Download A release-competent influenza A virus mutant lacking the coding

Journal
of General Virology (2002), 83, 2683–2692. Printed in Great Britain
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A release-competent influenza A virus mutant lacking the
coding capacity for the neuraminidase active site
Larisa V. Gubareva,1, 2 Marina S. Nedyalkova,1, 2 Dmitri V. Novikov,1 K. Gopal Murti,3
Erich Hoffmann3† and Frederick G. Hayden1
1
Department of Internal Medicine, University of Virginia, 1300 Jefferson Park Avenue, Jordan Hall Room 2231, PO Box 800473,
Charlottesville, VA 22908, USA
2
DI Ivanovsky Institute of Virology, 16 Gamaleya Str., Moscow 123098, Russia
3
Department of Virology and Molecular Biology, St Jude Children’s Research Hospital, 332 North Lauderdale Str., Memphis,
TN 38108, USA
Both influenza A virus surface glycoproteins, the haemagglutinin (HA) and neuraminidase (NA),
interact with neuraminic acid-containing receptors. The influenza virus A/Charlottesville/31/95
(H1N1) has shown a substantially reduced sensitivity to NA inhibitor compared with the A/WSN/33
(H1N1) isolate by plaque-reduction assays in Madin–Darby canine kidney (MDCK) cells. However,
there was no difference in drug sensitivity in an NA inhibition assay. The replacement of the HA
gene of A/WSN/33 with the HA gene of A/Charlottesville/31/95 led to a drastic reduction in
sensitivity of A/WSN/33 to NA inhibitor in MDCK cells. Passage of A/Charlottesville/31/95 in cell
culture in the presence of an NA inhibitor resulted in the emergence of mutant viruses (delNA)
whose genomes lacked the coding capacity for the NA active site. The delNA mutants were plaqueto-plaque purified and further characterized. The delNA-31 mutant produced appreciable yields
(" 106 p.f.u./ml) in MDCK cell culture supernatants in the absence of viral or bacterial NA activity.
Sequence analysis of the delNA mutant genome revealed no compensatory substitutions in the HA
or other genes compared with the wild-type. Our data indicate that sialylation of the oligosaccharide chains in the vicinity of the HA receptor-binding site of A/Charlottesville/31/95 virus
reduces the HA binding efficiency and thus serves as a compensatory mechanism for the loss of NA
activity. Hyperglycosylation of HA is common in influenza A viruses circulating in humans and has
the potential to reduce virus sensitivity to NA inhibitors.
Introduction
Both influenza A virus surface glycoproteins, the haemagglutinin (HA) and neuraminidase (NA), interact with neuraminic acid-containing receptors. The HA binds to neuraminic
Author for correspondence : Larisa Gubareva (at University of Virginia).
Fax j1 434 924 9065. e-mail LVG9B!virginia.edu
†Present address : MedImmune, Inc., 297 North Bernando Avenue,
Mountain View, CA 94043, USA.
The GenBank accession numbers of the sequences reported in this
paper are A/Charlottesville/31/95 : PB1 AF398865, PB2 AF398866,
PA AF398862, HA AF398878, NP AF398867, defective NA
AF398870, M AF398876, NS AF398877 ; A/Charlottesville/28/95 :
PA AF398864, HA AF398874, defective NA AF398873.
0001-8453 # 2002 SGM
acid-containing receptors and the NA destroys HA receptors
by cleaving the terminal neuraminic acid residues from adjacent
oligosaccharide chains. Both viral glycoproteins are sialylated
by cellular enzymes, and the activity of viral NA is required to
prevent self-aggregation of progeny virions and to promote
release from the cellular membrane (Colman, 1994). Each
subunit of the NA homotetramer consists of a cytoplasmic tail,
a transmembrane domain, a stalk and a head region. The crystal
structure of the NA head, which contains the enzyme active
site, has been extensively studied and used in rational drug
design (Colman, 1994). The recently developed NA inhibitors
zanamivir and oseltamivir have demonstrated therapeutic
benefit in clinical trails, and the inhibitor RWJ-270201 (BCX1812) is undergoing clinical evaluation (Gubareva et al., 2000).
