct ELISA Previous experiments have demonstrated that the binding of NoV VLPs to HBGA expressing bacteria protects the VLPs from a heat treatment (Hirneisen & Kniel, 2013; Li et al., 2015). This study aims to find out whether there is a difference in this protective role if the bacteria are grown in different atmospheres. The direct ELISA test is performed in order to measure the antigenicity of the VLP antigens after heat treatment of 2 minutes at 90°C. The results of the test for E. coli LMG 8223 and LFMFP 861 are listed in Figure 13. 33 For both E. coli strains, detection of the VLP epitopes increased after heat treatment in the presence of the bacteria. However, no significant difference was present for E. coli LMG 8223 grown in different atmospheres (p value 0.134). E. coli LFMFP 861 gave a significantly higher result after heat treatment when the bacteria were grown in anaerobic conditions compared to aerobic conditions (p value 0.006). Similarly, no consistent conclusions on the effect of the different atmosphere could be drawn from these results for both strains. Direct ELISA 2,50 OD 450 nm 2,00 1,50 1,00 0,50 0,00 6h aer + VLP 6h aer + VLP, after 24h anaer + VLP 24h anaer + VLP, heat after heat E. coli LMG 8223 Only VLP Only VLP, after heat E. coli LFMFP 861 Figure 13: Protection of HBGA-expressing bacteria (E. coli LMG 8223 [dots] and E. coli LFMFP 861 [striped]) on the antigenicity of human NoV GII.4 VLPs towards heat treatment (90°C for 2 min). Antigen detection of NoV GII.4 in the presence of 6 h cultured bacteria in aerobic conditions (TSB, 37°C), 24 h cultured in anaerobic gas atmosphere (TSB, 37°C) or in absence of bacteria; each time before and after heat treatment. Every bar is an average of three independent tests, and every error bar represents the data range. B) Mucin-binding ELISA The second test to investigate the protective role of bacteria grown in different atmospheres is a mucin-binding ELISA. Porcine gastric mucin contains a mixture of HBGAs. This test determines if the VLPs lose their ability to bind receptors (HBGAs) after the same heat treatment as before: 2 min at 90°C. The same two E. coli strains were tested as for the direct ELISA test. The results are presented in Figure 14. Heating decreased the mucin-binding ability drastically. However, the measured optical density was always slightly higher in the tests with bacteria. When E. coli LMG 8223 was used, the aerobic cultures could preserve the binding ability of the VLPs better than anaerobic cultures (p value 0.001). For E. coli LFMFP 861, no significant difference could be made between incubation of the VLPs with aerobic or 34 anaerobic cultures (p value 0.374). Still, no consistent conclusions on the effect of the different atmosphere could be drawn from these results for both strains. Mucin-binding ELISA 2,00 OD 450 nm 1,60 1,20 0,80 0,40 0,00 6h aer + VLP 6h aer + VLP, after 24h anaer + VLP 24h anaer + VLP, heat after heat E. coli LMG 8223 Only VLP Only VLP, after heat E. coli LFMFP 861 Figure 14: Protection of HBGA-expressing bacteria (E. coli LMG 8223 [dots] and E. coli LFMFP 861 [striped]) on the receptor binding ability of human NoV GII.4 VLPs towards heat treatment (90°C for 2 min). Mucin-binding ability detection of NoV GII.4 in the presence of 6 h cultured bacteria in aerobic conditions, 24 h cultured in anaerobic gas atmosphere or in absence of bacteria; each time before and after heat treatment. Each bar is an average of three independent tests, and each error bar represents the data range. 4.2 Interaction between Norovirus and probiotic bacteria 4.2.1 B. longum growth 4.2.1.1 Growth in bacteria culture medium TSB The anaerobic growth of B. longum in bacteria culture medium TSB at 37°C was followed (Figure 15). ANAEROGENTM bags created the anaerobic atmosphere. The number of alive B. longum cells showed a noticeably increase after 12 h of incubation. The bacteria culture could be considered as fully grown after 24 h, when approximately 9 to 10 log CFU/mL were present and a plateau phase was reached. In addition, the cells mass remained constant from 24 h on. The OD600 measurements reached a value of 0.2 in this case. 35 12 0,4 10 0,3 8 6 0,2 4 OD 600 nm Concentration (Log CFU/mL) Growth of B. longum 0,1 2 0 0 0 5 10 15 20 25 30 35 40 45 50 Time (h) log(CFU/mL) OD 600 nm Figure 15: Growth of B. longum over 48 h, visualizing the log(CFU/mL) with circles and OD600 with triangles. The bacteria were anaerobically grown (ANAEROGENTM