Mutations That Hamper Dimerization of Foot-and-Mouth Disease
Virus 3A Protein Are Detrimental for Infectivity
Mónica González-Magaldi,aRaúl Postigo,aBeatriz G. de la Torre,bYuri A. Vieira,aMiguel Rodríguez-Pulido,aEduardo López-Viñas,a,c
Paulino Gómez-Puertas,aDavid Andreu,bLeonor Kremer,dMaría F. Rosas,aand Francisco Sobrinoa,e
Centro de Biología Molecular Severo Ochoa (CSIC-UAM), Cantoblanco, Madrid, Spaina; Departament de Ciències Experimentals i de la Salut, Universitat Pompeu Fabra,
Barcelona, Spainb; Biomol-Informatics SL, Madrid, Spainc; Centro Nacional de Biotecnología (CNB-CSIC), Cantoblanco, Madrid, Spaind; and CISA-INIA, Valdeolmos, Madrid,
FMDV 3A homodimerization was evidenced by an in situ protein fluorescent ligation assay. A molecular model of the FMDV 3A
(5, 24, 52) and the etiological agent of a devastating disease of
livestock (34). The viral particle is composed of a protein capsid
that contains a positive-sense RNA molecule of about 8,500 nu-
proteins (9, 62). The viral genome encodes four structural capsid
and P3 (3A, 3B, 3C, and 3D) regions (9).
Replication of picornaviruses occurs associated to cell endo-
teins are involved in crucial aspects of the viral cycle and patho-
genesis, such as rearrangements of intracellular membranes
tion complex and intracellular membranes appears to be accom-
plished by proteins 3A and 2C, which have membrane-binding
properties (11, 60). When expressed as a recombinant protein in
transfected cells, PV 3A cofractionates with endoplasmic reticu-
lum markers (66), and its single transient expression can disrupt
the secretory apparatus (23) and decrease major histocompatibil-
ity complex (MHC) class I expression (22). On the other hand,
3AB presumably anchors 3B in intracellular membranes origi-
nated de novo during the early steps of RNA replication, where
uridylylated 3B primes the synthesis of nascent viral RNAs (2, 37,
ciates with the cloverleaf structure in the 5= end of viral RNA and
ber of the aphthovirus genus within the family Picornaviridae
with the 3CD precursor to form a ribonucleoprotein complex
required for PV RNA synthesis (32, 74, 76).
While FMDV shows considerable functional and structural
analogies with PV and other picornaviruses, some differences
have been reported, such as its resistance to the Golgi disruption
2BC, instead of 3A, to inhibit the secretory pathway in cultured
immune responses (58). In addition, FMDV is the only picorna-
virus encoding 3 copies of 3B protein, required for both optimal
replication in cell culture (26) and for virulence in natural hosts
(50). All 3 copies have been shown to undergo uridylylation in
vitro (44). In FMDV, 3A protein has been reported to play a role
on viral host range, as a single amino acid replacement (Q44R)
conferred FMDV the ability to cause vesicular lesions in guinea
a hydrophobic domain that in other picornaviruses mediates its
sible for the location of the replication complex within a mem-
brane context (20, 23, 27, 69). The C-terminal fragment of 3A
Received 6 March 2012 Accepted 3 July 2012
Published ahead of print 11 July 2012
Address correspondence to Francisco Sobrino, email@example.com.
M.G.-M. and R.P. contributed equally to the work.
Copyright © 2012, American Society for Microbiology. All Rights Reserved.
October 2012 Volume 86 Number 20Journal of Virologyp. 11013–11023 jvi.asm.org
longer in FMDV (77 amino acids [aa]) than in other picornavi-
ruses (i.e., 7 aa for PV), and deletions and mutations in 3A are
known to contribute both to viral attenuation in cattle (8) and to
decreased replication rates in bovine epithelial cells (49). How-
ever, little is known on the interactions of 3A with other viral and
cellular proteins, and no structural data are available for this pro-
Homodimer formation has been revealed by the NMR struc-
domain (aa 1 to 59) of PV 3A (65). Each monomer has a struc-
tured region consisting of two amphipathic ?-helices (aa 23 to 29
flanked by nonstructured N and C regions. The N terminus of 3A
top of the dimer structure, in the loop between the two ?-helices,
as well as three solvent-exposed charged residues (E38, K39, and
homodimer formation in both RNA replication and inhibition of
cellular protein transport has also been reported for coxsackievi-
rus (CV) B3. In this case, while the general organization of the
CVB3 dimer was similar to that of PV, the establishment of salt
bridges between residues D24 and K41 was found critical for
dimer stability; using an optimized PV 3A structure, these salt
bridges were also found in equivalent PV residues (D23 and K40)
To gain insight into the structure-function relationship of
FMDV 3A protein, we devised a molecular model for the N-ter-
minal region of this protein, using as the template the structure
reported for PV 3A. This model predicted hydrophobic interac-
tions between residues at two ?-helices in each monomer as the
main homodimerization determinant. Here, we show that amino
phobic dimerization interface, and expected to contribute to
dimer stability, decrease 3A dimerization in cells transiently ex-
pressing 3A, and abolish dimer/multimer formation in peptides
reproducing the N terminus of 3A. Replacements L38E and L41E
significantly reduced the homodimerization signal detected for
transiently expressed 3A by means of an in situ proximity ligation
assay (63). In addition, replacements L38E and L41E were detri-
mental for virus growth, leading to selection of viruses that for
mutants L38E and L41E restored the hydrophobicity of the resi-
dues, suggesting that 3A dimer formation plays a relevant role in
FMDV replication. On the other hand, replacement Q44R that
favors or replacement Q44E that impairs the polar interactions
that, according to the model, Q44 could establish with residue
transiently expressed 3A, indicating that these polar interactions
are not critical for 3A dimerization. Nevertheless, while Q44R led
to infectious virus recovery, Q44D resulted in the selection of
infective viruses with substitution D44E with acidic charge but
with structural features similar to those of the parental virus, sug-
gesting that residue Q44, despite not being essential for 3A
MATERIALS AND METHODS
Modeling procedures of FMDV 3A proteins. Three-dimensional (3D)
were generated using comparative modeling procedures. As a structural
template, the soluble domain of human PV 3A protein was used, attend-
Data Bank code 1NG7) (65). Best representative conformer in the NMR
spectroscopy ensemble was selected as a source of coordinates for the
FMDV 3A wild type and subsequent models. All models were built using
.org//SWISS-MODEL.html, and their structural quality was checked using
model quality estimation was performed in terms of QMEAN4 raw score
(10). To optimize geometries, models were energy minimized using the
steepest descent minimization followed by 500 steps of conjugate-gradient
minimization. Figures were generated using the Pymol Molecular Graphics
System (Schrödinger, LLC). Structurally based sequence alignment of wild-
type model templates and other 3A proteins was refined with T-COFFEE
well as four analogues with the M29R, L38E, L41E, and L38E-L41E re-
placements (N-ter peptides; see Table 1), were made in C-terminal car-
(Protein Technologies, Tucson, AZ). Standard 9-fluorenylmethoxy car-
amounts of Fmoc-amino acids and O-benzotriazole-N,N,N=,N=-tetram-
ethyluronium hexafluorophosphate and 10-fold molar amounts of N,N-
diisopropylethylamine, with systematic double coupling for all residues.
pylsilane (95:2.5:2.5 [vol/vol], 90 min, room temperature), peptides were
precipitated with chilled diethyl ether, redissolved in water, lyophilized,
and purified by preparative reverse-phase high-performance liquid chro-
matography (HPLC) to a minimum purity of 95% by analytical HPLC.
