A DNA-launched reverse genetics system for rabbit
hemorrhagic disease virus reveals that the VP2
protein is not essential for virus infectivity
Guangqing Liu, Zheng Ni, Tao Yun, Bin Yu, Liu Chen, Wei Zhao,
Jionggang Hua and Jianping Chen
Institute of Virology and Biotechnology, Zhejiang Academy of Agricultural Sciences, Hangzhou
310021, PR China
Received 25 April 2008
Accepted 30 July 2008
Rabbit hemorrhagic disease virus (RHDV), a member of the family Caliciviridae comprising
positive-stranded RNA viruses, is a highly virulent pathogen of rabbits. Until recently, studies into
the molecular mechanisms of RHDV replication and pathogenesis have been hindered by the lack
of an in vitro culture system and reverse genetics. This study describes the generation of a DNA-
based reverse genetics system for RHDV and the subsequent investigation of the biological role
of the RHDV VP2 protein. The full-length RHDV genome was assembled as a single cDNA clone
and placed under the control of the eukaryotic human cytomegalovirus promoter. Transfection of
cells with the DNA clone resulted in a clear cytopathic effect and the generation of infectious
progeny virions. The reconstituted virus was stable and grew to titres similar to that of the parental
virus. Although previous reports have suggested that the minor structural protein (VP2) of other
caliciviruses is essential for the production of infectious virions, using the DNA-launch-based
RHDV reverse genetics system described here it was demonstrated that VP2 is not essential for
RHDV infectivity. Transfection of cells with a cDNA clone of RHDV lacking VP2 resulted in the
generation of infectious virions. These studies indicate that the presence of VP2 could reduce the
replication of RHDV, suggesting that it may play a regulatory role in the life cycle of RHDV.
Rabbit hemorrhagic disease virus (RHDV) is an emerging
disease in rabbits and is considered the single most
economically important disease of rabbits worldwide. The
disease was first recognized in China (Liu et al., 1984), but
was subsequently isolated from other areas of Asia, various
et al.,1991; Nowotny et al., 1997). In 1995, the virus
reached mainland Australia after escaping from an island
where it had been kept for experimental purposes (Meyers
et al., 1991). Two years later, rabbit haemorrhagic disease
was also observed in New Zealand (Ohlinger et al., 1990).
The aetiological agent was identified as a calicivirus (Mitro
& Krauss, 1993), a positive-sense, single-stranded RNA
virus that is antigenically related to European brown hare
syndrome virus (Wirblich et al., 1994; Mutze et al., 1998).
The complete genome of the virus has been elucidated for
the German isolate (Meyers et al., 1991) and shown to
comprise a genome of 7437 nt. The genome contains two
open reading frames, the first of which, ORF1, contains
2344 codons and encodes a large polyprotein containing
the viral non-structural proteins as well as the viral coat
protein at the C terminus. The genome also has a virus-
encoded protein, VPg, attached covalently to the 59 end
(Gregg et al., 1991) and is polyadenylated at the 39 end
(Morales et al., 2004).
In RHDV-infected cells, a 2.2 kb subgenomic mRNA is
transcribed that is collinear with the 39 third of the
genomic RNA (Meyers et al., 1991). This mRNA is thought
to represent the major source of the RHDV capsid protein,
VP60; however, VP60 is also generated via cleavage of the
ORF1-encoded polyprotein (Parra et al., 1993; Wirblich
et al., 1995; Boga et al., 1999). A second ORF, ORF2, is
located at the extreme 39 end of the genomic and
subgenomic RNAs. The start codon for ORF2 is located
at nt 7025 and shares a 17 nt overlap with ORF1, but has a
21 frame shift relative to the capsid ORF. RHDV ORF2 is
117 aa and encodes a polypeptide of 12.7 kDa (VP2),
which is considered a component of RHDV virions
(Wirblich et al. 1996; Meyers et al., 2000; Meyers, 2003).
VP2 is conserved throughout the caliciviruses, suggesting
that it may play a role in virus replication or assembly.