bags) at 37°C. 4.2.1.2 Growth in a food matrix Since dairy products are the main vehicles for probiotic supplementation, the choice was made to test skim milk as a food matrix model. Around 106 B. longum cells were inoculated in skim milk and incubated anaerobically at 37°C. After 48 h, 8 log CFU/mL were present in the milk culture (Table 8). This was still approximately 2 log less than growth in TSB. Consequently, an extra carbon source (5 % sucrose or 5 % honey) was added to the milk to stimulate bacterial growth. Adding sucrose resulted in the biggest increase of growth, almost 1 log CFU/mL. Therefore, the skim milk supplemented with 5% sucrose was chosen as the food model in the following tests. Table 8: The growth of B. longum cultured in different media. The bacteria were grown anaerobically for 48 h at 37°C. Log CFU/mL TSB Skim milk Skim milk + 5 % sucrose Skim milk + 5 % honey 9.86 8.00 8.95 8.60 4.2.2 Effect of B. longum on NoVs The survival of MNV-1 in TSB, milk with 5 % sucrose, and cell-free bacteria culture filtrate was firstly investigated. Bacteria cultures were centrifuged at 6000 rpm for 5 min in an Eppendorf centrifuge 5804 R (Novolab, Geraardsbergen, Belgium) and the supernatants were recovered and filtered over a 0.22 36 µm Millex-GV syringe filter (Merckx Millipore, Carrigtwohill, Co. Cork, Ireland) to remove residual bacteria and end up with the filtrate. In each case, 1 mL of the virus was added to 10 mL of the medium and anaerobically incubated for 48 h at 37°C before performing the plaque assays. The results are presented in Table 9. Incubating the virus with cell-free filtrate, containing secondary metabolites of a fully grown B. longum culture, delivered the highest viral reduction: 2.4 log PFU/mL. This is almost 1 log more than in TSB or milk with sucrose. Figure 16 displays an example of the plaque assay, with the transparent spots as plaques caused by MNV-1. Table 9: Results of the survival of MNV-1 in different media detected by cell plaque assays. Each time the mean log PFU/mL and standard deviation of three independent tests are given. 0h 12 h 24 h 48 h Log reduction after 48 h 10 mL TSB + 1 mL MNV-1 10 mL skim milk with 5 % sucrose + 1 mL MNV-1 10 mL cell-free bacteria culture filtrate (in TSB) + 1 mL MNV-1 4.15 ± 0.06 3.40 ± 0.09 3.06 ± 0.06 2.50 ± 0.17 6.60 ± 0.12 6.73 ± 0.12 5.17 ± 0.15 4.33 ± 0.35 1.65 ± 0.18 1.43 ± 0.19 2.40 ± 0.37 Figure 16: An example of the plaque assay, with the transparent spots as plaques caused by MNV-1. 37 The direct effect of B. longum cells on MNV-1 was investigated in the following. As a first test, 1010 B. longum cells were added to 1 mL of the virus. After 1 h of incubation at 37°C, no reduction in the number of PFU was found (Test 1, Table 10). This test was repeated with a lowered virus concentration, assuming MNV-1 might become more susceptible for the treatment. However similarly, only a very small reduction was observed (Test 2, Table 10). Table 10: Effect of B. longum on MNV-1 with 1010 bacterial cells, incubated anaerobically for 1 h at 37°C. MNV1 lysate was used for Test 1, 1,000 times diluted virus lysate for Test 2. Each time the mean log PFU/mL and standard deviation of three independent tests are given. 1 mL MNV-1 + 1010 B. longum cells 0h 1h Log reduction after 1 h Test 1 Test 2 8.15 ± 0.06 8.58 ± 0.06 5.15 ± 0.06 4.89 ± 0.04 No reduction 0.26 ± 0.07 Since the effect of the bacteria after 1 h was very limited, the treatment time was prolonged up to 48 h in a next phase of the experiments. For Test 3, 1 mL of MNV-1 was added to 10 mL of a growing bacteria culture in TSB. Based on the bacterial growth indicated in Figure 14, bacteria numbers starting from 106 going up to 1011 after 48 h were expected. Test 3 showed only a 1.28 ± 0.27 log reduction in PFU/mL after 48 h. For similar reasons as with Test 2, the virus concentration was lowered to repeat the test. In this case (Test 4), a 1.74 ± 0.21 log reduction in PFU/mL occurred after 12 h, but no plaques were observed after 24 and 48 h. The uncountable results remained questionable, as it is known that higher B. longum cell numbers were present in the specific wells for plaque assay compared to other wells (with higher virus concentrations and therefore further dilutions or less bacteria culture time), and it is known based on our previous experience that the presence of B. longum in the plaque assay wells could inhibit virus multiplication, therefore inducing smaller and less plaques (data not