PEER-REVIEWED ARTICLE
Food Protection Trends, Vol 33, No. 5, p. 290–299
Copyright©2013, International Association for Food Protection
6200 Aurora Ave., Suite 200W, Des Moines, IA 50322-2864
Norovirus Attachment:
Implications for Food Safety
Kirsten A. Hirneisen and Kalmia E. Kniel*
Dept. of Animal and Food Sciences, University of Delaware, Newark, DE 19716, USA
ABSTRACT
Human noroviruses are the leading cause of foodborne gastroenteritis in the United States. Recent research has focused on norovirus attachment to
host-cell carbohydrates. This area of research increases understanding of the first step of norovirus replication in host cells; it also has been extended to
explaining norovirus capsid structure and virus evolution, as well as norovirus attachment to foods and has been used in developing methods of detecting
human noroviruses. This article provides a review of research conducted on human norovirus attachment, specifically on the attachment of noroviruses on
foods, including original research addressing the role of spinach and green onions in disease caused by human norovirus and a common human norovirus
surrogate, murine norovirus. The objective of this study is to assess the applicability of an ELISA attachment assay to predict infectivity of a genogroup II
human norovirus and to determine if spinach and green onion homogenates interfere with the attachment of the virus to the carbohydrate coating. Attachment
assay results for heat-treated human norovirus indicated decreasing attachment with increasing heat treatment, which likely correlates with loss of infectivity.
Additionally, the presence of spinach and green onion homogenate did not affect attachment of human norovirus or murine norovirus to porcine gastric mucin.
*Author for correspondence: Phone: +1 302.831.6513; Fax: +1 302.831.2822; E-mail: [email protected]
290 FOOD PROTECTION TRENDS SEPTEMBER-OCTOBER 2013
INTRODUCTION
Human noroviruses cause 58% of gastroenteritis in the United
States and are suspected to be the leading cause of foodborne illness
(37). Noroviruses typically cause a self-limiting acute diarrheal disease
lasting 2–3 days; however, more serious cases have been responsible
for 26% of hospitalizations and 11% of deaths caused by foodborne
illnesses (37). Although the ways in which noroviruses infect human
gastrointestinal cells are not completely understood, in 2002, human
noroviruses were discovered to bind to histo-blood group antigens
on human gastrointestinal cells (32). This finding opened a door to
research aimed at understanding the beginning stages of norovirus
replication, structural differences and evolutionary strategies and
applying this understanding to development of more precise detection
methods as well as evaluation of human norovirus attachment to foods.
Norovirus attachment to histo-blood group antigens
The initial steps of virus infection involve attachment of the virus
to host cellular receptors, followed by entry of the virus through the
cell membrane and subsequent release of the genetic material into the
cytoplasm. Carbohydrate moieties of host cell glycoproteins, glycolipids
such as sialic acid, and proteoglycans comprise a widely used strategy
of virus attachment to epithelial cells (5). The attachment of virus
to host cellular receptors involves recognition of the specific cellular
receptor. Histo-blood group antigens (HBGA) are receptors on human
gastrointestinal cells to which NoV binds (19, 20, 32). Often, viral
penetration is mediated by a membrane protein that interacts with the
virus specifically as a receptor or co-receptor, for example, the norovirus
surrogate feline calicivirus binds to sialic acid on host cells (39) and
also to a cellular membrane protein, junctional adhesion molecular 1
(JAM1) (31, 34). With human noroviruses, a 105-kDa membrane protein,
designated as NV attachment (NORVA) protein, present in mammalian
cell lines including Caco-2 cells was shown to interact with human
norovirus-like particles (40). The function of this protein-protein
interaction between NORVA and NoVLPs has yet to be defined (43) and
attempts to extract and purify the protein have been unsuccessful (41).
Knowledge of HBGA binding patterns of human noroviruses have
been elucidated through studies using in vitro binding and blocking
assays with recombinant virus-like particles, human volunteer
challenge studies, and outbreak investigations (43). Histo-blood group
antigens, a major group of carbohydrates that are determinants of
human and animal blood types, are abundant in bodily fluids including
blood, saliva and milk. HBGAs, found on cellular surfaces and mucosal
epithelium, are complex carbohydrates present on the outermost
ends of N- or O- linked glycans or glycolipids (43). Different norovirus
strains have varying receptor-binding profiles associated with the ABO,
secretor, and Lewis HBGA types as determined by human volunteer
challenge studies (43). In vitro binding assays revealed that most
GII.4 strains recognized saliva of all ABO secretors, which represent
approximately 80% of the world’s population, indicating an important
reason for the predominance of this genotype over those that have a
narrower target population (43). Of all norovirus genotypes tested, GII.4
was observed to bind to the widest variety of HBGAs (19).
In addition to HBGAs, human noroviruses have been shown to bind
to porcine gastric mucin (PGM) (48) and heparan sulfates (41). Porcine
gastric mucin (PGM) contains type A, type O and type Lewis b HBGA, to
which NoV has been shown to bind (48, 49). PGM also contains protein
(20%) and carbohydrates such as hexosamine (37%), hexoses (27%),
fucose (10%) and sialic acid (6%) (14). Studies using PGM to assess
the mechanism of norovirus attachment showed that the binding of
NoVLP to PGM was completely inhibited after periodate oxidation of
carbohydrates at all concentrations, while boiling the proteins had no
effect (14). This provided further confirmation that NoV attachment to
PGM occurs through the carbohydrate moieties and not through proteinprotein interactions (49).
Norovirus capsid structure and interactions with antibodies and
receptors
Following the studies linking noroviruses to various cell surface
receptors, X-ray crystallography, cryoelectron microscopy, NMR and
genetic mutagenesis techniques were used to advance knowledge of
the crystal structures and binding interfaces of noroviruses with host
cellular receptors (4, 6, 10, 11, 42, 45, 48). The major structural protein
of noroviruses, VP1, is composed of two domains, the shell (S) domain,
which forms the core of the icosahedral shell (3), and the protruding (P)
domain, which forms arches extending from the shell. The P domain,
located at the most exterior surface of the viral particle, is responsible
for host interactions and the immune response (6). The P domain is
composed to two subdomains, P1 and P2, which correspond to the leg
and head of the arch-like P dimer (35). Binding has been observed
to occur at the outer pocket of the P domain protrusions (6, 42). This
HBGA-binding pocket includes a group of scattered amino acids that
interacts with the oligosaccharide residues of the HBGA receptor
(6, 10, 11).