Their identities were confirmed by matrix-assisted laser desorption ion-
ization–time of flight (MALDI-TOF) mass spectrometry in a Voyager
namic acid matrix and linear mode acquisition (Table 1).
For SDS-PAGE analysis, aliquots containing 1.87 ?g and 0.187 ?g of
each peptide were prepared from 5-mg/ml stock solutions of the lyophi-
lized peptides in Milli-Q water, mixed with Laemmli sample buffer con-
taining 2% SDS and 100 mM dithiothreitol (DTT), and incubated for 25
stained with Coomassie blue, transferred to nitrocellulose membranes,
a goat anti-rabbit horseradish peroxidase-coupled secondary Ab (GE
Healthcare). Samples were subsequently developed using a chemilumi-
nescence kit (Perkin-Elmer).
TABLE 1 Synthetic peptide sequences
Wild type or mutationAmino acid sequencea
Avg mol wt (calculated/found)
aSequences correspond to residues 1 to 52 of the 3A protein. Bold, underlined letters refer to replacements relative to the 3A sequence of FMDV C-S8c1.
González-Magaldi et al.
jvi.asm.orgJournal of Virology
and maintained in Dulbecco’s modified Eagle’s medium (DMEM)
(Gibco-BRL), supplemented with 5% fetal bovine serum (Gibco-BRL), 1
?g/ml streptomycin, and 1 ?g/ml penicillin. A viral stock from a type C
FMDV C-S8c1 (61) isolate was produced by amplification in BHK-21
cells. Procedures for infections and virus titration in semisolid agar me-
dium were as described previously (54).
protein 3A (21) was used in the proximity ligation assay. Rabbit polyclonal
anti-?-tubulin 196 was used in Western blot analysis (4). Anti-rabbit IgGs
assays, was from Molecular Probes (Invitrogen). Polyclonal Ab 443 against
GMVHDSIKEELRPLIQ) coupled to Cys-KLH. To generate polyclonal
Ab 346 against the C terminus of 3A, plasmid pRSET-C (Invitrogen) was
used for the inducible isopropyl-?-D-thiogalactopyranoside (IPTG) ex-
pression in Escherichia coli BL21(DE3)pLysS of a His-tagged polypeptide
spanning 3A residues 81 to 153 fused to 3BBB, whose sequence was am-
GAAGATGGTG-3= and 5=-GCAGATCTTTACTCAGTGACAATCAA-
3=. The restriction enzymes NheI and BglII (New England Biolabs) were
used for cloning. The His-tagged protein was purified in an Ni column
(Probond resin; Invitrogen) with a pH 4.5 elution buffer, and two New
Zealand White rabbits were immunized with the purified protein. Rabbit
immunizations were as described previously (54).
Plasmids. To assess their effect on FMDV infectivity, 3A mutations
were introduced in plasmid pMT28 encoding the type C FMDV isolate
C-S8c1 full-length sequence (29). Substitutions of selected amino acids
were performed by site-directed mutagenesis (41), using the primers
shown in Table 2. To this end, a 3A-containing fragment was amplified
from pMT28 using primer 3A-F and the corresponding mutated reverse
primer (-R) or primer 3A-R and the corresponding mutated forward
primer (-F) at a 1 ?M concentration, in a reaction, including the Expand
high-fidelity PCR system BioTaq (Bioline) polymerase (1.25 U), plus Pfu
triphosphate (dNTPs) (Roche). After amplification, each PCR fragment
was purified and an overlap extension PCR was performed using outer
and RsrII and cloned into pMT28 digested with the same restriction en-
of the resulting plasmids were confirmed by sequencing with primers
3A-F and 3A-R (Table 2), as well as primers 3A-1 and 3ABB-2 (30). For
transient expression, plasmid pRSV3Awt (30) and derivatives of pRSV/L
different mutants described above were used. To this end, 3A-containing
sequences were amplified from the corresponding pMT28 derivatives us-
ing oligonucleotides 3A-1 and 3A-2 (30), which included restriction sites
for cloning into pRSV (HindIII and KpnI).
RNA synthesis, transfection, and infectivity. For RNA synthesis,
plasmids were linearized with NdeI (New England BioLabs) and in vitro
transcribed using SP6 RNA polymerase (Promega). After transcription,
the reaction mixture was treated with RQ1 DNase (1U/?g RNA; Pro-
mega). Then, RNA was extracted with phenol-chloroform and precipi-
tated with ethanol. The RNA integrity and concentration were deter-
described previously (40). Cells were maintained at 37°C in Dulbecco’s
modified Eagle’s medium (DMEM)-supplemented fetal bovine serum
(FBS). To assess RNA infectivity, at 4 h posttransfection (hpt) cells were
maintained in semisolid DMEM-0.5% agar supplemented with 5% FBS.
At 24 hpt, the viral titer was determined by plaque assay (54).
Viral RNA extraction, cDNA synthesis, and DNA sequencing. Viral
RNA was extracted from supernatants of cell cultures or tissue homoge-
nates from suckling mice (7), using TRI reagent (Sigma), as described by
3A-R. cDNA was amplified by PCR using the same primers and BioTaq
ity polymerase (Biotools) for proofreading activity. PCR products were
purified with Wizard SV gel and PCR clean-up system (Promega) and
their sequences determined by automatic DNA sequencing. DNA se-
5 of 3B2 viral proteins, were confirmed by at least two independent se-
quencing reactions using the primers indicated above. Nucleotide posi-
tions correspond to those previously described for the FMDV C-S8c1
isolate (GenBank no. AJ133357).
15 min at room temperature, as described previously (56). Translated
proteins were analyzed by 10% SDS-PAGE.