published). The results of Tests 3 and 4 are shown in Table 11. From the countable results of the four tests described above, not a single one gave a reduction that was similar or higher to the one obtained with cell-free bacterial filtrate. The starting number of B. longum bacteria was increased to 1011 bacterial cells for the next tests. The bacteria were put together with 1 mL of MNV-1 and 10 mL of TSB, incubated anaerobically for 48 h at 37°C, and the plaque assay was performed subsequently in Test 5. All the same was done in Test 6, but this time milk with 5 % sucrose was the matrix of choice. Table 12 gives a summary of the results obtained by these two tests. Test 5 resulted in a 2.14 ± 0.29 log PFU/mL reduction after 48 h, whereas a procedure as in Test 6 led to a reduction that might be bigger than 5.73 log. 38 Table 11: Results of the cell plaque assays for MNV-1 in a growing bacteria culture in TSB, incubated anaerobically up to 48 h at 37°C. MNV-1 lysate was used for Test 3, 1,000 times diluted virus lysate for Test 4. Each time the mean log PFU/mL and standard deviation of three independent tests are given. 10 mL growing bacteria culture in TSB + 1 mL MNV-1 0h 3h 6h 12 h 24 h 48 h Log reduction Test 3 Test 4 5.86 ± 0.04 5.66 ± 0.02 5.58 ± 0.03 5.44 ± 0.13 5.13 ± 0.10 4.58 ± 0.27 4.76 ± 0.13 3.79 ± 0.10 3.40 ± 0.17 3.02 ± 0.16 <1? <1? 1.28 ± 0.27 log reduction after 48 h 1.74 ± 0.21 log reduction after 12 h; > 3.76 log after 24 h? Table 12: Results of the cell plaque assays for MNV-1 with 1011 bacterial cells, incubated anaerobically in different media for 48 h at 37°C. TSB was used for Test 5, milk with 5 % sucrose for Test 6. Each time the mean log PFU/mL and standard deviation of three independent tests are given. 10 mL matrix + 1 mL MNV-1 + 1011 B. longum cells 0h 48 h Log reduction after 48 h Test 5, in TSB Test 6, in milk + 5 % sucrose 6.73 ± 0.12 4.59 ± 0.26 6.73 ± 0.13 <1? 2.14 ± 0.29 > 5.73 ? 39 40 Chapter 5: Discussion 5.1 Interaction between Norovirus and HBGA expressing bacteria Previous research at LFMFP investigated the HBGA expression of multiple bacterial strains (Li et al., 2015). The study also indicated that if NoV VLPs were heated for 2 min at 90°C in the presence of HBGA expressing E. coli, a higher antigen integrity as well as mucin-binding ability of the VLPs was detected compared to the same test with the same cell numbers of non-HBGA expressing bacteria. This led to the conclusion that HBGA expressing bacteria can have a protective role on human NoV from acute heat stress. However, it was also noticed that large variations can occur in HBGA expression between different bacterial strains, even if they are closely related. Different cultural batches from one particular bacterial strain were susceptible to varying HBGA levels as well. HBGA-like structures can be secondary metabolites of bacteria and their production could be regulated by some currently unknown factors. In this thesis, further explorations were done to tackle the rising question of which factors are influencing HBGA expression levels. E. coli are facultative anaerobe bacteria, meaning that they are capable of growing in both aerobic or anaerobic environments. It has already been demonstrated that the production of secondary metabolites in E. coli can be influenced by the gaseous atmosphere in which the bacteria grow (Gray et al., 1966). In this study, the goal is to investigate the role or influence of the gaseous atmosphere on both the expression of HBGAs and the viral protective effect of the bacteria. Due to a limitation in the available facilities, flow cytometry could not be used to measure HBGA expression like was done in the previously mentioned article from Li et al. (2015). The first challenge was thus to create a new HBGA detection method. An ELISA test was chosen as a substitute. Therefore, the amounts of antibodies, blocking reagent and bacterial cell numbers had to be optimised in the first place. Based on the results of Li et al. (2015), the choice was made to use E. coli LMG 8223 and E. coli LFMFP 861 since they showed the highest HBGA expression and could both bind to NoV VLPs. Besides the ELISA-based HBGA expression test, direct ELISA assays were used to measure the NoV VLP antigen integrity after heat treatment and porcine gastric mucin-binding assays were performed with the prospect of measuring the receptor binding ability of the NoV VLPs after the same treatment. Table 13 gives an overview of the main findings