Influenza viruses have demonstrated a broad range of
sensitivities to NA inhibitors in cell culture, despite the efficient
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L. V. Gubareva and others
inhibition of the NA activities (Woods et al., 1993 ; Zambon &
Hayden, 2001). Prolonged passage in MDCK cells in the
presence of NA inhibitor can lead to the emergence of drugresistant viruses that have acquired amino acid substitutions in
the NA active site (McKimm-Breschkin, 2000). Additionally,
amino acid changes in the HA are frequently detected in NA
inhibitor-resistant mutants selected in vitro and often precede
drug-selected changes in the NA (McKimm-Breschkin, 2000).
Influenza viruses with substitutions in the NA active site and
reduced sensitivity in enzyme inhibition assays have occasionally been recovered from people treated with NA
inhibitors (Gubareva et al., 1998, 2001a ; Treanor et al.,
2000 ; Whitley et al., 2001). In accordance with the observed
interdependence of the HA and NA functions during influenza
virus propagation (Yang et al., 1997 ; Kaverin et al.,
1998 ; Mitnaul et al., 2000 ; Wagner et al., 2000 ; Baigent &
McCauley, 2001), the decreased efficiency of HA binding to
the cellular receptors is believed to facilitate the release and
spread of the virus when its NA activity is low. To this end,
previous studies by others have shown that NA-lacking
mutants, generated by supplying the bacterial NA and
antibodies to viral NA, undergo multiple rounds of replication
only in the presence of exogenous NA or after sonication of
membrane-attached virus aggregates (Liu & Air, 1993 ; Liu et
al., 1995). It has also been shown that substitutions in the HA
of the NA-lacking mutants can provide compensation for the
loss of the endogenous NA activity (Hughes et al., 2000).
Previously, we have reported that clinical isolates of
influenza A (H1N1) viruses A\Charlottesville\31\95 and
A\Charlottesville\28\95 passaged in MDCK cells in the
presence of NA inhibitors BCX-1812 or oseltamivir carboxylate acquired no point mutations in either the HA or NA genes.
Yet, with the use of RT–PCR analysis, we were able to detect
the emergence and accumulation of the defective RNA
segments encoding the NA but not HA in these viruses
(Nedyalkova et al., 2002). We suggested that the accumulation
of defective NA genes was a consequence of the reduced
requirement of the viruses for the NA activity, although the
mechanism of such reduction has not been explained.
In the present study, we demonstrate the key role
of the HA in the virus requirements for NA activity and in
the sensitivity to NA inhibitors by substituting the HA gene
of the A\WSN\33 virus with that of the A\Charlottesville\
31\95 virus. Our findings indicate that sialylation of oligosaccharide chains in the vicinity of the receptor-binding site
(RBS) of the HA is likely to provide a compensatory mechanism
for the NA activity deficit and allows emergence of NA-lacking
mutants.
Methods
Compounds. The influenza virus NA inhibitor BCX-1812 (RWJ270201) was provided by BioCryst.
Viruses and cells. A\WSN\33 (H1N1) was generated from eight
plasmids containing the entire virus genome (see below). Influenza A
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virus isolates A\Charlottesville\31\95 (H1N1) and A\Charlottesville\
28\95 (H1N1) were from the repository at the University of Virginia.
Madin–Darby canine kidney (MDCK) cells were cultured in minimal
essential medium (MEM) supplemented with 10 % foetal bovine serum.
Viruses were grown in monolayers of MDCK cells in MEM supplemented
with 0n3 % BSA and 1 µg\ml TPCK-treated trypsin. A\Charlottesville\31\95 underwent 18 passages in the presence of BCX-1812. The
clinical isolate A\Charlottesville\28\95 (H1N1) underwent five passages
in MDCK cells in the presence of oseltamivir carboxylate (Nedyalkova et
al., 2002).
All comparisons of properties were made between the properties of
the mutant and its wild-type (wt) virus grown in MDCK cells for the
same number of passages in the absence of the NA inhibitor.
Gene reassortment using reverse genetics. The eight plasmids
containing the cDNA of the virus A\WSN\33 (H1N1) were kindly
provided by Dr Robert G. Webster at St Jude Children’s Research
Hospital (Hoffmann et al., 2000). Transfection of co-cultivated 293T and
MDCK cells was performed as described (Hoffmann et al., 2000). The
reassortant virus was produced by transfection of cells with seven
plasmids containing the genome segments of A\WSN\33 and one
plasmid containing the HA gene of A\Charlottesville\31\95. The
plasmid pHW-HA31 containing the cDNA of the HA gene of
A\Charlottesville\31\95 was constructed by RT–PCR amplification of
the viral RNA. cDNA was amplified by PCR using oligonucleotide
primers containing the site of digestion for BsmBI. To amplify the
HA gene, the primers RGHA1 (5h TATTCGTCTCAGGGAGCAAAAGCAGGGGAAAAT 3h) and RGNS2\HA2 (5h ATATCGTCTCGTATTAGTAGAAACAAGGGTGTTT 3h) were used. Viral
sequences are underlined and restriction sites are shown in bold.