(OLINK Bioscience). Briefly, IBRS cells were grown on glass coverslips
and transfected with 1 ?g of pRSV3Awt and derivatives using Lipo-
fectamine (Invitrogen) as described by the manufacturer or infected with
FMDV C-S8c1 at a multiplicity of infection (MOI) of 5 PFU/cell. At 24
hpt, or 3 h postinfection (hpi), monolayers were washed with phosphate-
buffered saline (PBS), fixed in 4% paraformaldehyde for 15 min at room
X-100, 1% bovine serum albumin [BSA], and 1 M glycine in PBS) for 15
min at room temperature. Primary Ab was prepared using the Probe-
maker kit (OLINK, Bioscience) by using the manufacturer’s recommen-
dations: 1 mg/ml of monoclonal Ab (IgG class) 2C2, affinity purified
through a protein G column, was independently conjugated to each of a
pair of oligonucleotides to generate plus and minus proximity ligation
assay (PLA) probes. Cells on coverslips were incubated with conjugated
primary antibodies diluted in PBS with 1% BSA for 1 h at room temper-
0.05% Tween 20) for 5 min. The signal development (ligation, amplifica-
tion, and hybridization) was performed according to the manufacturer’s
instructions in PLA Duolink II. Briefly, a ligation-ligase solution was
added to each sample, and slides were incubated in a preheated humidity
chamber for 30 min at 37°C and washed twice in buffer A for 2 min with
gentle agitation. Then, the amplification-polymerase solution, including
the fluorescent probe (?excof 554 nm), was added and slides were incu-
slides were washed three times in buffer B (0.2 M Tris, 0.1 M NaCl [pH
TABLE 2 Oligonucleotides used for construction of mutants in the 3A
aForward and reverse primers are indicated by F and R, respectively.
bSequences in bold and nucleotide numbering correspond to those of FMDV C-S8c1
(GenBank no. AJ133357), and substituted nucleotides are in italics. Restriction sites are
indicated, and their nucleotides are underlined.
Determinants of Dimerization on FMDV 3A Protein
October 2012 Volume 86 Number 20jvi.asm.org 11015
7.5]) for 10 min with agitation and protected from light. In the case of
polyclonal Ab 346 and anti-rabbit IgG secondary antibodies coupled to
Alexa Fluor 647 (Invitrogen) to detect 3A protein. Slides were mounted
with DAPI (4=,6-diamidino-2-phenylindole). Cells were observed with a
confocal LSM710 vertical (Bio-Rad/Zeiss) microscope. As reported for
ImageJ software (analyze particles plug-in). The percentage of the fluo-
rescence intensity, relative to that determined for cells expressing 3Awt
protein, was plotted for each mutant protein ? standard error.
plates and transfected as described above with 1 ?g of different plasmids.
At 24 hpt, the cells were scraped on ice into NP-40 lysis buffer (10 mM
EGTA, 2.5 mM MgCl2, 1% NP-40, 20 mM HEPES [pH 7.4]) and soni-
cated. Equal amounts of total protein of each sample, estimated by Brad-
ford (13), were mixed with Laemmli sample buffer and boiled. Samples
were separated by 12% SDS-PAGE and transferred onto a nitrocellulose
membrane. The membrane was blocked, and proteins were detected by
incubation with the corresponding 3A-specific polyclonal rabbit Ab as
ling mice was tested as described previously (7). Briefly, in vitro-tran-
scribed viral RNA (from 102to 105ng) mixed with 20 ?l of Lipofectin
(Invitrogen) in a final volume of 100 ?l in PBS was inoculated intraperi-
toneally in litters (4 to 5 mice per RNA) of Swiss newborn mice. Dead
animals were scored up to 11 days after infection. Mice showing severe
signs of disease (tremors ataxia, paralysis of the hind limbs) were eutha-
nized. All animals in this study were handled in the BSL-3 facilities at
CISA-INIA (Madrid, Spain), in strict accordance with the guidelines of
the European Community 86/609/CEE. The protocol was approved by
the Committee on the Ethics of Animal Experiments of INIA (permit
number CBS 2008/016).
Data analysis. One-way analysis of variance was performed with sta-
means ? standard errors.
Homology model of the 3A protein and dimerization interface.
of the FMDV 3A protein, 3D structural models of its N-terminal
region (residues 1 to 94) were generated by bioinformatic proce-
dures, using the analogous structure of the soluble domain of
human poliovirus 3A protein (65) as the template. As a result,
compatible ungapped models were rendered for residues 15 to 48
of the FMDV 3A protein, in correspondence to the structured
region in solution from the experimental template. Quality
QMEAN4 raw scores for the model of wild-type protein mono-
mers in this region were 0.622 (Z score ? 0.09) and 0.621 (Z
mainly on the structural basis of two ?-helices (?1, residues 25 to
33, and ?2, residues 37 to 44) connected by a 3-residue loop (Fig.
1A and B), hypothetically responsible for the dimerization pro-
predictions (6, 33, 70) points to the region comprised between
residues 59 to 76 as putatively responsible for 3A protein anchor-
IDs are given in parenthesis): 1NG7, poliovirus type 1 strain Mahoney (P03300), sequence used as the template for homology modeling procedures (65); PV1,
poliovirus type 1, CHN-Guangdong/92-2 isolate (Q9E912); CVB3 (P03313); CVA11 (Q7T7P4); CVA20 (Q7T7N8); FMDV type C, C-S8c1 isolate, (Q9E2G4);
FMDV type O (P03305); FMDV type A (P49303); FMDV type Asia 1 (Q7TDB3). Positions in the alignment are colored according to sequence conservation by
average BLOSUM 62 score (similar residues according to BLOSUM 62 score are colored as the most conserved one using standard Belvu colors: ?0.50, white;
0.51 to 1.50, light gray; 1.51 to 3.00, medium blue; ?3.00, cyan) (http://sonnhammer.sbc.su.se/Belvu.html) (64). Positions of D32 and Q44 residues in the
FMDV-3A modeled sequence are indicated by magenta arrows. Position of hydrophobic residues M29, L38, and L41 are marked by green arrows. (B) General
left to right and top to bottom: M29R, L38E, L41E, and the relative positions of D32 and Q44 residues (mutants Q44D and Q44R).
González-Magaldi et al.
jvi.asm.orgJournal of Virology
quaternary organization of the N-terminal region protein similar
dimerization interface in FMDV 3A would include all residues in
the 25-to-44 stretch of the two antiparallel monomers. As in PV
could provide physical stability to the dimer. Such interactions
would involve residues F25, F26, M29, V30, L38, L41, and I42, in
both monomers. Genotypes carrying mutations introducing sim-
ilar electrostatic repulsive interactions in the hydrophobic core
tial local accumulation of heterogeneous repulsive interactions.