Through comparing the structures of the P domain bound to A
trisaccharides, evidence of different receptor-binding motifs were
observed for both GI and GII noroviruses (6). Bu et al.(6) determined
that both GI and GII noroviruses bound to the same A and H antigens
and the binding interfaces were located on the same region of the
P2 domain, but the amino acids involved were completely different.
Sequence alignment of noroviruses showed that key residues for HBGA
binding are highly conserved among strains within the genogroups,
but not between two genogroups; the remaining sequences of the P2
subdomain are highly variable (46). This suggests that interactions
with HBGAs play an important role in the evolution of noroviruses (44).
Norovirus attachment in foods
Knowledge gained from attachment research has also focused
on the possible attachment of human noroviruses to foods. Because
noroviruses are transmitted through the fecal-oral route, it would be
of evolutionary advantage for the viruses to bind easily to a broad range
of vehicles, including foods, so as to be spread more efficiently (26).
Additionally, the attachment of noroviruses to foods likely promotes
persistence and survival. The study of viral attachment to foods has
presently focused on two groups of foods: shellfish, particularly oysters,
and produce, particularly romaine lettuce. Both of these groups are
commonly implicated in norovirus outbreaks worldwide (1, 2, 7, 15,
29, 33, 57, 62). Understanding virus attachment mechanisms is
key to developing effective detection methods as well as optimizing
intervention strategies.
SEPTEMBER-OCTOBER 2013 FOOD PROTECTION TRENDS 291
Early efforts to understand the mechanism of virus attachment
to lettuce focused on electrostatic forces, in particular the role of the
isoelectric point of the virus (58, 59). Virus attachment to lettuce
was first assessed by Vega et al. (59) using bacteriophages MS2 and
ΦX174, feline calicivirus, and echovirus 11 at various pH levels. The
authors hypothesized that the attachment of these viruses to lettuce
would depend on the isoelectric point of the virus, because the ultimate
charge of a virus would depend on the amino acid residues on the
virus surface and on the pH of the surrounding medium. The four
viruses had varying attachment patterns to lettuce despite having
similar isoelectric points, which indicated that the isoelectric point
of the virus was not the factor governing their attachment to lettuce.
The authors then compared viral attachment to lettuce with Tween
80 added and NaCl added, to test for hydrophobic interactions and
electrostatic interactions, respectively (58). Vega et al.(58) concluded
that electrostatic forces are the primary force involved in the adsorption
to lettuce.
Since the discovery of norovirus attachment to histo-blood group
antigens, research on the attachment of enteric viruses on foods
has shifted from non-specific to specific interactions. Early on, the
attachment of poliovirus to oysters was shown to involve binding of the
virus particles to carbohydrate moieties (13). Similarly, GI NoV were
also found to bind to carbohydrates in oysters, specifically histo-blood
group antigens present in the gastrointestinal cells of oysters (24, 48,
51), as observed in ELISA attachment assays and confocal microscopy.
Oyster stomach and digestive tissues were reported to contain type
A-like HBGAs to which NoVs bind (24, 48). Type A-like HBGAs have been
reported to bind to multiple NoV stains (21), suggesting that other
strains of NoVs bind to oysters. In competitive binding assays, human
saliva type A HBGA and anti-type A HBGA MAb inhibited the binding
of NoVLPs to the oyster stomach and digestive tissues. Specifically in
oysters, enteric viruses, including MNV and HAV, have been observed to
bind to phagocytic blood cells (hemocytes) (36). These results suggest
that NoV binding to oyster tissue is mediated by specific ligandreceptor interactions and not just physical entrapment. Through this
HBGA binding, it is hypothesized, oysters bio-accumulate NoVs during
the course of filter feeding (24, 51). Mutant human norovirus virus-like
particles (NoVLPs) with an alanine substitution in the P2 domain of VP1
prevented the binding to oyster tissue (24), suggesting similarities to
the mechanisms of recognition of human tissues by NoVs.
Tian et al.(51) assessed NoV binding to HBGAs in a variety of
bivalve filter feeders, including a range of oyster species, clams
and mussels. Through ELISA binding assays using MAbs specific for
different HBGAs, it was revealed that both type A-like and type O-like
HBGAs were present in all oyster species and in Manila clams; however,
blue mussels contained only type A-like HBGAs. Although both type
A-like and type O-like HBGAs are present, competitive assays indicated
that type A-like HBGAs appear to have a dominant role in the NoVLP
binding in oysters (51). The genetic diversity of NoVs, highlighted by
differences between GI and GII binding to human HBGAs, is similarly
observed in the binding capacity to HBGA structures within oysters. GI
NoVs bind to type A-like HBGAs in oyster digestive tissues, whereas GII
NoVs, in addition to binding to digestive tissue by a type A-like HBGA,
are also able to bind to gills and mantle tissue sections by a sialic acid
α2,3 linkage (30).
292 FOOD PROTECTION TRENDS SEPTEMBER-OCTOBER 2013
The role of seasonality and glycogen levels in NoV attachment
to oysters was hypothe-sized, as oyster glycogen is present in high
concentrations from November to March, which coincides with the time
of seasonal outbreaks of norovirus infections (24, 52). In assessing the
role glycogen plays in norovirus binding, it was observed that norovirus
attachment to oysters does not have a seasonal pattern (52). Contrary
results were observed by Maalouf et al. (30); GI VLPs were able to bind
more much efficiently to oysters during the first five months of the
year than during the rest of the year, and GI strains are most regularly
involved in oyster-related outbreaks from January to May.