Virus concentration. The viruses were propagated in MDCK cells
by a standard procedure, and cell culture supernatants were clarified by
centrifugation at 1000 g for 20 min. The virus particles were pelleted by
centrifugation at 27 000 g in a Beckman SW 28 ultracentrifuge rotor
through a 25 % sucrose cushion for 2 h at 4 mC and resuspended in a
saline solution supplemented with 4 mM CaCl .
#
NA activity measurement. A standard fluorimetric assay with
minor modifications (Potier et al., 1979) was used to measure NA activity
of the virus in cellular supernatants and in concentrated virus preparations. NA activity was measured with 2h-(4-methylumbelliferyl) α--Nacetyl neuraminic acid (Sigma) as a substrate at a final concentration of
100 µM. The reactions were carried out in 33 mM MES buffer, pH 6n5,
supplemented with 4 mM CaCl . The assessment of the virus sensitivity
#
to NA inhibitors was performed in an NA inhibition assay, as described
previously (Gubareva et al., 2001b). The IC value reflects the
&!
concentration of drug that reduces the NA activity by 50 %.
Plaque-reduction assay. Virus sensitivity to NA inhibitors was
tested in a standard plaque-reduction assay, as described previously
(Gubareva et al., 2001a). Briefly, monolayers of MDCK cells were infected
with virus at an m.o.i. of approximately 30 p.f.u. per well. After 1 h
adsorption, the virus inoculum was discarded and cells were overlaid with
an MEM–agarose mixture containing trypsin and the NA inhibitor. The
diameter of plaques and their number were determined after 72 h of
incubation at 36 mC. Drug sensitivity was assessed by two criteria : the
concentration of inhibitor causing 50 % reduction of the plaque diameter
(IC plaque diameter) and the concentration causing reduction of the plaque
&!
number by 50 % (IC plaque number) compared with the control without
&!
inhibitor.
Immunofluorescent staining. Efficiency of virus adsorption was
examined as described previously for NA inhibitor-resistant viruses (Blick
et al., 1998) with minor modifications. MDCK cells were seeded on tissue
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Drug-selected NA deletion mutant
culture glass slides (Becton Dickinson). Confluent monolayers were
washed with cold PBS and inoculated with virus. Adsorption was allowed
to proceed for 20, 40 and 60 min on ice, followed by removal of the
inoculum and washing the cells. As a positive control for maximum
adsorption, the virus inoculum was removed after incubation for 60 min
on ice followed by additional incubation at 37 mC for 60 min. The
inoculum was removed and infected cells were incubated in a CO
#
incubator at 37 mC for 7 h followed by washing with PBS. The washed
cells were air-dried and fixed with cold acetone for 10 min. The cells were
stained with a mixture of anti-NP mAbs (kindly provided by Dr Robert
G. Webster, St Jude Children’s Research Hospital, Memphis, TN, USA),
followed by the addition of FITC-labelled goat anti-mouse immunoglobulins. The stained cells were viewed with a fluorescence microscope
(Olympus BH-2 RFCA).
Viral protein analysis. Radioisotopic labelling of viral proteins
was performed during virus propagation in MDCK cells. Four hours after
infection, cells were starved in methionine-free medium for 30 min. The
infected cultures were then incubated in maintenance medium containing
half the usual concentration of unlabelled methionine and [$&S]methionine
(25 µCi\ml) for 24 h. The growth medium did not contain trypsin to
prevent proteolytic cleavage of the HA precursor (HA ) into the two
!
subunits, HA and HA . The culture fluids were collected and clarified and
"
#
the labelled virus was pelleted through a 25 % sucrose cushion. Viral
proteins in the pellets were analysed by SDS–PAGE under reducing and
non-reducing conditions.
When specified, an aliquot of the concentrated virus was pretreated
with bacterial sialidase (5 units\ml, from Clostridium perfringens ; Glyko)
for 2 h at 37 mC or with peptide-N-glycosidase F (N-glycanase,
0n5 units\ml ; Glyko) overnight.