from all the tests. Looking at the two strains separately, no consistent link could be made between the HBGA expression and the protection of the virus by the bacteria that were cultured in either an aerobic or anaerobic gaseous atmosphere. In addition, the 41 conclusions that were observed for E. coli LMG 8223 were not consistent with those for E. coli LFMFP 861. Table 13: Summary of the differences of aerobically or anaerobically grown E. coli LMG 8223 and E. coli LFMFP 861 on their HBGA expression on the one hand, and the NoV VLP antigen integrity and receptor binding ability after heat treatment on the other hand. The p values were calculated by an independent samples t-test that compared the means of the 24 h anaerobic and 6 h aerobic groups. HBGA expression NoV VLP antigen integrity after heat treatment Receptor binding ability after heat treatment E. coli LMG 8223 E. coli LFMFP 861 24 h anaerobic > 6 h aerobic p = 0.009 No significant difference p = 0.134 24 h anaerobic < 6 h aerobic p = 0.001 24 h anaerobic < 6 h aerobic p = 0.034 24 h anaerobic > 6 h aerobic p = 0.006 No significant difference p = 0.374 This inconsistency in the results might be described partially to the normalisation of bacteria. In this study, it was determined to normalise the optical density of a bacteria culture at 600 nm to 0.3 before each experiment. The purpose was to make sure that the cell mass was similar in each sample to be compared with. However as shown in the results, the growth dynamics of E. coli in aerobic and anaerobic atmospheres, determined by measuring the viable cell numbers and optical density respectively, were rather different. Therefore, although the cell mass was normalised, it is possible that other factors exist to influence the HBGA expression and NoV protective effects that we investigated. In this study, the HBGA expression was measured by an ELISA based method. Compared with the flow cytometry used previously (Li et al., 2015) which measures the fluorescence of the cells one by one, ELISA measures the total fluorescence signal generated from all the bacterial cells from one sample. Therefore, it is possible that bigger variation can be caused in the ELISA based method due to aggregation of bacterial cells or background noise. A clear link between the gaseous atmosphere and the level of HBGA expression that accounts for both E. coli strains was not found (see Table 13). However, it was noticed that after 24 h of aerobic growth, for both of the two bacterial strains, it was difficult to differentiate positive HBGA expression signals from the negative controls (too high background signals, Table 6). Greater differences were present for 24 h of growth in the anaerobic case (Table 7). It has been demonstrated in this study that the E. coli bacteria grow faster aerobically than anaerobically. It was also shown that the gaseous atmosphere can influence the production of secondary metabolites (Gray et al., 1966). Taken together, we can assume that certain secondary metabolites are earlier formed when the bacteria are grown aerobically, and some of these metabolites may as well have affinity towards the antibodies that would interfere with the detection of HBGAs. It is also possible that other factors apart from the gaseous 42 environment, such as the prevalent pH in the intestines of the human body, have a greater influence on HBGA expression. This can be the subject for further examination on the HBGA topic. The direct ELISA assay revealed that NoV VLPs gained a higher immunoreactivity after exposure to heat stress. This can be due to partial degradation of the VLPs when they are heated, causing otherwise masked epitopes to now reach the surface. The results from the direct ELISA and mucin-binding ELISA suggest that HBGA expressing bacteria can protect NoVs during heat treatment, even though these assays do not completely reflect an authentic NoV infection. This outcome is consistent with the findings of Li et al. (2015). 