Wei et al. (61) observed, by use of confocal microscopy, that
SYBR gold labeled MNV attached to Romaine lettuce surfaces. Virus
was pipetted directly onto lettuce or the lettuce was agitated in
virus suspension, and the virus was found in stomata, along the cut
edges and on leaf surfaces; biosolids were shown to promote this
attachment of MNV to lettuce (61). Porcine sapovirus, as a surrogate
for human noroviruses, were also found to attach to Romaine lettuce
and remain infectious to lettuce one week after storage at refrigeration
temperatures (60). Confocal microscopy has also shown that NoVLPs
localize in clusters along the veins rather than being equally distributed
through the leaf (16). The specific attachment of NoVLPs to romaine
lettuce extract was further elucidated through ELISA methods (16).
Extracts of Romaine lettuce leaves coated to ELISA plates were observed
to bind the NoVLPs in a dose-dependent manner, but Romaine lettuce
extract did not bind the norovirus-like particles by HBGA. Additionally,
Romaine lettuce extract was not competitive with NVLP binding to PGM,
suggesting that non-HBGA molecules in romaine lettuce bind to NoVLPs
via binding sites different from the defined binding pocket of the virus
particle. Proteins were also suggested to have some role in the binding
to NoVLP to Romaine lettuce exudate, as binding was increased slightly
by oxidation but decreased slightly by boiling.
To further understand the attachment of human noroviruses to
Romaine lettuce, Esseili et al. (14) assessed the binding of NoVLPs to
cell wall material carbohydrates of romaine lettuce leaves. Through
studies of oxidation of carbohydrates and boiling of proteins, it was
determined that NoVLP attachment in young leaves was primarily
associated with proteins whereas in older leaves it was primarily
through carbohydrates (14). This difference in the binding mechanisms
of NoVLPs to cell wall material could be the result of different sugar
abundances or differences in composition between young and old
leaves. This is supported by previous studies showing that the plant
cell wall changes continuously in different developmental stages (8).
NoV-lettuce interaction is mediated by various carbohydrate moieties
present in the plant cell walls (14). Minor binding to cell wall proteins
was also observed (14), which is supported by the observed binding of
NoV G1.1 binding to unknown proteinaceous components released from
the surface of lettuce plants (16).
Esseili et al. (14) investigated the specific inhibition of
carbohydrate moieties, using specific monoclonal antibodies against
carbohydrates of human HBGAs. The antibody against the A antigen
was the only antibody that significantly inhibited VLP binding to PGM,
and none of the MAbs inhibited binding of VLPs to lettuce leaves.
This further confirms the hypothesis of Gandhi et al. (16) that NoVLPs
bind to Romaine lettuce leaves differently from the binding to HBGAs
and PGM. In addition to assessing norovirus attachment to Romaine
lettuce; extracts prepared from cilantro, iceberg lettuce, celery,
spinach, green onions, clover sprouts, and raspberries were used to
coat ELISA wells to measure NoVLP binding; however, none of these
bound NoVLPs as well as Romaine lettuce (16). In a study assessing the
ability of washing to remove human noroviruses from different types
of produce, it was observed that a greater percentage of GI NoV was
removed from raspberries (95%) than from lettuce (75%), suggesting
stronger binding of NoV to lettuce (55). The amount of virus removed by
washing significantly decreased when produce was washed with acidic
electrolyzed water (AEW, pH 2.2–2.4), suggesting enhanced binding of
norovirus to raspberries and lettuce due to ionic interactions (55).
Using attachment to detect norovirus detection and to
assess infectivity
The attachment properties of noroviruses have been exploited
for use in detection assays (9, 17, 50, 53, 54) in which HBGAs or PGM
were conjugated to magnetic beads that were then used to isolate
noroviruses from water, sewage/wastewater and lettuce. Advantages
of isolating noroviruses through HBGA attachment methods include
both ability to detect intact capsids and increased sensitivity.
Recently, a study used viral binding properties in tandem with RT-PCR
to discriminate between infectious and non-infectious noroviruses
in a binding-based RT-PCR assay (26). Binding-based RT-PCR was
investigated for its ability to distinguish between infectious and noninfectious virus particles, with the hypothesis that binding to a receptor
followed by RT-PCR would better indicate viral infectivity, eliminating
the detection of viruses with damaged capsid or RNA. In this study,
attachment of both infectious and inactivated (by H2O2 or heat)
MNV and NoV was assessed using ganglioside GD1a, the attachment
receptor of MNV on RAW cells, and PGM or Caco-2 cells, respectively.
MNV results were confirmed by plaque assay. For the inactivation
of MNV by heat (70°C, 3 min) and H2O2 (2.1%, 5 min), the plaque
assay showed reductions of > 6 log PFU/mL. Reductions of 1 log were
obtained by RT-PCR after heat inactivation as well as by cell binding
RT-PCR.
The RT-PCR assay was shown to be a better indicator of infectivity
for heat-treated MNV and human norovirus GII.4 (70°C, 2 min)
compared with RNase treatment of virus followed by RT-PCR (28).
The RNase RT-PCR methods measure the ability of the virus capsid to
protect the interior viral RNA genome from digestion by RNase (56),
while the cell-binding RT-PCR assay is based on the ability of the virus
capsid to bind to specific receptors. This binding is essential for viral
infection (26), likely because the structure of the protruding (P) domain
on the capsid is denatured by heat before the occurrence of any severe
damage on the capsid that would enable the entry of RNase One and
subsequently RNA degradation (28). Using virus attachment to PGM
coated beads coupled with viral detection by qRT-PCR, Dancho et al.
(12) investigated whether human norovirus GI.1 and GII.4 binding
ability after treatment with heat, high-pressure, and UV was correlated
to infectivity. Results showed that these virus treatments affected
capsid integrity enough to interfere with attachment to PGM (12);
however, human clinical trials for UV and heat-treated noroviruses have
not been conducted, and it cannot be said with certainty that altered
binding properties definitely resulted in loss of norovirus infectivity.
Although none of these assays identifies a reduction in virus titer
comparable to results obtained with infectivity assays, they represent
an improvement in evaluation of norovirus infectivity compared
with results obtained with standard RT-PCR and they enhance our
understanding of the steps involved in virus infectivity.