However, the precise biological function of RHDV VP2 is
not clear, and the inability to grow RHDV together with a
Journal of General Virology (2008), 89, 3080–3085
30802008/003525G2008 SGMPrinted in Great Britain
lack of reagents to detect ORF2 has hampered studies of
the VP2 protein. Studies on other members of the
calicivirus family have demonstrated that the minor capsid
protein, VP2, is critical for infectivity, as interruption of
the feline calicivirus (FCV) VP2 ORF results in a loss of
infectivity (Sosnovtsev et al., 2005).
Recently, an infectious cDNA clone of RHDV was con-
structed. The rescued virus is adapted to growth in rabbit
RK13 cells, providing a useful platform for the study of
RHDV (Liu et al., 2006). In the present paper, we describe a
DNA-launched reverse genetics system for RHDV and the
application of this system to investigate the biological
relevance of VP2. The entire ORF2 of RHDV was deleted
to generate a cDNA clone lacking ORF2. The role of the VP2
protein for virus infectivity was then investigated in vivo
using the reconstructed genomic clone. Our results indicated
that VP2 is not essential for the infectivity of RHDV.
Cells, virus antibody and plasmids. The rabbit kidney cell line
RK13 was obtained from the China Center for Type Culture
Collection. Construction of an infectious cDNA clone was based on
the CHA/JX/97 strain of RHDV. CHA/JX/97 is a virulent strain of
RHDV isolated in 1997 from an outbreak of rabbit haemorrhagic
disease in Jiaxing, China. Monoclonal antibodies (mAbs) specific for
VP60 and VP10 were prepared in our laboratory. Plasmid pcDNA
3.1(+) (Invitrogen) was used for the assembly of a full-length cDNA
clone of RHDV. The core sequence of self-cleaving ribozyme of the
hepatitis delta virus (HDV) was kindly provided by Haixue Zheng
(Lanzhou Veterinary Institute, Gansu, China).
Mutagenesis and reconstruction of pcDNA3.1. To ensure that the
transcripts had a precise 59-terminal sequence, 2 nt downstream of the
human cytomegalovirus (HCMV) promoter were mutated (A833AG
and A835AT) using a QuikChange Site-directed Mutagenesis kit
according to the manufacturer’s protocol (Stratagene) to introduce a
KpnI site near the putative transcriptional start site of pcDNA3.1.
Subsequently, pcDNA3.1 was digested with KpnI and an 80 bp DNA
fragment was released. Finally, the core sequence of self-cleaving
ribozyme of HDV was subcloned into the above plasmid and the
resulting plasmid was designated pCMV-HDV.
Assembly of a full-length RHDV cDNA clone. A full-length cDNA
clone of CHA/JX/97 was assembled by following a multistep strategy.
First, the pCMV-HDV plasmid was digested with KpnI and EcoRV to
allow insertion of the CD fragment from the RHDV genome (nt 1–
2909), previously amplified from pRHDV using the specific primers
F1+(59-ACTGGTACCGTGAAAATTATGCCGCCTATG-39; KpnI site
underlined) and F12(59-GTTGACAAGGTGGTTCGCACACAG-39).
This yielded a recombinant plasmid, pCMV-CD, containing nt 1–2909
from the RHDV genome. The remainder of the RHDV genome
(fragment AB) was subsequently amplified from pRHDV with another
pair of primers, F2+(59-CCATCGATGATATCACACCTGTGCGCA-
AA-39) and F22(59-CATGGGCCCTCTAGAGCGGCCGCTTTTTT-
TTTTTTTTTT-39; ApaI and NotI sites underlined, respectively, and
XbaI site in italics). After digestion with EcoRV and NotI, the DNA
fragment was inserted into pCMV-CD, which had been cleaved by the
same enzymes. The recombinant plasmid was named pRHDV.
Generation of a mutant RHDV cDNA clone without the VP2-
coding region. To delete the VP2-coding region (ORF2) from the
RHDV genome, two NarI cleavage sites were introduced on either
side of ORF2. The first mutation was introduced at nt 7013–7018,
where 2 nt were mutated (G7015AC and A7018AC) to generate a NarI
site. The second NarI site was introduced at nt 7361–7366 by two
point mutations (7365AG7366ACC). Four specific primers were used
during the mutagenesis: NarI7013+(sense primer: 59-GTTCAACC-
NarI7361+(sense primer: 59-TACCACTGGCGCCTCCAGTGA-39)
and NarI73612(antisense primer: 59-TCACTGGAGGCGCCAGTG-
GTA-39). The mutations were carried out using a QuikChange Site-
directed Mutagenesis kit. Subsequently, the mutated recombinant
plasmid was digested with NarI and the VP2-coding region was
released. The resulting plasmid was designated pRHDVDVP2 .