5.2 Interaction between Norovirus and probiotic bacteria Several key challenges remain in assessing the efficacy of vaccines and antiviral drugs for human NoV infections. There is for example no robust cell culture system for human NoVs, limiting the direct study of these viruses. Additionally, the genetic heterogeneity among NoV strains makes it difficult to find a good vaccine. Consequently, the use of probiotics as a possible method to control and treat a NoV infection is an interesting topic for investigation. Nowadays, research on the effect of probiotic bacteria on NoVs is still in a preliminary phase. This is mainly due to two reasons. First of all, as a result of the non-cultivability of human NoVs, surrogates are used to conduct in vitro experiments (e.g. FCV, Aboubakr et al., 2014; MNV, Lee et al., 2012; TV, Shearer et al., 2014; NoV P particles, Rubio-del-Campo, 2014). Secondly, because of a lack of enough in-depth studies on this topic, it is difficult to draw general conclusions. As previously shown in part 2.2.3, the decrease in viral infectivity caused by probiotic bacteria can be very variable. The inconsistency in the results could be caused by differences between the used LAB strains and viruses (or its surrogates), as well as the experimental set-ups. The purpose of this study was to investigate the potential antiviral effect of B. longum against NoVs. MNV-1 was employed as a surrogate. B. longum is a recognized probiotic (Tamaki et al., 2016; Vieira et al., 2016) and it was also reported to show antiviral effect against rotavirus (Kang et al., 2015). The survival of MNV-1 in different media without bacteria was firstly tested. In a next phase, the potential antiviral effect of B. longum was examined by looking into the outcome of multiple co-incubation setups with the bacteria and virus. Skim milk was chosen as a food matrix, because it is relatively easy to define its compounds and dairy products are known to be the main vehicles of probiotic supplementation. Bifidobacteria are normal inhabitants of the human gastrointestinal tract and when they are added to foods, exterior 43 environmental factors (e.g. the compatibility with the starter lactic acid culture, oxygen stress, acid stress, etc.) can greatly influence the stability and functionality of these bacteria (reviewed by Granato et al., 2010). Indeed, in this study, B. longum grew slower in skim milk compared with the growth in TSB. Previous research reported enhanced growth by bifidobacteria grown in skim milk in the presence of carbohydrate components such as honey, fructooligosaccharides, and galactooligosaccharides (Shin et al., 2000; Ustunol & Gandhi, 2001; Ustunol, 2000). Based on a preliminary test, 5% of sucrose was added into the slim milk in this study to support the growth of B. longum. Based on the results in this research, it was indicated that the direct effect of B. longum is limited. Within all the countable results of the plaque assays, the highest reduction was obtained with cell-free bacteria culture filtrate. Even with high bacterial cell numbers, not much difference was noticed compared to the filtrate. This filtrate contained secondary metabolites that were secreted by the bacteria. It is known that probiotic bacteria can produce metabolites such as organic acids (lactic and acetic acid), hydrogen peroxide, ethanol, diacetyl, acetaldehyde, carbon dioxide, reuterin, and bacteriocins (Suskovic et al., 2010). There is not much information available concerning bacteriocins produced by B. longum. Kang et al. (1989) described the bacteriocin bifilong, Saleh and El-Sayed (2004) informed about bifilong Bb-46, and Lee et al. (2011) identified the lantibiotic bisin. Some characteristics that were determined for these B. longum bacteriocins are given in Table 14. Recently, metabolites of B. adolescentis were shown to have an antiviral effect on rotavirus by decreasing the production of the viral protein NSP4, being a viral enterotoxin protein (Olaya Galán et al., 2016). What specific metabolites caused this reduction and how they did it, was not mentioned. No studies have been performed yet that investigated a possible antiviral effect of B. longum metabolites. Table 14: Bacteriocins from B. longum and their main characteristics. (Martinez et al., 2013) Bacteriocin Species Mol. wt. and strain (kDa) Heat range stability pH range stability Production phase Inhibitory spectrum Bifilong B. longum 120 (100°C – 30 min) 2.5 – 5.0 (-) Bifilong Bb46 B. longum Bb-46 25-127 (121°C – 15 min) 4.0 – 7.0 (-) Lantibiotic (Bisin) B. longum DJO10A (-) (-) (-) 1–8h gram-positive & gram-negative bacteria Staphylococcus aureus, Salmonella typhimurium, Bacillus cereus, Escherichia coli Streptococcus thermophilus ST403, Clostridium perfringens, Staphylococcus epidermidis, Bacillus subtilis, E. coli DH5a. Reference Kang et al. (1898) Saleh & El-Sayed (2004) Lee et al. (2011) 44 Incubation of the bacteria with the virus for 1 h resulted in no or only very small