Recently, we published research using murine norovirus (MNV)
as a surrogate for human norovirus, in which an ELISA method was
utilized to assess attachment through binding to host cell receptors,
and MNV attachment was correlated to infectivity as determined by a
plaque assay (18). ELISA plates were coated with porcine gastric mucin
and untreated, heat-, high pressure-, ozone- or UV-treated MNV was
added, followed by monoclonal anti-MNV IgG antibody. Through use of
this ELISA attachment assay on heat-treated Murine norovirus (MNV),
a positive correlation was observed between virus attachment patterns
and inactivation of virus. While this correlation was not observed for
other processing methods, including high pressure-, ozone- and UVtreated MNV, this assay has the potential to be useful for predicting
human norovirus infectivity after heat treatment. The objective of the
short study described here is to assess the applicability of the ELISA
attachment assay to a GII NoV and to assess whether food matrices
interfere with attachment of the virus to the carbohydrate coating on
the ELISA plate. In particular, attachment of MNV and NoV suspended
in spinach and green onions was assessed.
MATERIALS AND METHODS
Human norovirus
GII NoV were isolated from fecal samples (generously provided by
Megan Davis, South Carolina Department of Health and Environmental
Control). NoV was extracted from stool samples by a protocol similar
to Straub et al. (38). Stool samples were suspended in PBS (0.01 M)
to obtain a 10–20% stool suspension. The suspension was vortexed
for 60s, centrifuged at 1000 x g and processed through at 0.22
micron-filter to remove bacterial contamination. NoV was purified by
a polyethylene glycol (PEG) precipitation as described by Lewis and
Metcalf (25). Briefly, viral suspensions were added to PEG 8000 (Fisher
Scientific, Waltham, MA) to obtain a final concentration of 8% (wt/vol),
stirred for 2 h at 4°C, and centrifuged at 10,000 x g for 20 min. The PEG
supernatant was discarded and the viral pellet resuspended in 0.15
M Na2HPO4 (pH 9.0) and then shaken for 20 min at 250 rpm before
centrifugation at 10,000 x g for 30 min. The pH of the supernatant
was adjusted to 7.4 and stored at -80°C until used. NoV was heat
treated at 20, 50, 60, 70, 80 and 100°C for 5 min in 200 µL aliquots
of the supernatant in 0.2 mL PCR tubes in an Eppendorf Mastercycler
(Eppendorf, Duesseldorf, Germany).
Attachment – enzyme-linked immunosorbant assay (ELISA)
ELISA plates (CoStar, Corning, NY) were pre-coated with a
10% solution of poly-L-lysine (Sigma Aldrich, St. Louis, MO) in Tris
buffered saline (TBS, 100 mM, pH 7.4). After a 30-min incubation,
ELISA plates were washed three times with sterile ddH2O, and coated
with 100 μL of 10 μm/mL PGM (Sigma Aldrich, St. Louis, MO) in TBS
and incubated for 24–48 h at 4°C. Unbound PGM was removed by
washing three times with ddH2O and the wells were then blocked for
2 h at room temperature with Animal-Free Blocking Buffer (Vector
Labs, Burlingame, CA). After removal of the blocking buffer, wells were
SEPTEMBER-OCTOBER 2013 FOOD PROTECTION TRENDS 293
TABLE 1. RNase treatment of heat-treated GII human norovirus to determine capsid integrity followed by RT-PCR. Nucleic acid presence is expressed as the number of intact capsid samples/destroyed capsid (n=2)
20˚C
50˚C
60˚C
70˚C
80˚C
100˚C
2/0
2/0
2/0
2/0
0/2
0/2
4
3.5
Attachment (P/N Ratio)
3
2.5
2
1.5
1
0.5
0
20
50
60
70
80
100
Temperature (C˚)
FIGURE 1. Attachment of heat-treated GII human norovirus to porcine gastric mucin coated ELISA plates. Positive attachment is indicated by
a P/N ratio ≥ 2, shown by a bold line, confirming that treated virus can be recognized by monoclonal antibodies. P/N Ratios are not
statistically different (P > 0.05).
294 FOOD PROTECTION TRENDS SEPTEMBER-OCTOBER 2013
washed three times with ddH2O. MNV or GII Human norovirus samples
diluted in TBS (10-1 dilution) were added to wells and incubated at
37°C for 1 h. Unbound virus was removed and wells were washed three
times with ddH2O before the addition of 1:1000 dilution of monoclonal
mouse anti-MNV IgG (generously provided by Dr. Christine Wobus,
University of Michigan) or anti-NoV mouse IgG P4D8 monoclonal
antibody (Kim Laboratories, Rantoul, IL) diluted in TBS. Anti-MNV IgG
or anti-NoV IgG was allowed to incubate at 37°C for 1 h before unbound
anti-MNV IgG was removed by washing the wells with TTBS (TBS with
0.5% Tween-20). Anti-MNV or anti-NoV antibodies were detected by
horseradish peroxidase conjugated goat anti-mouse IgG secondary
antibody (Promega, Madison, WI) diluted 1:2500 in TTBS. The secondary
antibody was incubated at 37°C for 1 h. Unbound secondary antibody
was removed by washing three times with TTBS, and bound HRPconjugated goat anti-mouse antibodies were detected by adding 80 μL
of 3,3’,5,5’– tetramethylbenzidine (TMB) liquid substrate (Promega).
Samples were incubated at room temperature for 5 min before the
reaction was stopped by addition of 1N HCl, and absorbance was read
at 405 nm on a plate reader.
RNase treatment of heat-treated NoV GII
To determine whether viral capsids were intact after treatment,
heat-treated NoV GII was subjected to RNAse prior to RT-PCR, as
previously described (18). Briefly, NoV GII was subjected to RNase (20
units) for 30 min at 37°C, after which RNase inhibitor (20 units) was
added. Viral RNA was released by treatment at 99°C for 5 min. RNase
inhibitor was added again and One-step RT-PCR (Qiagen, Valencia, CA)
was performed on serially diluted virus in 25 µL volumes containing
5 µL 5✕ Buffer, 1 µL dNTP (10mM), 2 µL RNA, 1 µL enzyme mix, 0.2 µL
RNase Inhibitor (Fisher Scientific, Waltham, MA) and 9.8 µL RNasefree H2O. Primers for NoV GII were designed by Primer3 and target the
VP1 region (VP1-FP3 – 5’-TGGGTGCTCCCAAGTTATTC-3’ and VP1-RP35’-CTGGAGCTGCCTCTTGGTAG-3’) to produce a product of 196-bp.