Electron microscopy. For electron microscopy studies, confluent
monolayers of infected cells were washed three times with PBS, fixed
with 2 % glutaradehyde in PBS for 2n5 h at 4 mC and post-fixed with 2 %
osmium tetroxide. The fixed cells were dehydrated with increasing
concentrations of ethanol (50–100 %) and embedded in a mixture of
epoxy resin. Ultrathin sections of cells were cut with a diamond knife on
a Sorvall MT 6000 Ultramicrotome. The sections were stained with
uranyl acetate and lead citrate and examined using a Philips EM301
electron microscope at 80 kV.
RT–PCR and sequence analysis. Extraction of viral RNA was
performed as described previously (Gubareva et al., 2001a). The sequences
of the primers used for PCR amplification of the eight segments of the
influenza A (H1N1) viruses and for their sequence analysis are available
on request.
The synthetic primer 5h AGCAAAAGCAGG 3h was used to generate
cDNA with reverse transcriptase (Promega). cDNA was amplified by a
standard PCR method and purified with the QIAquick PCR purification
kit (Qiagen). Purified PCR products were sequenced using Taq Dye
Terminator chemistry, according to the manufacturer’s instructions
(Applied Biosystems) and then analysed on an ABI 373 DNA sequencer
(Applied Biosystems) at the Center of Biotechnology at the University of
Virginia. Sequencher 4.0 software (Gene Codes Corporation) was used
for the analysis of nucleotide sequence data. The analysis did not include
the sequence of the 30 nucleotides at the 3h and 5h end of each segment.
Results
Sensitivity to NA inhibitors
To investigate the mechanisms that allow compensation for
the loss of the NA activity, we initially compared the drug
Fig. 1. Electron micrographs of influenza virus budding at the surface of
MDCK cells 12 h post-infection. Normal budding and release of
A/WSN/33 (A) and A/Charlottesville/31/95 (C) virions occurred in the
absence of the NA inhibitor. In the presence of 100 µg/ml of the NA
inhibitor BCX-1812, A/WSN/33 progeny virions were bound in clumps or
clusters to the plasma membranes of the cells (B), whereas
A/Charlottesville/31/95 progeny virions did not form large clusters but
tended to be bound in high numbers singly (D). Arrows indicate virus
aggregates. Bar, 1 µm.
sensitivities of the clinical isolate A\Charlottesville\31\95
(H1N1) and the laboratory-adapted virus A\WSN\33 (H1N1).
In the plaque-reduction assays, the IC plaque number for the
&!
A\WSN\33 virus was 0n1 µg\ml of BCX-1812. However, no
inhibition was detected for the A\Charlottesville\31\95
virus at a 100-fold higher concentration (IC plaque number &!
10 µg\ml). Additionally, the diameter of plaques produced
by A\WSN\33 was reduced by 50 % at a drug concentration
at least 100-fold lower than that required for the A\
Charlottesville\31\95 virus (IC plaque size 0n0001 µg\ml
&!
and 0n01 µg\ml, respectively). When tested in the NA enzyme
inhibition assay, the viruses A\WSN\33 and A\Charlottes-
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L. V. Gubareva and others
(a)
(b)
(c)
(d)
Fig. 2. For legend see opposite.
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Drug-selected NA deletion mutant
ville\31\95 both showed similar sensitivity to BCX-1812,
with the IC values equal to 0n20–0n23 ng\ml. These data
&!
indicate that the difference observed in the drug sensitivities
between the two wild-type viruses in MDCK cells was not
related to the enzyme.
The observed difference in the virus sensitivity to NA
inhibitor BCX-1812 was corroborated by the electron microscopy results. When propagated in the presence of a high
concentration of NA inhibitor (100 µg\ml), A\WSN\33
produced large clusters of progeny virions attached to the
surface of the infected cells (Fig. 1A, B). In contrast, the
aggregation produced by the A\Charlottesville\31\95 was
limited (Fig. 1C, D).
Drug sensitivity of the reassortant virus
To assess the HA impact on the virus sensitivity to NA
inhibitor in cell culture, we performed a gene reassortment
with the use of a reverse genetics technique (Hoffmann et al.,
2000). The reassortant virus carrying the HA gene of
A\Charlottesville\31\95, with the remaining genes from
A\WSN\33, showed reduced sensitivity to BCX-1812
(IC plaque number and diameter 10 µg\ml). These results in&!
dicated that the HA of A\WSN\33 was largely responsible
for the high drug sensitivity of this virus in cell culture.