Thus, mutations L38E, L41E, and M29R within the hydrophobic
core could disrupt the binding interaction in the region by creat-
ing repulsions from facing negatively (or positively) charged res-
idues (Fig. 1C). On the other hand, polar contacts such as that of
residues D32 and Q44 (Fig. 1C), flanking the C-terminal ends of
to mediate FMDV adaptation to the guinea pig (48), could signif-
icantly modify the electrostatic surface of the region, favoring the
formation of salt bridges between both R44 and D32 residues and
eventually increasing dimer stability.
highly conserved among FMDVs of different serotypes (16), were
selected to be replaced by a negatively charged E residue, which
would interfere in the hydrophobic interactions predicted in this
Substitutions at the predicted hydrophobic interface affect
3A dimerization. Identification of protein bands potentially cor-
responding to 3A dimers in FMDV-infected cells is impaired by
the presence of protein precursors from the 3ABBB region (30).
Therefore, we first addressed the effect of the substitutions se-
tive of 3A dimer/multimer formation were also observed. When
cells were transfected with pRSV derivatives expressing 3A with
mutations M29R, L38E, or L41E or double-mutant L38EL41E,
observed in cells expressing mutant proteins Q44D and Q44R,
with the intensity of the dimer band in both cases slightly lower
than that of 3Awt.
To further characterize 3A homodimerization, we set up an in
situ protein ligation assay previously used to visualize protein-
protein interaction in the cell by fluorescence microscopy (3, 28,
45). The strategy, detailed in Materials and Methods and summa-
rized in Fig. 3A, is based on coupling a monoclonal Ab that rec-
ognizes a single epitope on each 3A molecule to two different
oligonucleotide probes, ? and ?, which subsequently hybridize
would ligate, forming a circle; amplification of this circle upon
addition of fluorescently labeled nucleotides allows visualization
of each single interaction by fluorescence microscopy (71). The
monoclonal Ab selected for this assay was directed against an
epitope on the C-terminal region of 3A, distant from the N-ter-
minal region analyzed to avoid loss of recognition due to the mu-
tations introduced. The assay included a biological control of
one of the probes each. As shown in Fig. 3B, when cells were
FMDV infected and incubated with probes 2C2? and 2C2?, a
dimer fluorescent signal was observed. Fluorescence was also no-
the fluorescence intensity, of about 40% of that of 3Awt, were
detected in cells transfected with mutant plasmids pRSV3AL38E
and pRSV3AL41E (Fig. 3C and D). When plasmid pRSV3AL38E-
tion observed was higher, at about 80% (Fig. 3C). These results
support the capacity of 3A to form homodimers and suggest that
replacements L38E and L41E impaired dimer formation.
mation by synthetic peptides spanning the 3A dimerization in-
terface. To study the potential of the N terminus of 3A to form
FIG 2 Analysis of 3A proteins and peptides, including mutations potentially af-
incubated with polyclonal Ab 346 generated against the C terminus of 3A. The
protein bands whose migrations correspond to the monomeric and the dimeric
forms of 3A are indicated with arrows. Membranes were reprobed with a MAb
against ?-tubulin, as a control for protein loading. (B) Analysis of synthetic pep-
parallel gels, one of which was stained with Coomassie blue. The second gel was
transferred to nitrocellulose membrane and blotted with polyclonal Ab 443 pro-
duced against the N terminus of 3A. The migrations and sizes of the molecular
Determinants of Dimerization on FMDV 3A Protein
October 2012 Volume 86 Number 20jvi.asm.org 11017
dimers and multimers, peptides spanning residues I1 to F52
(termed N peptides) were synthesized by solid-phase procedures,
as described in Materials and Methods. The electrophoretic mo-
bility of peptide N-wt, spanning the sequence corresponding to
the parental C-S8c1 FMDV, revealed by Western blotting with an
Ab against the N terminus of 3A protein, showed a band of a size
about that corresponding to the monomeric peptide, as well as
additional bands corresponding to higher-order forms (Fig. 2B).
A pattern of bands similar to that observed by Western blotting
was revealed by mass staining; in this case, the monomeric form
was the major band, indicating that the Ab used preferentially
recognized dimer/multimeric peptide forms. The inverse rela-
of Ab recognition due to epitope multimerization. On the other
hand, only monomeric bands were detected by mass staining of
mutant peptides, which showed slight differences in mobility, as
reported to occur among 48-mer peptides differing in single resi-
of higher-order forms, N-M29R being the only peptide for which
a faint dimeric band was observed. The poor Ab recognition of
peptide N-L38E could be explained by either an altered confor-
mutated residue in the peptide-Ab interaction.
form dimers and multimers, (ii) its N-terminal fragment is in-
volved in this process, and (iii) replacement of the hydrophobic
ity. To assess the effect of replacements L38E and L41E located at
the predicted hydrophobic dimerization interface on the FMDV
life cycle, the infectivity of mutant RNAs transcribed from plas-
mids pMT28L38E and pMT28L41E carrying these substitutions
was compared with that of the corresponding parental C-S8c1
was determined after 24 h of transfection with different amounts
in semisolid medium. No infectious virus was recovered after
transfection of cells with up to 1 ?g of RNA transcribed from
pMT28L38E or pMT28L41E (data not shown), indicating that
RNAs with these mutations showed an infectivity at least five log
lower than that of the RNA control pMT28. When transfected
cells were incubated in liquid medium, a delay in the recovery of
infectious virus, relative to transcript pMT28, was observed for
transcripts pMT28L38E and pMT28L41E (no infective virus was
were detected at 72 hpt, respectively) (Fig. 4). Upon two addi-
tional passages of the transfection supernatant, the viral titers re-
FIG 3 Proximity ligation assay to confirm homodimer protein complexes. (A) The scheme summarizes the procedure: (i) conjugation of primary monoclonal
Ab 2C2, with PLA probes ? (red) and ? (blue); (ii) incubation of the sample where dimerized proteins are represented in green and cherry, with the two
conjugates of monoclonal Ab 2C2, probes ? and ?; (iii) hybridization and ligation reaction with oligonucleotides complementary to the PLA probes; (iv) final
from left to right: (i) PLA reaction in cells infected with FMDV (MOI of 5 PFU/ml); negative controls of infected cells incubated with (ii) 2C2-probe ? or (iii)
2C2-probe ?; and (iv) PLA reaction in mock-infected cells with probes ? and ?. (C) Plot of percentage of fluorescence intensity, relative to that of cells
n ? 40 cells scored in the experiment. Mean values and standard errors are represented. Statistically significant differences, relative to pRSV3Awt, are indicated
by * (P ? 0.05) or ** (P ? 0.005). (D) PLA reaction (orange) in cells transfected with the plasmids in panel C (24 hpt) and incubated with rabbit polyclonal Ab
346 (red) detected with Alexa Fluor 647 anti-rabbit Ab. Nuclei were stained with DAPI. Scale bar, 20 ?m.