Effect of spinach and green onions on viral attachment to PGM
To assess MNV and NoV attachment to spinach and green onion
plants, one mature plant was homogenized in TBS (10 mL), coated on
ELISA plates and incubated for 24–48 h at 4°C. The attachment ELISA
assay was performed as described previously (18). To assess the effect
of plant homogenate on results of the MNV and NoV attachment assay,
ELISA plates were pre-coated with poly-L-lysine and coated with PGM
and then incubated for 24–48 h at 4°C as described previously. MNV
and NoV suspended in spinach and green onions homogenized in TBS
were added to wells after blocking and washing. Attached MNV and NoV
were detected by anti-MNV and anti-NoV IgG and HRP-conjugated goat
anti-mouse secondary antibody as described previously.
Data analysis
Virus attachment is represented as the positive to negative ratio
(P/N), in which the OD405 readings of sample wells were compared
to those of the negative control wells. Negative control wells included
wells without PGM, spinach or green onion homogenate, or wells without
the primary antibody. A P/N Ratio ≥ 2 was considered to be positive
attachment (18, 48). Three technical replicates were performed for
three biological replicates and the results recorded as the mean and
standard deviation of the mean. Statistical analysis included the
difference of means t-test on Microsoft Excel 2010. Attachment was
considered significant if P ≤ 0.05.
RESULTS AND DISCUSSION
Attachment of heat-treated human norovirus GII
Human norovirus genogroup II attached positively (P/N Ratio > 2)
to porcine gastric mucin after heat treatment (Fig. 1), P/N ratios were
2.70 ± 0.83, 2.30 ± 0.39, 2.26 ± 0.77, 2.01 ± 0.75, 2.09 ± 0.44, and
2.04 ± 0.29 for 20, 50, 60, 70, 80, and 100°C, respectively. However,
attachment P/N ratios were close to the cut-off value of 2 at 70, 80 and
100°C. Results with RNase treatment (Table 1) indicate the NoV capsid
does not remain intact at 80 and 100°C; norovirus RNA is not detected
at these temperatures, since the viral capsid is no longer intact,
which allows RNase degradation of the viral genome. This pattern
of attachment with heat-treated human norovirus is similar to that
observed for heat-treated MNV (18), which may indicate that NoV is no
longer infectious when the P/N ratio of attachment to porcine gastric
mucin is less than or equal to 2. However, this is difficult to predict, as
to date human clinical trials assessing NoV infectivity have not been
performed on heat-treated NoVs.
Effect of spinach and green onion homogenate on MNV and NoV
attachment to porcine gastric mucin
The effects of food matrices on results of the attachment assay
were assessed using spinach and green onion plants for MNV (Fig. 2)
and NoV (Fig. 3). MNV attached strongly to spinach and green onion
when the plant homogenate was coated on the ELISA plates, with
P/N values of 13.60 ± 0.24 and 14.50 ± 0.33, respectively. Despite
the significantly greater (P ≤ 0.05) attachment of MNV to both plant
homogenates, MNV suspended in spinach and green onion homogenate
was able to positively attach to porcine gastric mucin, where P/N ratios
were 2.75 ± 0.13 and 3.05 ± 0.29 (Fig. 2). Similarly, NoV was shown to
positively attach to spinach and green onions when plant homogenate
was coated to ELISA plates, with P/N values of 3.82 ± 0.27 and 3.73
± 0.46, respectively (Fig. 3). While there is a positive attachment
(P/N ≥ 2), this interaction of NoV with the spinach and green onion
homogenates, as indicated by the P/N Ratio, is signifcantly less
(P ≤ 0.05) than it was for MNV, possibly because of the difference in
the binding/attachment properties and structure of the two viruses
(22, 23, 43, 47). When NoV was suspended in spinach and green onion
homogenate, NoV positively attached to PGM, with P/N ratios of 3.64 ±
0.76 and 2.56 ± 0.52, respectively.
These results indicate that, despite the attachment of these
viruses to produce, this attachment does not interfere with attachment
to PGM, suggesting that this assay could be used for assessing
infectivity of heat-treated virus or of virus in some food commodities.
It is important to assess any inhibition by compounds in plant material
that could interfere with the assay. Attachment of MNV and NoV to
extracts of spinach was demonstrated by Gandhi et al. (16), who
showed that recombinant human norovirus-like particles (rNoVLPs)
attached positively to spinach; however, the attachment of rNoVLPs to
spinach was weaker than the attachment of rNoVLPs to Romaine lettuce
SEPTEMBER-OCTOBER 2013 FOOD PROTECTION TRENDS 295
16
Attachment (P/N Ratio)
14
12
10
8
6
4
2
0
spinach
green onion
FIGURE 2. Attachment of murine norovirus to spinach or green onion homogenate ( □ ) or attachment of murine norovirus suspended in spinach or green onion homogenate to porcine gastric mucin (■ ). Positive attachment is indicated by a P/N
ratio ≥ 2, shown by a bold line, confirming that murine norovirus virus can be recognized by monoclonal antibodies.
Attachment (P/N Ratio)
5
4
3
2
1
0
spinach
green onion
FIGURE 3. Attachment of GII human norovirus to spinach or green onion homogenate (□ ) or attachment of NoV suspended in
spinach or green onion homogenate to porcine gastric mucin (■ ). Positive attachment is indicated by a P/N ratio ≥ 2,
shown by a bold line, confirming that GII human norovirus can be recognized by monoclonal antibodies.