Transfection of RK13 cells. RK13 cells were grown to 80%
confluency and transfected for 4 h at 37 uC with a mixture of 2–5 mg
pRHDV or pRHDVDVP2 and 10 ml Lipofectmine 2000 (Invitrogen),
according the supplier’s instructions.
RT-PCR for RHDV genome and sequencing. RNA from the
recombinant and parental viruses was purified using an RNeasy
Extraction kit (Qiagen). A 443 bp fragment that included the genetic
marker was then amplified by RT-PCR using the primers JD+(59-
CCAACTGCACAATTCAAATCC-39) and JD2(59-TGAACATGACG-
GAGTCCTGGT-39). The RT-PCR products were digested with
EcoRV and analysed on a 1.5% agarose gel.
Immunofluorescence assay (IFA). Indirect IFAs were used to
detect viral protein expression in RK13 cells. Cells were fixed in 3.7%
paraformaldehyde in PBS (pH 7.5) at room temperature for 30 min
and permeabilized by incubation in 220 uC methanol for 30 min.
The fixed cells were washed with PBS and stained with a mAb specific
for VP60 (1:500 dilution), followed by goat anti-mouse immuno-
globulin G conjugated to fluorescein isothiocyanate. Finally, the
samples were observed under a fluorescence microscope equipped
with a video documentation system.
Detection of VP2 expression. Lysates were analysed by Western
blot analysis using a mouse hyperimmune serum to VP2 after
separation by SDS-PAGE and transfer to nitrocellulose membranes
(Hybond-C; Amersham Life Sciences).
Virus titration. To compare the growth kinetics of the mutant and
parental virus, RK13 cells were cultured in 96-well plates and then
infected at an m.o.i. of 0.003 TCID50per cell with stocks of virus
generated from four passages on RK13 cells. After 2 h of incubation,
the cells were washed twice and fresh growth medium was added
(time 0). The cells were incubated at 37 uC in a humidified 5% CO2
atmosphere and observed daily for the appearance of cytopathic effect
(CPE). From the onset of CPE, the titres of rescued viruses were
determined as TCID50values at 24, 36, 48, 60 and 72 h post-infection.
Quantification of RHDV genome RNA levels. A comparative
analysis of the increase in genome copies of recombinant RHDV
between virus recovered from pRHDV and from pRHDVDVP2 was
carried out. Two pairs of PCR primers were designed. The first pair,
F1 (59-AGGACAAAACGAGAATGAAGGA-39) and R1 (59-GCTGG-
GCTATGGAACACAAAC-39), were used to quantify the RHDV
genomic RNA levels, whilst the second pair, F+(59-TCGGGTT-
TGGTGGTATTTGG-39) and R2(59-GGTGGGCTGGAGGTTGTTT-
39), were used to detect RHDV subgenomic RNA levels. Viral RNA
was extracted from lysates of infected cells using an RNeasy Mini kit
and reverse-transcribed into cDNA using the specific primers.
Subsequently, the cDNA was amplified by real-time PCR using a
SYBR Green PCR mix (TaKaRa). PCR amplification was performed
using a program of 10 min at 95 uC followed by 40 cycles of 1 min at
A reverse genetics system of RHDV
94 uC, 1 min at 60 uC and 1 min at 72 uC. Each reaction was
performed in triplicate in a Perkin Elmer ABI Prism 7700 Sequence
Detection system (TaKaRa). Standards to establish genome equiva-
lents were synthetic RNAs transcribed from a clone of the full-length
cDNA of RHDV. The RNA was quantified by absorbance and 10-fold
serial dilutions were prepared from 106to 101copies.