González-Magaldi et al.
jvi.asm.orgJournal of Virology
covered from transcripts pMT28L38E and pMT28 were simi-
lar, while those recovered from transcript pMT28L41E were
about 1 log lower (Fig. 4). Emergence of infectious virus was
concomitant with the imposition in the viral populations of
replacements for each of the mutant RNAs analyzed—E38V
and E41A—that restored the hydrophobicity of the residues, as
determined by sequencing of the viral RNA at the second cell
passage given to the transfection medium (Table 3). Selection
of mutant E38V was observed in the two additional transfec-
tions of RNA from pMT28L38E performed. On the other hand,
infectious virus was recovered in only one of the three addi-
which replacement E41A was found imposed on the viral pop-
ulations (Table 3). These results indicate that replacement of
hydrophobic residues by charged residues at positions 38 and
41 of 3A drastically reduces viral multiplication, leading to
selection of substitutions to nonpolar V and A residues that
restore hydrophobicity and favor dimer stability.
To study whether the point mutations introduced in 3A could
alter the translation of the viral polyprotein, in vitro-transcribed
RNAs from pMT2-L38E and pMT28L41E were in vitro translated
in a rabbit reticulocytes lysate. The protein pattern obtained for
the different mutants was similar to that of pMT28 control RNA
(data not shown).
the effect of electrostatic charge acquisition at residue Q44 on
FMDV life cycle, the infectivity of mutants with replacements
Q44R and Q44D, which resulted in a slightly lower dimer detec-
tion in transiently expressed 3A mutants (Fig. 2A), was analyzed.
An infectivity of about 197 PFU/ng of RNA was determined for
transcript pMTQ44R, a value slightly higher than that observed
for the control RNA from pMT28 (data not shown). When trans-
fected cells were incubated in liquid medium, RNA from mutant
Q44R produced a cytopathic effect at a time similar to that of
supernatant (Fig. 4), and the viruses recovered maintained the
substitution over 3 additional cell passages. Similar results were
obtained from two additional transfections performed with
pMTQ44R RNA. These results confirm that replacement Q44R
does not exert a detrimental effect on FMDV infectivity in cul-
On the contrary, an infectivity of ?1 PFU/?g was found for
RNA from pMTQ44D, a delay in the recovery of infectious virus
natant of cells transfected were about 1 log lower than those re-
covered from pMT28 RNA. Emergence of infectious virus was
associated with the imposition of replacement D44E, which was
also observed in three additional transfections of RNA from
pMT28-Q44D (Table 3).
On the other hand, in vitro translation of RNAs from
pMT28Q44R and pMT28Q44D showed a protein pattern similar
to that of pMT28 control RNA (data not shown). Thus, while
replacement Q44R leads to viruses with infectivity similar to that
of the parental C-S8c1 virus, introduction of an acidic residue at
to that of the parental Q.
Mutations at residues L38, L41, and Q44 of 3A protein abol-
ish infectivity in suckling mice. To assess the effect of the muta-
model, RNAs transcribed from pMT28L38E, pMT28L41E,
and the emergence of clinical signs and instances of death were
followed for 10 days. RNA from pMT28 was used as a control. An
amount of 100 ng RNA from pMT28 killed 4 of the 5 animals
RNAs derived from full-length infectious clones (7). RNA from
pMT28-Q44R was also lethal for suckling mice, as 100 ng of RNA
region of RNA extracted from dead mice showed the presence of
replacement Q44R, confirming the capacity of the virus carrying
pMT28Q44D, pMT28L38E, and pMT28L41E were not lethal at
ments allow viral replication in cultured cells, enabling selection
model, such as suckling mice.
The self-association of proteins to form dimers and higher-order
TABLE 3 Replacements selected from RNAs with mutations in 3A
Substitution introducedSequence recoveredb
aAmino acid residue and position in 3A are indicated relative to the C-S8c1 sequence.
The corresponding codon is given in parenthesis.
bReplacement selected in the virus recovered upon RNA transfection of BHK-21 cells
(see Materials and Methods for details). In each case, the substitution shown was the
only mutation found in the RNA region sequenced.
cSequence of the viral population recovered from the 3 independent experiments
dSequence of the viral population recovered from 2 of the 4 independent experiments
FIG 4 Effect of 3A mutations on FMDV infectivity. (A) Viral titers of viruses
harboring different 3A mutations recovered after transfection, first or second
passage on BHK-21 cells, were determined at the time of complete CPE, as
follows: 24 hpt for pMT28 and pMT28Q44R or 72 hpt for pMT28L38E,
for all the viruses after the second passage. Data are the averages from three
independent transfections; one out of the three experiments for pMT28-L41E
RNA failed to yield infectious virus.
Determinants of Dimerization on FMDV 3A Protein
October 2012 Volume 86 Number 20 jvi.asm.org 11019
and biophysical studies show that protein dimerization or oli-
gomerization is a key factor in the regulation of different protein
functions (38), including proteins relevant for virus replication
(36). In this line, dimerization/multimerization has been shown
to be relevant for the biological role of nonstructural proteins of
this study, we describe experiments aimed to gain insight on the
tural protein relevant for virus replication, virulence, and host
range for which the molecular mechanisms that mediate its bio-
structural data from E. coli-expressed 3A and peptides corre-
sponding to the N terminus of 3A showed an aggregation ten-
dency that impaired subsequent analyses. In this work, using the
NMR structure of PV 3A as a template, a molecular model for the
N-terminal 94 residues of FMDV 3A was derived. The model
shows that 3A contains two ? helices (?1, residues 25 to 33, and
?2, residues 37 to 44) and that, as in PV and CVB3 3A structures,
a number of hydrophobic contacts in helices ?1 and ?2 could
provide physical stability to the dimer. In addition, in FMDV 3A,
and E36) that conforms a patch of charged residues. In the equiv-
According to the model, the analysis of the transiently ex-
pressed FMDV 3A protein suggested that 3A forms dimers/mul-
timers as shown by Western blotting analyses (Fig. 2A). Protein
oligomerization even in the presence of SDS, has been previously
described for PV 3AB (35, 74). This homodimerization of 3A was
also evidenced by an in situ protein ligation assay designed to
visualize protein-protein interactions in the cell by fluorescence
of 3A to the dimerization was detected using the N-wt synthetic
peptide, spanning residues 1 to 52 of 3A, whose mobility in SDS-
PAGE revealed by mass staining showed a major band of a size
about that corresponding to the monomeric peptide, as well as
bands corresponding to higher-order oligomers (Fig. 2B).
Based on the model and on sequence conservation among
FMDV isolates, hydrophobic interactions between residues at the
helical regions of both monomers were expected to be the main
M29R, involving charge acquisition at residues predicted to con-
tribute to the hydrophobic interface, abolished formation of
dimer bands in transiently expressed 3A (Fig. 2A). Moreover, the
single replacements L38E and L41E and, to a higher extent, the
double replacement L38EL41E showed a reduced fluorescence
3A was evidenced by conventional immunofluorescence with a
different anti-3A antibody (Ab 346). A similar decrease in the
dimerization signal in a double hybrid system has been reported
N terminus of 3A with substitutions L38E, L41E, M29R, or
L38EL41E showed a dramatic reduction of the electrophoretic
forms. Overall, these results suggest that, as reported for CVB3,
is essential for 3A dimer stability.