Isolation of NA-lacking mutants
In our previous study, we observed that prolonged passage
of A\Charlottesville\31\95 in the presence of high concentrations of BCX-1812 resulted in the rapid accumulation of
defective NA genes (Nedyalkova et al., 2002). The uncloned
virus preparation contained two kinds of the NA genes, fullsized and defective ones (Nedyalkova et al., 2002). If our
assumption was correct that the HA of A\Charlottesville\
31\95 plays a key role in lowering the virus requirements for
the NA activity, we should be able to recover a mutant
containing no full-sized NA genes, with an associated loss of
NA activity. In the current studies, we therefore sought to
isolate an NA-lacking mutant and to study its properties. Three
of 19 randomly picked plaques were produced by viruses that
contained no full-sized NA genes. The progeny of one of these
clones was plaque-to-plaque purified and designated as the
delNA-31 mutant. The other NA-lacking mutant (delNA-28)
was generated by a similar approach when a clinical isolate of
A\Charlottesville\28\95 (H1N1) was passaged in the presence of oseltamivir carboxylate (Nedyalkova et al., 2002). This
NA-lacking mutant was used to confirm the observations made
for the delNA-31 mutant.
Characterization of the NA-lacking mutants
The defective NA gene of the delNA-31 mutant encoded a
95 amino acid polypeptide of residues comprising the
cytoplasmic tail, the transmembrane domain and the stalk of
the NA ; however, this defective gene did not contain genomic
information encoding the head carrying the NA active site.
Similarly, the defective NA gene of the delNA-28 mutant
encoded a peptide of 74 amino acids. Importantly, sequence
analysis detected no changes in the HA of the delNA-31 and
the delNA-28 mutants compared with their wild-type counterparts. In order to identify any possible extragenic mutations in
the genome of the delNA-31 mutant, we analysed the
sequences of the remaining six genomic segments of this
mutant and compared them with the wild-type. Only two
amino acid substitutions (Ser-409 Asn and Leu-531 Ile)
were detected ; they were located in segment 3, which encodes
the polymerase subunit PA. Sequence analysis of segment 3 of
the delNA-28 mutant indicated no mutations.
As expected, the loss of coding capacity for the NA active
site led to the loss of NA activity. When tested in the highly
sensitive fluorimetric assay, NA activity was not detected in
either the cell supernatants containing 10' TCID \ml of the
&!
delNA-31 mutant or in the approximately 100-fold-concentrated virus preparation. The NA activity of the wild-type
virus preparation (10& TCID \ml) was easily detected under
&!
the same experimental conditions. The delNA-31 mutant
produced small plaques (approximately 1 mm) in MDCK cell
monolayers and the addition of NA inhibitor into the agarose
overlay (at 100 µg\ml) affected neither the size nor the number
of plaques (not shown).
In vitro replication of the NA-lacking mutant
It has been demonstrated in previous studies (Liu et al.,
1995) that the NA-lacking NWS-Mvi mutant produces large
aggregates at the cell surface and in cytoplasmic vacuoles. A
massive virus aggregation was also demonstrated when
A\WSN\33 was propagated in the presence of NA inhibitor
FANA (Palese & Compans, 1976) and BCX-1812 (Fig. 1B). In
contrast, the aggregation of the progeny of the delNA-31
mutant was inferior and restricted to production of small virus
clumps at the surface of the infected MDCK cells (Fig. 2C, D).
To evaluate the growth characteristics of the delNA-31
mutant, MDCK cells were inoculated with either wild-type or
the mutant virus at equal m.o.i. values. The titres of the delNA31 in the cell supernatants were approximately 100-fold lower
than those of the wild-type at 60 and 72 h after infection (Fig.
3a). Of note, the titres of the delNA-31 virus associated with
the cell membranes were approximately 10-fold lower than
those of the wild-type virus (Fig. 3b). Therefore, the delNA-31
Fig. 2. Electron micrographs of MDCK cells in the absence of NA inhibitor. (A) Uninfected cells ; (B) budding and release of
A/Charlottesville/31/95 ; (C, D) limited aggregation of virions at the surface of cells infected with the delNA-31 mutant virus.
Bar, 1 µm.