Construction of a full-length RHDV cDNA clone
and ORF2 knockout
cDNA fragments of RHDV were assembled into a single
full-length clone from two overlapping fragments using the
restriction sites NotI, EcoRV and KpnI derived from the
plasmid. The full-length genomic cDNA was placed under
the control of the HCMV promoter of pcDNA3.1,
engineered to eliminate the addition of non-viral nucleo-
tides at the 59 extremities of viral transcripts. To ensure
that the final transcripts generated had the correct 39 end,
the core sequence of HDV ribozyme was placed immedi-
ately downstream of the viral poly(A) tract. Thus, the RNA
transcripts generated from the plasmid contained a 16 nt
poly(A) tract followed by an additional GC. The final
plasmid was designated pRHDV. To knock out the ORF2
from the full-length genome, two NarI sites were
introduced on either side of ORF2 in pRHDV. In the
newly derived plasmid, the ORF1 sequence and 39-end
non-translated region were bordered by NarI restriction
sites and ORF2 was deleted from pRHDV by NarI
digestion and subsequent religation. The resulting plasmid
was designated pRHDVDVP2.
Infectivity of cDNA clones in cell culture
Transfection of RK13 cells with pRHDV resulted in the
appearance of CPE at 12 h post-transfection, which
became more prominent at 48 h post-transfection. The
generation of viral RNA and antigen was subsequently
confirmed by RT-PCR followed by sequencing, as well as
by IFA using a VP60-specific mAb. At 2 days post-
transfection, VP60, the major capsid protein, was clearly
detected in the cytoplasm of cells transfected with pRHDV,
whereas untransfected cells failed to cross-react with the
VP60 antiserum (Fig. 1).
To rule out the possibility of contamination with the
parental virus, RT-PCR and sequencing were performed
for virus recovered from DNA transfection. The recon-
stituted virus derived from pRHDV retained the genetic
marker (EcoRV site) located at nt 2907, as indicated by the
ability of EcoRV to digest the RT-PCR-amplified region.
These results confirmed that the infectivity in transfected
cells was derived from the DNA construct.
No VP2 expression in the mutant virus
The results of Western blot analysis demonstrated that VP2
was expressed by the virus rescued using either pRHDV or
in vitro-synthesized transcripts (Fig. 2). As expected, the
VP2 protein was not detected in cells infected with the
virus rescued from pRHDVDVP2 (Fig. 2), indicating that
the recombinant virus lacked VP2.
Infectivity of pRHDVDVP2
The mutant cDNA clone containing a deletion of the entire
VP2-coding region was transfected into RK13 cells and the
appearance of CPE was monitored. At 12 h post-transfec-
tion, visible CPE was observed, and when stained with a
VP60-specific mAb, cells showed bright fluorescence,
indicating the presence of the VP60 protein in the
cytoplasm (Fig. 1). In contrast, transfection of pcDNA3.1
did not produce any visible CPE for up to 72 h post-
transfection, suggesting that VP60 can be expressed in the
absence of VP2.
Fig. 1. IFA of viral protein expression in cells transfected with
pRHDV and pRHDVDVP2. (a) Normal RK13 cells; (b) RK13 cells
transfected with pRHDVDVP2; (c) RK13 cells transfected with
G. Liu and others
3082 Journal of General Virology 89
To examine further the infectivity of pRHDVDVP2, RT-
PCR was conducted from passage 3 viruses and a specific
fragment (nt 2729–3171) was RT-PCR amplified. As
anticipated, a 443 bp product was amplified from the
culture cells (data not shown), and sequencing results
confirmed the authenticity of the amplified product. These
results confirmed that the mutant virus was infectious in
the absence of VP2.
The growth curve of the mutant virus was analysed by
determination of TCID50values in comparison with the
wild-type parental virus. The results showed that the titres
of the rescued virus from pRHDVDVP2 were slightly lower
than the titres obtained for the parental virus, but both
viruses reached a similar maximum titre of 16104.32and
16104.68TCID50ml21by 3 days post-infection, respect-
ively (Fig. 3). These data indicated that, unlike previous
observations with FCV, deletion of RHDV VP2 did not
result in complete loss of RHDV replication ability,
although there was a slight decrease in infectivity.