296 FOOD PROTECTION TRENDS SEPTEMBER-OCTOBER 2013
extracts. Additionally, Gandhi et al.(16) did not observe attachment
of rNoVLPs to green onions. Food items such as oysters and Romaine
lettuce, which have been shown to bind to NoVs through carbohydrates
similarly to HBGA attachment of NoVs (14, 24, 49), may affect
attachment to PGM and pose a challenge to predicting infectivity using
this attachment ELISA. Li et al. (27) tested the binding ability
of GII.4 NoV P particles on various types of shellfish, vegetables, and
fruit juices in a saliva-binding enzyme-linked immunosorbant assay
(ELISA). As predicted, oysters had negative effects on NoV P particle
attachment to human saliva, suggesting that the shellfish extracts
blocked the binding sites of the NoV P particles, thus reducing the
HBGA-binding levels. No inhibition of binding effects were observed
for the fresh produce, including strawberry, raspberry, blackberry,
blueberry, cherry tomatoes, spinach, and romaine lettuce (27). It should
be taken into consideration that the behavior of NoV P particles may
not be indicative of human norovirus particles. The components of food
such as carbohydrates will vary between commodities and should be
assessed for each intended use.
CONCLUSION
The studies described here represent a broad view of the
challenges to understanding how NoV interacts with foods. This
knowledge can help lead to the development of more effective barriers
against initial contamination and limit spread or transmission of NoV
among food products. Attachment of NoV is complex, as shown by
the various carbohydrates and proteins that may be involved. These
challenges are greatly increased by the inability to culture human NoV
and the reliance on surrogates, which, although they may provide a
better model for inactivation, are weaker models for attachment.
ACKNOWLEDGMENTS
This project was funded in part by the United States Department of
Agriculture NoroCORE grant no. 1111-2011-0494.
REFERENCES
1. Baker, K., J. Morris, N. McCarthy, L. Saldana, J. Lowther, A. Collinson, and M. Young. 2011. An outbreak of norovirus infection linked to oyster
consumption at a UK restaurant, February 2010. J. Public Health 33:205–211.
2. Berger, C. N., S. V. Sodha, R. K. Shaw, P. M. Griffin, D. Pink, P. Hand, and G. Frankel. 2010. Fresh fruit and vegetables as vehicles for the
transmission of human pathogens. Environ. Microbiol. 12:2385–2397.
3. Bertolotti-Ciarlet, A., L. J. White, R. Chen, B. V. V. Prasad, and M. K. Estes. 2002. Structural requirements for the assembly of Norwalk virus-like
particles. J. Virol. 76:4044–4055.
4. Bhella, D., D. Gatherer, Y. Chaudhry, R. Pink, and I. G. Goodfellow. 2008. Structural insights into calicivirus attachment and uncoating. J. Virol.
82:8051–8058.
5. Bomsel, M., and A. Alfsen. 2003. Entry of viruses through the epithelial barrier: Pathogenic trickery. Nat. Rev. Mol. Cell Bio. 4:57–68.
6. Bu, W. M., A. Mamedova, M. Tan, M. Xia, X. Jiang, and R. S. Hegde. 2008. Structural basis for the receptor binding specificity of norwalk virus.
J. Virol. 82:5340–5347.
7. Buenaventura, E., R. Szabo, K. Mattison, O. Mykuczuk, A. Youssef, M. C. Lavoie, A. Todd, and R. Edwards. 2011. A norovirus outbreak in Vancouver,
British Columbia related to the consumption of raw oysters from a specific harvest site implicating an ill worker. J. Shellfish Res. 30:489–489.
8. Caffall, K.H., and D. Mohnen. 2009. The structure, function, and biosynthesis of plant cell wall pectic polysaccharides. Carbohyd. Res. 344:1879–1900.
9. Cannon, J. L., and J. Vinje. 2008. Histo-blood group antigen assay for detecting noroviruses in water. Appl. Environ. Microbiol. 74:6818–6819.
10. Cao, S., Z. Y. Lou, M. Tan, Y. T. Chen, Y. J. Liu, Z. S. Zhang, X. J. C. Zhang, X. Jiang, X. M. Li, and Z. H. Rao. 2007. Structural basis for the
recognition of blood group trisaccharides by norovirus. J. Virol. 81:5949–5957.
11. Choi, J. M., A. M. Hutson, M. K. Estes, and B. V. V. Prasad. 2008. Atomic resolution structural characterization of recognition of histo-blood group
antigens by Norwalk virus. P. Natl. Acad. Sci. USA 105:9175–9180.
12. Dancho, B. A., H. Chen and D. H. Kingsley. 2012. Discrimination between infectious and non-infectious human norovirus using porcine gastric
mucin. Int. J. Food Microbiol. 155:222–226.
13. Digirolamo, R., J. Liston, and J. Matches. 1977. Ionic bonding, mechanism of viral uptake by shellfish mucu. Appl. Environ. Microbiol. 33:19–25.
14. Esseili, M. A., Q. H. Wang, and L. J. Saif. 2012. Binding of human GII.4 norovirus virus-like particles to carbohydrates of Romaine lettuce leaf
cell wall materials. Appl. Environ. Microbiol. 78:786–794.
15. Ethelberg, S., M. Lisby, B. Bottiger, A. C. Schultz, A. Villif, T. Jensen, K. E. Olsen, F. Scheutz, C. Kjelso, and L. Muller. 2010. Outbreaks of
gastroenteritis linked to lettuce, Denmark, January 2010. Eurosurveillance 15:2–4.
16. Gandhi, K. M., R. E. Mandrell, P. Tian. 2010. Binding of virus-like particles of Norwalk virus to Romaine lettuce veins. Appl. Environ. Microbiol.
76:7997–8003.
17. Harrington, P. R., J. Vinje, C. L. Moe, and R. S. Baric. 2004. Norovirus capture with histo-blood group antigens reveals novel virus-ligand
interactions. J. Virol. 78:3035–3045.
18. Hirneisen, K. A., and K. E. Kniel. 2012. Comparison of ELISA attachment and infectivity assays for murine norovirus. J. Virol. Met. 186:14–20.