viral reductions. Because of this, the decision was made to prolong the treatment time. It was not known beforehand if and when a significant antiviral effect could happen, so multiple incubation durations were tested ranging from 3 h up to 48 h. A NoV infection is self-limiting within two to three days. Testing treatment times longer than 48 h would thus not be very useful, because a NoV infection might already be over again by that time. From a clinical point of view, it would be most interesting if a meaningful viral reduction could be found at low treatment periods. It was demonstrated in this study that the presence of B. longum could inhibit the multiplication of MNV-1 on RAW 264.7 cells. The experiments support the idea that B. longum does not show direct virucidal effect or prevent the viruses from binding to the cells, but that it is more likely that the viral replication phase would be influenced. Several reasons can be brought up to confirm this statement. First of all, previous research from our laboratory has revealed that the presence of probiotic bacteria could induce smaller plaques in the plaque assay of MNV-1 (Li et al., unpublished). When uncountable results occurred in the test, always a high number of bacterial cells were present in the cell culture wells for plaque assay. In addition, in Test 4, for the groups incubated with 12 h cultured bacteria, the plaques were found to be too small to count in some replicates. In Test 5, where MNV-1 was coincubated with 1011 bacterial cells in TSB, plaques could only be counted when a ten-thousand-time dilution of the original viral-bacterial sample was made. Lower dilutions did not allow for counting plaques, since they were too small to be seen. Co-incubation of MNV-1 with the highest number of bacterial cells in milk supplemented with sucrose led to viral numbers below the detection limit. However, in this case it was not sure whether the reduction was caused by the antiviral effect of B. longum, by the viral inhibiting effect of certain milk components, or by a combination of these two. Some milk proteins were reported to interfere with viral infections, which might lead to greater virus inhibition effect in milk. Lactoferrin could for instance prevent viral entry of host cells by blocking cellular receptors or by binding to the virus particles (Pan et al., 2006). In addition, it was found that human milk glycans inhibits binding between NoV and its host glycan receptor (Shang et al., 2013). This might be an interesting lead to combat with NoV infection in young children. Rotaviruses and NoVs cause gastroenteritis and the symptoms of both infections are similar. Although some of the articles mentioned in Table 2 studied the effect of probiotics on rotaviruses, these viruses cannot be considered as real surrogates for NoVs. The viruses differ too much in their virion structure and genome structure (Estes & Cohen, 1989). As a result, mechanisms that are effective against rotaviruses might not always work against NoVs. 45 46 Chapter 6: General Conclusions Overall we observed that growing the bacteria in different gas atmospheres had indeed influenced the HBGA expression as well as the protection of the integrity and functionality of NoV VLPs during heat stress. However, performing the tests for two different E. coli strains always led to different conclusions that could be drawn. So in the end, the inconsistency in these results makes it impossible to make general conclusions that account for both strains. The gas atmosphere might not be the biggest factor influencing HBGA expression and virus protection. Perhaps for further research on this topic, attention could be paid to the influence of pH. As the bacteria should be capable of passing through the stomach and intestines, it might be useful to mimic this route and expose HBGA expressing bacteria to fluids similar to those present in the stomach (pH 1.5 - 3.5) and small intestines (pH 6.8), before checking their receptor expression and virus protection ability. Currently, research on the effect of probiotics on NoVs is still preliminary. This is mainly caused by the non-cultivability of human NoVs and the lack of sufficient in-depth studies on this topic. This study intended to demonstrate the antiviral effect of B. longum against NoVs. The direct effect was found to be limited. The actual effect should mainly be found in the replication phase of the virus. 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