When FMDV RNA with replacement L38E was transfected in
BHK-21 cells, a delay, relative to cells transfected with parental
C-S8c1 RNA, in both the emergence of cytopathic effect and the
recovery of infectious virus was observed in three independent
experiments. Viruses recovered from the second passage in cul-
tured cells of the transfection medium displayed replacement
E38V that resembled the nonpolar (L) residue of the parental
C-S8c1 virus. On the other hand, infectious virus could be recov-
which again restored the nonpolar nature of this position. The
lower viral titers recovered from transfections with L41E RNA,
relative to those produced by L38E RNA, as well as the lack of
recovery of infectious virus from two of the transfections with
RNA L41E, suggest that this replacement could affect FMDV rep-
lication more severely than L38E. While direct reversion to the
parental residue L at E38 and E44 residues requires at least 2 nu-
cleotide substitutions, single transversions mediate replacements
E38V (A5411T) and E41A (A5420C) (Table 3), making their se-
lection more likely. These results indicate that the presence of
nonpolar hydrophobic residues at positions 38 and 41 of 3A is
essential for virus replication and that no second-site suppressor
mutations at other 3A residues, unlike as reported for coxsacki-
evirus RNA with replacements at analogous 3A hydrophobic res-
RNA mutants L38E and L41E. Thus, taken together, our results
suggest that the conservation of hydrophobic interactions at the
predicted dimerization interface is required for efficient FMDV
replication in cultured cells.
In CVB3, polar interactions, other than the hydrophobic con-
of salt bridges between residues D24 and K41 was found critical
for dimer stability, RNA replication, and inhibition of protein
transport (72). K41 is part of a cluster of charged residues located
at the C terminus of CVB 3A ?-helix 2. A similar cluster of
mutations at these residues (E38, K39, and K40) yield nonviable
viruses, indicating the biological relevance of these polar, charged
residues located at the ?-helix 2 (75). In contrast, the equivalent
a cluster of polar but not charged residues: Q43, Q44, and T45.
According to our model, residues D32 and Q44 of FMDV 3A,
exposed to the solvent, could establish polar interactions between
both monomers. Replacement Q44R, which mediates adaptation
TABLE 4 RNA infectivity in suckling mice of FMDV 3A mutants
RNARNA dose (ng)
No. of dead mice/total no. of mice
González-Magaldi et al.
jvi.asm.org Journal of Virology
salt bridge with D32 with the consequent dimer stabilization.
Conversely, replacement Q44D would impair polar interactions
produced a marked effect on the dimer formation of transiently
expressed 3A, suggesting that the polar interactions between Q44
and D32 are not critical for 3A dimerization, albeit they could
modulate dimer stability. A similar observation was reported for
CVB3 3A in which replacements to A at polar residues S28 and
Y37, predicted to establish an intermolecular hydrogen bond, did
not affect dimerization and protein transport, while only replace-
ment Y37A impaired virus replication, leading to recovery of a
second-site suppressor mutation (72).
When the effect of electrostatic charge acquisition at position
44 on the infectiveness of FMDV RNA was analyzed, transfected
RNA with replacement Q44R produced a cytopathic effect and
infectious virus in a manner similar to that of the parental RNA
pMT28, and the viruses recovered maintained the substitution
over 3 additional cell passages, indicating that this change is not
detrimental for virus replication in cultured cells. This result is
in the pig (47). On the contrary, in cells transfected with RNA
carrying the replacement Q44D, a delay in the production of in-
fectious virus was observed and the recovered virus displayed re-
placement D44E. In this case, substitution of D by E did not re-
charge and structural features similar to those of the parental Q.
Direct reversion to the parental residue Q requires 2 nucleotide
substitutions, while a single transversion mediates replacement
D44E (T5430A) (Table 3). Residue Q44 is rather conserved
among FMDVs, and replacements at these residue have been
despite the fact that replacements at Q44 do not substantially af-
fect the capacity of transiently expressed 3A to form dimers, this
residue is relevant for virus replication. Thus, while replacement
the parental C-S8c1, Q44D leads to the recovery of the D44E mu-
Despite the fact that replication impairment introduced by
mutants able to grow in cultured cells, RNAs carrying these sub-
stitutions caused no death or clinical signs when inoculated in
suckling mice, opposite to what was observed for RNA from
C-S8c1 (pMT28) and from mutant Q44R. These results indicate
that, as previously reported (7), in vivo multiplication frequently
imposes different constrains for virus replication and disease
emergence compared to those found in cultured cells.
dimerization inhibition on the detrimental effect of the mutants
of FMDV 3A protein among picornaviruses, FMDV requires 3A
dimerization for efficient replication. Our data support that the
hydrophobic interactions established between the ?-helices of
both monomers are the main determinant for dimerization and
its impairment is detrimental for virus multiplication. On the
other hand, mutations affecting polar interactions between resi-
dues at the ?-helices can affect FMDV replication, without abol-
ishing 3A dimerization.
We thank E. Domingo and C. Escarmís for infectious clone pMT28. We
are indebted to B. Borrego for advice and help with animal experiments
ing of the manuscript.
Work at the F.S. laboratory was supported by grants from Ministerio
de Ciencia e Innovación (MICINN, Spain): BIO2008-0447-C03-01,
CSD2006-0007, and BIO2011-24351. Work at the D.A. laboratory was
supported by MICINN grants BIO2008-0447-C03-01 and SAF2011-
24899 and by Generalitat de Catalunya (SGR2009-0492). Work at the
P.G.-P. laboratory was supported by MICINN grants SAF2007-61926,
IPT2011-0964-900000, and SAF2011-13156-E and by the European
Commission through grants FP7 HEALTH-F3-2009-223431 (EU project
“Divinocell”) and FP7 HEALTH-2011-278603 (EU project “Dorian”).
Support from the Fundación Ramón Areces and the Centro de Com-
edged. Work at Biomol-Informatics was partially financed by the Euro-
grants PI 201120E007 and PI 201120E089 and by MSC grant FIS 2010-
1. Andino R, Boddeker N, Silvera D, Gamarnik AV. 1999. Intracellular
determinants of picornavirus replication. Trends Microbiol. 7:76–82.
2. Andino R, Rieckhof GE, Achacoso PL, Baltimore D. 1993. Poliovirus
RNA. EMBO J. 12:3587–3598.
receptors in the form of homotrimers. Br. J. Pharmacol. 163:1069–1077.
4. Armas-Portela R, Parrales MA, Albar JP, Martinez AC, Avila J. 1999.
Distribution and characteristics of betaII tubulin-enriched microtubules
in interphase cells. Exp. Cell Res. 248:372–380.
biology and immunogenicity, p 3–32. In Romberger JA (ed), Beltsville
Osmund, Monclair, NJ.