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9
(a)
8
Virus titre, Log10 TCID50/ml
Virus titre, Log10 TCID50/ml
9
7
6
5
4
3
2
1
0
(b)
8
7
6
5
4
3
2
1
0
0
12
24
36
48
60
72
0
Time after infection (hours)
12
24
36
48
60
Time after infection (hours)
Fig. 3. Growth curves of the wild-type virus (
) and the delNA-31 mutant ($). MDCK cells were infected with either the wildtype A/Charlottesville/31/95 or the mutant virus, and virus yields were harvested at the indicated times. The virus titres in the
supernatant (a) and the cell-associated (b) fractions were determined. M.o.i., 0n01 TCID50/cell.
(a)
(c)
(b)
(d)
Fig. 4. Adsorption of A/Charlottesville/31/95 (a, b) and the delNA-31 mutant (c, d) to the MDCK cell monolayer. In (a) and
(c), adsorption was carried out at 0 mC for 20 min. In (b) and (d), adsorption was carried out at 0 mC for 60 min, followed by
adsorption at 37 mC for 60 min. Adsorption was followed by removal of the inoculum and incubation at 37 mC for 7 h. The
samples were then processed for immunofluorescence, and infected cells were detected using a mixture of anti-NP monoclonal
antibodies.
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Drug-selected NA deletion mutant
(a)
wt 31
delNA-31
wt 28
(b)
delNA-28
HA0
NA
NP
NA dimer
M1
M1
(c)
wt 31
delNA-31
wt 28
delNA-28
HA0
NP
wt 31
+PNG -F
delNA-31
+PNG -F
HA0
NA
NP
(d)
wt 31
+Sialidase
HA0
HA0*
delNA-31
+Sialidase
HA0-sia
NP
NA*
M1
M1
Fig. 5. Protein profile analysis by SDS–PAGE. Plates of MDCK cells were infected with the wild-type viruses and the
corresponding delNA mutants in the absence of trypsin. Four hours after infection, 25 µCi [35S]methionine was added to the
cells and the supernatants were harvested at 20 h post-infection. After the cellular debris had been removed, the virus particles
were pelleted and subjected to electrophoresis on SDS–polyacrylamide gels. (a) 12 % SDS–PAGE under reducing conditions.
Lane 1, A/Charlottesville/31/95 ; lane 2, delNA-31 mutant ; lane 3, A/Charlottesville/28/95 ; lane 4, delNA-28 mutant. All
viruses contained HA0, NP and M1 proteins, but NA could be detected only in the wild-type viruses. NP migrated as two bands.
(b) 12 % SDS–PAGE under non-reducing conditions. Lane designations are the same as those in (a). The mobility of HA0 from
the delNA viruses was reduced in comparison to those of the wild-type viruses. The NA monomers and NA dimers were seen in
the preparations of the wild-type viruses but not in those of the delNA mutants. (c) 12 % SDS–PAGE under reducing conditions
following pretreatment. Lanes 1, A/Charlottesville/31/95 ; lane 2, A/Charlottesville/31/95 pretreated with PNG-F ; lane 3,
delNA-31 mutant ; lane 4, delNA-31 mutant treated with PNG-F. A shift in the mobilities of the HA0 monomers of both viruses
was observed following pretreatment (HA0*) ; the pretreated HA0 of both viruses exhibited similar mobilities. A shift in the
mobility of the NA monomer was also seen in the wild-type virus (NA*) following pretreatment. (d) 10 % SDS–PAGE under
non-reducing conditions. Pretreatment of the wild-type virus and the delNA mutant with bacterial sialidase reduced the HA0
mobility of the delNA-31 mutant (HA0-sia HA0) but not that of the wild-type virus, indicating the presence of terminal
neuraminic (sialic) acid residues on the HA0 of the NA-lacking mutant.
mutant was capable of release, although less efficiently
compared with the wild-type.
Binding efficiency of the NA-lacking mutant
To confirm the reduced HA binding efficiency for MDCK
cell receptors, we tested the viruses in an assay previously
utilized by others to characterize NA inhibitor-resistant
mutants (Blick et al., 1998). MDCK cells were inoculated with
either the wild-type or the delNA-31 mutant at the same m.o.i.
After time-controlled adsorption, the infected cells were
incubated for 7 h and stained with antibodies against the viral
nucleoprotein (NP). For the wild-type virus, 20 min of
adsorption was sufficient to produce approximately 50 % NPpositive cells compared with its control (Fig. 4a, b). In contrast,
only 15 % of cells were positive for NP in the delNA-31-
infected monolayers after 20 min of adsorption (Fig. 4c)
compared with its control (Fig. 4d). Therefore, these data
indicate that the delNA-31 mutant demonstrated substantially
reduced adsorption efficiency compared with the wild-type.