19. Huang, P. W., T. Farkas, W. M. Zhong, S. Thornton, A. L. Morrow, and J. Xi. 2005. Norovirus and histo-blood group antigens: Demonstration of a
wide spectrum of strain specificities and classification of two major binding groups among multiple binding patterns. J. Virol. 79:6714–6722.
SEPTEMBER-OCTOBER 2013 FOOD PROTECTION TRENDS 297
20. Hutson, A. M., R. L. Atmar, D. Y. Graham, and M. K. Estes. 2002. Norwalk virus infection and disease is associated with ABO histo-blood group
type. J. Infect. Dis. 185:1335–1337.
21. Hutson, A. M., R. L. Atmar, D. M. Marcus, and M. K. Estes. 2003. Norwalk virus-like particle hemagglutination by binding to H histo-blood group
antigens. J. Virol. 77:405–415.
22. Katpally, U., N. R. Voss, T. Cavazza, S. Taube, J. R. Rubin, V. L. Young, J. Stuckey, V. K. Ward, H. W. Virgin, C. E. Wobus, and T. J. Smith. 2010.
High-resolution cryo-electron microscopy structures of murine norovirus 1 and rabbit hemorrhagic disease virus reveal marked flexibility in the
receptor binding domains. J. Virol. 84:5836–5841.
23. Katpally, U., C. E. Wobus, K. Dryden, H. W. Virgin, and T. J. Smith. 2008. Structure of antibody-neutralized murine norovirus and unexpected
differences from viruslike particles. J. Virol. 82:2079–2088.
24. Le Guyader, F. S., F. Loisy, R. L. Atmar, A. M. Hutson, M. K. Estes, N. Ruvoen-Clouet, M. Pommepuy, and J. Le Pendu. 2006. Norwalk virus-specific
binding to oyster digestive tissues. Emerg. Infect. Dis. 12:931–936.
25. Lewis, G. D., and T. G. Metcalf. 1988. Polyethylene-glycol precipitation for recovery of pathogenic viruses, including hepatitis-a virus and human
rotavirus from oyster, water and sediment samples. Appl. Environ. Microbiol. 54:1983–1988.
26. Li, D., L. Baert, E. Van Coillie, and M. Uyttendaele. 2011. Critical studies on binding-based RT-PCR detection of infectious Noroviruses. J. Virol. Met. 177:153–159.
27. Li, D., L. Baert, M. Xia, W. Zhong, X. Jiang, and M. Uyttendaele. 2012a. Effects of a variety of food extracts and juices on the specific binding
ability of norovirus GII.4 P particles. J. Food Prot. 75:1350–1354.
28. Li, D., L. Baert, M. Xia, W. Zhong, E. Van Coillie, X. Jiang, and M. Uyttendaele. 2012b. Evaluation of methods measuring the capsid integrity
and/or functions of noroviruses by heat inactivation. J. Virol. Met. 181:1–5.
29. Maalouf, H., M. Pommepuy, and F. S. Le Guyader. 2010a. Environmental conditions leading to shellfish contamination and related outbreaks.
Food Environ. Virol. 2:136–145.
30. Maalouf, H., M. Zakhour, J. Le Pendu, J. C. Le Saux, R. L. Atmar, and F. S. Le Guyader. 2010b. Distribution in tissue and seasonal variation of
norovirus genogroup I and II ligands in oysters. Appl. Environ. Microbiol. 76:5621–5630.
31. Makino, A., M. Shimojima, T. Miyazawa, K. Kato, Y. Tohya, and H. Akashi. 2006. Junctional adhesion molecule 1 is a functional receptor for feline
calicivirus. J. Virol. 80:4482–4490.
32. Marionneau, S., N. Ruvoen, B. Le Moullac-Vaidye, M. Clement, A. Cailleau-Thomas, G. Ruiz-Palacois, P. W. Huang, X. Jiang, and J. Le Pendu. 2002.
Norwalk virus binds to histo-blood group antigens present on gastroduodenal epithelial cells of secretor individuals. Gastroenterology. 122:1967–1977.
33. Mesquita, J. R., and M. S. J. Nascimento. 2009. A foodborne outbreak of norovirus gastroenteritis associated with a Christmas dinner in Porto,
Portugal, December 2008. Eurosurveillance. 14:19–21.
34. Ossiboff, R. J., and J. S. L. Parker. 2007. Identification of regions and residues in feline junctional adhesion molecule required for feline
calicivirus binding and infection. J. Virol. 81:13608–13621.
35. Prasad, B. V. V., M. E. Hardy, T. Dokland, J. Bella, M. G. Rossmann, and M. K. Estes. 1999. X-ray crystallographic structure of the Norwalk virus
capsid. Science 286:287–290.
36. Provost, K., B. A. Dancho, G. Ozbay, R. S. Anderson, G. P. Richards, and D. H. Kingsley. 2011. Hemocytes are sites of enteric virus persistence
within oysters. Appl. Environ. Microbiol. 77:8360–8369.
37. Scallan, E., R. M. Hoekstra, F. J. Angulo, R. V. Tauxe, M. A. Widdowson, S. L. Roy, J. L. Jones, and P. M. Griffin. 2011. Foodborne illness acquired in
the United States-Major pathogens. Emerg. Infect. Dis. 17:7–15.
38. Straub, T. M., K. H. Z. Bentrup, P. Orosz-Coghlan, A. Dohnalkova, B. K. Mayer, R. A. Bartholomew, C. O. Valdez, C. J. Bruckner-Lea, C. P. Gerba,
M. Abbaszadegan, and C. A. Nickerson. 2007. In vitro cell culture infectivity assay for human noroviruses. Emerg. Infect. Dis. 13:396–403.
39. Stuart, A. D., and T. D. K. Brown. 2007. Alpha 2,6-linked sialic acid acts as a receptor for Feline calicivirus. J. Gen. Virol. 88:177–186.
40. Tamura, M., K. Natori, M. Kobayashi, T. Miyamura, and N. Takeda. 2000. Interaction of recombinant Norwalk virus particles with the
105-kilodalton cellular binding protein, a candidate receptor molecule for virus attachment. J. Virol. 74:11589–11597.