6. Bagos PG, Liakopoulos TD, Hamodrakas SJ. 2006. Algorithms for in-
membrane proteins. BMC Bioinformatics 7:189.
7. Baranowski E, Molina N, Nunez JI, Sobrino F, Saiz M. 2003. Recovery
inoculation with in vitro-transcribed RNA. J. Virol. 77:11290–11295.
8. Beard CW, Mason PW. 2000. Genetic determinants of altered virulence
of Taiwanese foot-and-mouth disease virus. J. Virol. 74:987–991.
Microbiol. Immunol. 288:43–70.
10. Benkert R, Templin T, Schim SM, Doorenbos AZ, Bell SE. 2011. Testing
a multigroup model of culturally competent behaviors among underrep-
resented nurse practitioners. Res. Nurs. Health 34:327–341.
11. Bienz K, Egger D, Pasamontes L. 1987. Association of polioviral proteins
of the P2 genomic region with the viral replication complex and virus-
nocytochemistry and autoradiography. Virology 160:220–226.
12. Bienz K, Egger D, Rasser Y, Bossart W. 1983. Intracellular distribution
of poliovirus proteins and the induction of virus-specific cytoplasmic
structures. Virology 131:39–48.
microgram quantities of protein utilizing the principle of protein-dye
binding. Anal. Biochem. 72:248–254.
14. Buenz EJ, Howe CL. 2006. Picornaviruses and cell death. Trends Micro-
15. Cameron CE, Suk Oh H, Moustafa IM. 2010. Expanding knowledge of
P3 proteins in the poliovirus lifecycle. Future Microbiol. 5:867–881.
16. Carrillo C, et al. 2005. Comparative genomics of foot-and-mouth disease
virus. J. Virol. 79:6487–6504.
17. Cuconati A, Xiang W, Lahser F, Pfister T, Wimmer E. 1998. A protein
linkage map of the P2 nonstructural proteins of poliovirus. J. Virol. 72:
18. Cho MW, Teterina N, Egger D, Bienz K, Ehrenfeld E. 1994. Membrane
Determinants of Dimerization on FMDV 3A Protein
October 2012 Volume 86 Number 20 jvi.asm.org 11021
rearrangement and vesicle induction by recombinant poliovirus 2C and
2BC in human cells. Virology 202:129–145.
19. Choe SS, Kirkegaard K. 2004. Intracellular topology and epitope shield-
ing of poliovirus 3A protein. J. Virol. 78:5973–5982.
20. Datta U, Dasgupta A. 1994. Expression and subcellular localization of
poliovirus VPg-precursor protein 3AB in eukaryotic cells: evidence for
glycosylation in vitro. J. Virol. 68:4468–4477.
21. De Diego M, Brocchi E, Mackay D, De Simone F. 1997. The non-
structural polyprotein 3ABC of foot-and-mouth disease virus as a diag-
nostic antigen in ELISA to differentiate infected from vaccinated cattle.
Arch. Virol. 142:2021–2033.
22. Deitz SB, Dodd DA, Cooper S, Parham P, Kirkegaard K. 2000. MHC
I-dependent antigen presentation is inhibited by poliovirus protein 3A.
Proc. Natl. Acad. Sci. U. S. A. 97:13790–13795.
23. Doedens JR, Giddings TH, Jr, Kirkegaard K. 1997. Inhibition of endo-
plasmic reticulum-to-Golgi traffic by poliovirus protein 3A: genetic and
ultrastructural analysis. J. Virol. 71:9054–9064.
24. Domingo E, et al. 1990. Genetic variability and antigenic diversity of
tel MHV (ed), Applied virology research, vol 2. Plenum Publishing Cor-
poration, New York, NY.
25. Egger D, Teterina N, Ehrenfeld E, Bienz K. 2000. Formation of the
poliovirus replication complex requires coupled viral translation, vesicle
production, and viral RNA synthesis. J. Virol. 74:6570–6580.
26. Falk MM, et al. 1990. Foot-and-mouth disease virus protease 3C induces
specific proteolytic cleavage of host cell histone H3. J. Virol. 64:748–756.
27. Fujita K, et al. 2007. Membrane topography of the hydrophobic anchor
sequence of poliovirus 3A and 3AB proteins and the functional effect of
3A/3AB membrane association upon RNA replication. Biochemistry 46:
28. Gajadhar A, Guha A. 2009. A proximity ligation assay using transiently
transfected, epitope-tagged proteins: application for in situ detection of
dimerized receptor tyrosine kinases. Biotechniques 48:145–152.
29. García-Arriaza J, Manrubia SC, Toja M, Domingo E, Escarmís C. 2004.
plementation. J. Virol. 78:11678–11685.
30. Garcia-Briones M, et al. 2006. Differential distribution of non-structural
proteins of foot-and-mouth disease virus in BHK-21 cells. Virology 349:
Biochem. Sci. 24:364–367.
32. Hope DA, Diamond SE, Kirkegaard K. 1997. Genetic dissection of
interaction between poliovirus 3D polymerase and viral protein 3AB. J.
33. Kall L, Krogh A, Sonnhammer EL. 2007. Advantages of combined trans-
membrane topology and signal peptide prediction—the Phobius Web
server. Nucleic Acids Res. 35:W429–W432.
34. Knowles NJ, Samuel AR, Davies PR, Kitching RP, Donaldson AI. 2001.
a pandemic strain. Vet. Rec. 148:258–259.
35. Lama J, Paul AV, Harris KS, Wimmer E. 1994. Properties of purified
recombinant poliovirus protein 3aB as substrate for viral proteinases and
as co-factor for RNA polymerase 3Dpol. J. Biol. Chem. 269:66–70.
36. Li S, et al. 2009. Dimerization of hepatitis E virus capsid protein E2s
domain is essential for virus-host interaction. PLoS Pathog. 5:e1000537.
37. Lyle JM, et al. 2002. Similar structural basis for membrane localization
and protein priming by an RNA-dependent RNA polymerase. J. Biol.
38. Marianayagam NJ, Sunde M, Matthews JM. 2004. The power of two:
protein dimerization in biology. Trends Biochem. Sci. 29:618–625.
39. Martin-Acebes MA, et al. 2008. Subcellular distribution of swine vesicu-
lar disease virus proteins and alterations induced in infected cells: a com-
virus. Virology 374:432–443.
40. Martin-Acebes MA, Rincon V, Armas-Portela R, Mateu MG, Sobrino F.
2010. A single amino acid substitution in the capsid of foot-and-mouth
disease virus can increase acid lability and confer resistance to acid-
dependent uncoating inhibition. J. Virol. 84:2902–2912.