Sialylation of the HA
Next we analysed the wild-type viruses and the delNA
mutants by SDS–PAGE. As expected, there were no bands
corresponding to the full-sized NA in the lanes containing the
delNA-31 and delNA-28 mutants under reducing (Fig. 5a) and
non-reducing conditions (Fig. 5b). The electrophoretic mobilities of the HA (HA precursor) bands of the wild-type viruses
!
were slightly greater than those of the delNA mutants (Fig.
5a) ; this difference in mobility was especially apparent under
non-reducing conditions (Fig. 5b). To remove oligosaccharide
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L. V. Gubareva and others
chains attached to asparagines of the viral glycoproteins, we
pretreated virus preparations with peptide-N-glycosidase F
(PNG-F). Removal of oligosaccharide chains was accompanied
by a shift in the band corresponding to the HA and NA of the
!
wild-type virus. After treatment, the mobilities of the HA of
!
the wild-type and the delNA-31 viruses became equal (Fig. 5c).
The pretreatment of viruses with exogenous sialidase (from
Clostridium perfringens) also abolished the difference in the
electrophoretic mobilities of the HAs (Fig. 5d). Therefore, the
difference in the HA mobilities of the wild-type and the
!
delNA viruses appeared to be caused by the presence of
terminal neuraminic acid moieties on the HA of the delNA
mutants in the absence of NA activity.
Discussion
Here, we present direct evidence that the HA of a recent
human influenza A (H1N1) virus plays a key role in its low
sensitivity to NA inhibitors in MDCK cells. Moreover, the
reduced need for NA activity led to the emergence of NAlacking mutants when this virus was passaged in cell culture in
the presence of an NA inhibitor. Baigent et al. (1999) showed
that sensitivity to NA inhibitors of naturally occurring avian
influenza A viruses is determined by both the HA and NA. To
demonstrate the role of HA in the human virus sensitivity to
NA inhibitor, we used a reverse genetics technique to generate
a reassortant virus, carrying the HA gene from A\Charlottesville\31\95 (H1N1) and the remaining genes of A\WSN\33
(H1N1). This approach ensured that no unwanted changes had
been introduced into the genome of the reassortant virus that
could affect the drug phenotype. We demonstrated that
replacement of the HA gene of the highly drug-sensitive
A\WSN\33 (H1N1) drastically reduced the sensitivity of
A\WSN\33 in cell culture. Furthermore, we found that
glycosylation of HA from the recent clinical isolate differed
substantially from that of A\WSN\33. Analysis of the HA
sequences of the A\WSN\33 and A\Charlottesville\31\95
isolates revealed that the recent human strain had twice as
many potential glycosylation sites in its HA subunit (eight
"
compared with four). Some of these oligosacharide chains
reside in the vicinity of the RBS (i.e. Asn-129 and Asn-163 ; N1
numbering). HA hyperglycosylation is not confined only to
the two clinical isolates used in the present study but is a
characteristic trait of the recent human influenza viruses of
H1N1 subtype (Inkster et al., 1993). The regulatory function of
the N-glycans on the HA molecule in virus growth and in its
requirement for NA activity has been extensively investigated
in several laboratories (Baigent et al., 1999 ; Matrosovich et al.,
1999 ; Wagner et al., 2000 ; Baigent & McCauley, 2001).
Ohuchi et al. (1997) demonstrated that adsorption of erythrocytes to the viral HA could be reduced by the presence of
oligosaccharide chains in the vicinity of the RBS, and the
presence of uncleaved neuraminic acid moieties on these chains
reduces the HA binding still further (Ohuchi et al., 1997).