41. Tamura, M., K. Natori, M. Kobayashi, T. Miyamura, and N. Takeda. 2004. Genogroup II noroviruses efficiently bind to heparan sulfate proteoglycan
associated with the cellular membrane. J. Virol. 78:3817–3826.
42. Tan, M., P. W. Huang, J. Meller, W. M. Zhong, T. Farkas, and X. Jiang. 2003. Mutations within the P2 domain of norovirus capsid affect binding to
human histo-blood group antigens: Evidence for a binding pocket. J. Virol. 77:12562–12571.
43. Tan, M., and X. Jiang. 2010. Norovirus gastroenteritis, carbohydrate receptors, and animal models. Plos. Pathog. 6:e1000983.
44. Tan, M., and X. Jiang. 2011. Norovirus-host interaction: Multi-selections by human histo-blood group antigens. Trends Microbiol. 19:382–388.
45. Tan, M., M. Xia, S. Cao, P. W. Huang, T. Farkas, J. Meller, R. S. Hegde, X. M. Li, Z. H. Rao, and X. Jiang. 2008. Elucidation of strain-specific
interaction of a GII-4 norovirus with HBGA receptors by site-directed mutagenesis study. Virol. 379:324–334.
46. Tan, M., M. Xia, Y. T. Chen, W. M. Bu, R. S. Hegde, J. Meller, X. M. Li, and X. Jiang. 2009. Conservation of carbohydrate binding interfaces —
Evidence of human HBGA selection in norovirus evolution. Plos. One 4:e5058.
47. Taube, S., J. R. Rubin, U. Katpally, T. J. Smith, A. Kendall, J. A. Stuckety, and C. E. Wobus. 2010. High-resolution x-ray structure and functional
analysis of the murine norovirus 1 capsid protein protruding domain. J. Virol. 84:5695–5705.
298 FOOD PROTECTION TRENDS SEPTEMBER-OCTOBER 2013
48. Tian, P., A. H. Bates, H. M. Jensen, and R. E. Mandrell. 2006. Norovirus binds to blood group A-like antigens in oyster gastrointestinal cells.
Lett. Appl. Microbiol. 43:645–651.
49. Tian, P., M. Brandl, and R. Mandrell. 2005. Porcine gastric mucin binds to recombinant norovirus particles and competitively inhibits their
binding to histo-blood group antigens and Caco-2 cells. Lett. Appl. Microbiol. 41:315–320.
50. Tian, P., A. Engelbrektson, and R. Mandrell, R. 2008a. Two-log increase in sensitivity for detection of norovirus in complex samples by
concentration with porcine gastric mucin conjugated to magnetic beads. Appl. Environ. Microbiol. 74:4271–4276.
51. Tian, P., A. L. Engelbrektson, X. Jiang, W. M. Zhong, and R. E. Mandrelli. 2007a. Norovirus recognizes histo-blood group antigens on
gastrointestinal cells of clams, mussels, and oysters: A possible mechanism of bioaccumulation. J. Food Prot. 70:2140–2147.
52. Tian, P., A. L. Engelbrektson, and R. E. Mandrell. 2008b. Seasonal tracking of histo-blood group antigen expression and norovirus binding
in oyster gastrointestinal cells. J. Food Prot. 71:1696–1700.
53. Tian, P., D. Yang, X. Jiang, W. Zhong, J. L. Cannon, W. Burkhardt, J. W. Woods, G. Hartman, L. Lindesmith, R. S. Baric, and R. Mandrell. 2010.
Specificity and kinetics of norovirus binding to magnetic bead-conjugated histo-blood group antigens. J. Appl. Microbiol. 109:1753–1762.
54. Tian, P., D. Yang, and R. Mandrell. 2011a. A simple method to recover Norovirus from fresh produce with large sample size by using histo-blood
group antigen-conjugated to magnetic beads in a recirculating affinity magnetic separation system (RCAMS). Int. J. Food Microbiol. 147:223–227.
55. Tian, P., D. Yang, and R. Mandrell. 2011b. Differences in the binding of human norovirus to and from Romaine lettuce and raspberries by water
and electrolyzed waters. J. Food Prot. 74:1364–1369.
56. Topping, J. R., H. Schnerr, J. Haines, M. Scott, M. J. Carter, M. M. Willcock, K. Bellamy, D. W. Brown, J. J. Gray, C. I. Gallimore, and A. I. Knight.
2009. Temperature inactivation of Feline calicivirus vaccine strain FCV F-9 in comparison with human noroviruses using an RNA exposure
assay and reverse transcribed quantitative real-time polymerase chain reaction-A novel method for predicting virus infectivity. J. Virol. Met.
156:89–95.
57. Truyols, A. G., J. G. Duran, A. N. Riutort, G. A. Cerda, C. B. Isabel, M. P. Arbona, and J. V. Berga. 2011. Norovirus outbreak in Majorca (Spain)
associated with oyster consumption. Gac. Sanit. 25:173–175.
58. Vega, E., J. Garland, and S.D. Pillai. 2008. Electrostatic forces control nonspecific virus attachment to lettuce. J. Food Prot. 71:522–529.
59. Vega, E., J. Smith, J. Garland, A. Matos, and S. D. Pillai. 2005. Variability of virus attachment patterns to butterhead lettuce. J. Food Prot.
68:2112–2117.
60. Wang, Q., Z. Zhang, and L. J. Saif. 2012. Stability of and attachment to lettuce by a culturable porcine sapovirus surrogate for human
Calicivirus. Appl. Environ. Microbiol. 78:3932–3940.
61. Wei, J., Y. Jin, T. Sims, and K. E. Kniel. 2010. Manure- and biosolids-resident murine norovirus 1 attachment to and internalization by Romaine
lettuce. Appl. Environ. Microbiol. 76:578–583.
62. Woods, J. W., and W. Burkhardt. 2010. Occurrence of norovirus and hepatitis A virus in U.S. oysters. Food Environ. Virol. 2:176–182.
SEPTEMBER-OCTOBER 2013 FOOD PROTECTION TRENDS 299