41. Martin-Acebes MA, Vazquez-Calvo A, Rincon V, Mateu MG, Sobrino
disease virus can increase acid resistance. J. Virol. 85:2733–2740.
42. Moffat K, et al. 2005. Effects of foot-and-mouth disease virus nonstruc-
2BC but not 3A blocks endoplasmic reticulum-to-Golgi transport. J. Vi-
43. Monaghan P, Cook H, Jackson T, Ryan M, Wileman T. 2004. The
ultrastructure of the developing replication site in foot-and-mouth dis-
ease virus-infected BHK-38 cells. J. Gen. Virol. 85:933–946.
44. Nayak A, Goodfellow IG, Belsham GJ. 2005. Factors required for the
tides by the RNA-dependent RNA polymerase (3Dpol) in vitro. J. Virol.
proximity ligation on angiogenic sprouts. EMBO J. 29:1377–1388.
46. Notredame C, Higgins DG, Heringa J. 2000. T-Coffee: a novel method
for fast and accurate multiple sequence alignment. J. Mol. Biol. 302:205–
47. Nunez JI, et al. 2007. Guinea pig-adapted foot-and-mouth disease virus
with altered receptor recognition can productively infect a natural host. J.
48. Nuñez JI, et al. 2001. A single amino acid substitution in nonstructural
guinea pig. J. Virol. 75:3977–3983.
49. O’Donnell VK, Pacheco JM, Henry TM, Mason PW. 2001. Subcellular
distribution of the foot-and-mouth disease virus 3A protein in cells in-
fected with viruses encoding wild-type and bovine-attenuated forms of
3A. Virology 287:151–162.
50. Pacheco JM, Henry TM, O’Donnell VK, Gregory JB, Mason PW. 2003.
Role of nonstructural proteins 3A and 3B in host range and pathogenicity
of foot-and-mouth disease virus. J. Virol. 77:13017–13027.
51. Peitsch MC. 1996. ProMod and Swiss-Model: Internet-based tools for
Virus diseases of food animals. Academic Press Inc., London, United
53. Rath A, Glibowicka M, Nadeau VG, Chen G, Deber CM. 2009. Deter-
gent binding explains anomalous SDS-PAGE migration of membrane
proteins. Proc. Natl. Acad. Sci. U. S. A. 106:1760–1765.
54. Rosas MF, et al. 2008. Susceptibility to viral infection is enhanced by
stable expression of 3A or 3AB proteins from foot-and-mouth disease
virus. Virology 380:34–45.
55. Ryan MD, et al. 2004. Foot-and-mouth disease virus proteinases. In
Horizon Bioscience, Norfolk, United Kingdom.
56. Saiz M, Gomez S, Martinez-Salas E, Sobrino F. 2001. Deletion or
substitution of the aphthovirus 3= NCR abrogates infectivity and virus
replication. J. Gen. Virol. 82:93–101.
57. Samuilova O, Krogerus C, Poyry T, Hyypia T. 2004. Specific interaction
between human parechovirus nonstructural 2A protein and viral RNA. J.
Biol. Chem. 279:37822–37831.
58. Sanz-Parra A, Sobrino F, Ley V. 1998. Infection with foot-and-mouth
J. Gen. Virol. 79(Pt 3):433–436.
59. Schwede T, Kopp J, Guex N, Peitsch MC. 2003. SWISS-MODEL: an
automated protein homology-modeling server. Nucleic Acids Res. 31:
60. Semler BL, Anderson CW, Hanecak R, Dorner LF, Wimmer E. 1982. A
precipitation with antibodies directed against a synthetic heptapeptide.
61. Sobrino F, Dávila M, Ortín J, Domingo E. 1983. Multiple genetic
variants arise in the course of replication of foot-and-mouth disease virus
in cell culture. Virology 128:310–318.
but a current threat. Vet. Res. 32:1–30.
63. Soderberg O, et al. 2006. Direct observation of individual endogenous
protein complexes in situ by proximity ligation. Nat. Methods 3:995–
64. Sonnhammer EL, Hollich V. 2005. Scoredist: a simple and robust protein
sequence distance estimator. BMC Bioinformatics 6:108.
65. Strauss DM, Glustrom LW, Wuttke DS. 2003. Towards an understand-
ing of the poliovirus replication complex: the solution structure of the
soluble domain of the poliovirus 3A protein. J. Mol. Biol. 330:225–234.
66. Suhy DA, Giddings TH, Jr, Kirkegaard K. 2000. Remodeling the endo-
González-Magaldi et al.
jvi.asm.orgJournal of Virology
plasmicreticulumbypoliovirusinfectionandbyindividualviralproteins: Download full-text
an autophagy-like origin for virus-induced vesicles. J. Virol. 74:8953–
67. Sweeney TR, et al. 2010. Foot-and-mouth disease virus 2C is a hexameric
AAA? protein with a coordinated ATP hydrolysis mechanism. J. Biol.
68. Takegami T, Kuhn RJ, Anderson CW, Wimmer E. 1983. Membrane-
dependent uridylylation of the genome-linked protein VPg of poliovirus.
Proc. Natl. Acad. Sci. U. S. A. 80:7447–7451.
69. Towner JS, Ho TV, Semler BL. 1996. Determinants of membrane asso-
ciation for poliovirus protein 3AB. J. Biol. Chem. 271:26810–26818.
70. Tusnady GE, Simon I. 2001. Topology of membrane proteins. J. Chem.
Infect. Comput. Sci. 41:364–368.
proteomics toolbox. Expert Rev. Proteomics 7:401–409.
72. Wessels E, et al. 2006. Structure-function analysis of the coxsackievirus
protein 3A: identification of residues important for dimerization, viral
RNA replication, and transport inhibition. J. Biol. Chem. 281:28232–
73. Whitton JL, Cornell CT, Feuer R. 2005. Host and virus determinants of
picornavirus pathogenesis and tropism. Nat. Rev. Microbiol. 3:765–776.
74. Xiang W, Cuconati A, Hope D, Kirkegaard K, Wimmer E. 1998.
Complete protein linkage map of poliovirus P3 proteins: interaction of
polymerase 3Dpol with VPg and with genetic variants of 3AB. J. Virol.
75. Xiang W, Cuconati A, Paul AV, Cao X, Wimmer E. 1995. Molecular
dissection of the multifunctional poliovirus RNA-binding protein 3AB.
76. Xiang W, Harris KS, Alexander L, Wimmer E. 1995. Interaction between
the 5=-terminal cloverleaf and 3AB/3CDpro of poliovirus is essential for
RNA replication. J. Virol. 69:3658–3667.
77. Zell R, Seitz S, Henke A, Munder T, Wutzler P. 2005. Linkage map of
protein-protein interactions of porcine teschovirus. J. Gen. Virol. 86:
Determinants of Dimerization on FMDV 3A Protein
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