CGJA
The question arises as to whether sialylation of the HA
alone can render the virus resistant to NA inhibitors. The
number of plaques produced by A\Charlottesville\31\95 in
MDCK cells was not affected by the presence of the NA
inhibitor at the highest concentration tested (10 µg\ml). This
result demonstrated that this virus had over 100-fold reduced
sensitivity to NA inhibitor compared with A\WSN\33 in cell
culture. Nevertheless, the diameter of plaques produced by
A\Charlottesville\31\95 was inhibited at lower drug concentrations (0n01 µg\ml) indicating a restricted spread of the virus
under agarose overlay. None the less, in liquid media, the NA
independence of A\Charlottesville\31\95 allowed the emergence of a mutant completely lacking the coding capacity for the
NA active site. Therefore, it was important to investigate
whether the NA-lacking mutant had acquired any additional
compensatory mechanisms besides the sialylated oligosaccharide chains in the vicinity of the RBS. We found that the
genome of the mutant contained only the defective NA genes
that arose as a result of a massive internal deletion. In this
respect, it was similar to the NA-lacking mutants generated by
others under conditions when the endogenous NA activity
was not required (Liu & Air, 1993 ; Hughes et al., 2000, 2001).
We did not attempt to detect the products of the defective NA
genes in the infected cells or the delNA virus because the
previous extensive studies by Yang et al. (1997) produced no
evidence for the expression of defective NA genes in cells
infected with the other NA-lacking mutant, NWS-Mvi (Yang
et al., 1997). Importantly, sequence analysis of the delNA-31
genome detected no amino acid substitutions, which could
have provided additional compensation for the loss of NA
activity. The only two amino acid substitutions found were in
the PA polymerase subunit. No substitutions were detected in
the PA of the other NA-lacking mutant (delNA-28) generated
under similar conditions. Therefore, we believe that the lower
binding efficiency of the delNA-31 mutant compared with the
wild-type (Fig. 4) was directly by sialylation of the oligosaccharide chains of the HA (Fig. 5). The delNA-31 mutant was
able to elute from infected cells and produced only small
clumps of progeny virions at the cell surface (Fig. 2).
Importantly, the delNA-mutant produced a substantial yield of
infectious virus in the supernatant fraction, although not as
high as the wild-type virus (Fig. 3a). The considerable NA
independence of the delNA-31 mutant separates this virus
from the other NA-lacking viruses generated on the basis of
the NWS-G70c reassortant (H1N9) (Liu et al., 1995 ; Hughes et
al., 2000). The difference in the number (four versus eight) of
the oligosaccharide chains in the HA of NWS-Mvi and
"
delNA-31 could be one explanation. It is noteworthy that the
egg-grown A\Texas\36\91 (H1N1) variant was highly
sensitive to NA inhibitors in cell culture despite the presence of
oligosaccharide chains in the vicinity of its RBS. A plausible
explanation for this is the acquisition of amino acid substitutions in the RBS caused by egg-adaptation (Gubareva et al.,
2001a).
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Drug-selected NA deletion mutant
The emergence of NA-lacking mutants following virus
passage in the presence of NA inhibitor provides additional
evidence that NA activity is not obligatory when virus is
propagated in vitro (Liu et al., 1995). However, the reduced
need for NA activity demonstrated for the recent influenza A
viruses in the present study does not necessarily imply that
these viruses are resistant to NA inhibitors during replication
in humans. The NA function in vivo is not restricted to the
prevention of virus aggregation at the surface of infected cells,
but is also needed to avoid virus entrapment by the respiratory
tract secretions and thus elimination. Furthermore, the receptors present on MDCK cells do not adequately reflect those
present on the human respiratory tract epithelium (Baum &
Paulson, 1990 ; Govorkova et al., 1995). Treatment with NA
inhibitors was beneficial for individuals experimentally infected
with the egg-adapted influenza A\Texas\36\91 (H1N1)
(Hayden et al., 1996, 1999), although the HA of this virus has
"
the same number of potential glycosylation sites as A\Charlottesville\31\95. Thus, hyperglycosylation of the HA did not
abolish the inhibitory effect of the NA inhibitors on the eggadapted virus replicating in a human host. Whether non-eggadapted viruses with the hyperglycosylated HA are less
sensitive to NA inhibitors in humans remains to be seen.
We thank Dr Robert G. Webster (St Jude Children’s Research
Hospital, TN) for valuable discussions, Dr Julia Cay Jones (St Jude
Children’s Research Hospital) for editorial help and Douglas Schallon and
Kimberly Underwood from the University of Virginia for excellent
technical assistance. This work was supported in part by a grant (AI45782) from the National Institute of Allergy and Infectious Diseases
(L. V. G.), by a grant from the R. W. Johnson Pharmaceutical Research
Institute (L. V. G.) and by the American Lebanese Syrian Associated
Charities (K. G. M.).
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Received 12 March 2002 ; Accepted 13 June 2002
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