Quantification of genome replication by
To evaluate the effect of VP2 deletion on RHDV
replication, qRT-PCR was used to quantify genomic and
subgenomic RNA levels. A pair of primers (F1/R1) located
in the coding region of the non-structural protein p16 was
used to detect replication of RHDV genomic RNA, whilst a
pair of primers (F+/R2) targeting the VP60-coding region
was used to detect levels of subgenomic RNA. The results
demonstrated that the level of the viral subgenomic RNA
was decreased up to 60 h post-infection as result of VP2
deletion (Fig. 4a), although it showed an increase at 72 h
post-infection. The replication level of the VP2 deletion
mutant was reproducibly higher than that of the parental
virus (Fig. 4b).
Fig. 2. Western blot analysis of VP2 expression. Western blot of
cell lysates of RK13 cells transfected with infectious in vitro-
transcribed RHDV RNA (lane 1), pRHDV (lane 2) or pRHDVDVP2
(lane 3) stained with RHDV VP2-specific antiserum. M, Marker.
Fig. 3. Multi-step growth curves of recombinant viruses recovered
from pRHDV and pRHDVDVP2. RK13 cells were infected with
virus at an m.o.i. of 0.003 TCID50per cell. At various time points,
cells were harvested and the titres of the viruses determined as
Fig. 4. qRT-PCR analysis of genome replication. qRT-PCR was
used to evaluate the levels of subgenomic (a) and genomic (b)
RNA levels of viruses derived from pRHDV and pRHDVDVP2.
Quantification was performed in triplicate and the mean values of
genome equivalents, as determined by comparison with a standard
curve of in vitro-transcribed RNA, were plotted. The experiment
was performed three times and the data from one experiment is
A reverse genetics system of RHDV
It is well known that the use of reverse genetics systems is a
useful tool for studying RNA virus replication, pathogen-
esis and in vivo function of individual viral proteins, as well
as for developing new vaccines. For caliciviruses, several
infectious cDNA clones have been reported for FCV
(Thumfart & Meyers, 2002; Sosnovtsev et al., 2005),
RHDV (Liu et al., 2006), murine norovirus (Chaudhry et
al., 2007) and porcine enteric calicivirus (Chang et al.,
2005). These systems have the common feature that the
full-length cDNA molecule is placed downstream of the T7
or SP6 promoter, and the entire genomic RNA is either
synthesized in vitro and the transcripts introduced into
host cells in order to recover infectious virus or the RNA is
synthesized in vivo by the use of a T7 RNA polymerase-
expressing helper virus. In the present study, we placed the
full-length cDNA of RHDV under the control of the
HCMV immediate-early eukaryotic promoter to generate
genomic RNA after transfection into cells and rescue
infectious RHDV. To ensure the authenticity of the 59 end
of the virus genome derived from the HCMV promoter,
the nucleotides between the HCMV promoter and genome
start site were deleted, so that transcription by RNA
polymerase II began at or very near the 59 end of the viral
genome. Similarly, to ensure a precise 39 end of the viral
genome, a DNA fragment containing the core sequence of
the self-cleaving ribozyme of HDV was inserted between
the 39 end of the genome and the bovine growth hormone
poly(A) sequence. In the reverse genetics system described
here, plasmid DNA was used for direct transfection of
RHDV-susceptible cells, which makes the manipulation of
the RHDV genome technically easier and more consistent
than RNA transfection.
All caliciviruses are known to encode the VP2 protein in a
separate ORF near the 39 end of the genome (ORF2 for
sapoviruses and lagoviruses and ORF3 for noroviruses and
vesiviruses). Although the VP2 protein was proposed to be
a minor structural protein, the biological function of VP2
remains to be fully elucidated. In 2003, the function of
Norwalk virus VP2 was studied by the Estes group
(Bertolotti-Ciarlet et al., 2003; Glass et al., 2003); their
results showed that the protein could interact with capsid
protein and regulate the expression and stability of the viral
capsid protein VP1. Two years later, Sosnovtsev et al.
(2005) showed that FCV VP2 was essential for the
production of infectious virions, as deletion of VP2
resulted in complete loss of infectivity. In addition,
Kaiser et al. (2006) reported that the FCV VP2 protein
can interact with capsid protein and possibly the RNA
polymerase in a yeast two-hybrid system. For RHDV, the
function of VP2 is still unknown. In the present study, we
utilized a DNA-launch-based RHDV reverse genetics
system to investigate the role of VP2 in viral replication.
We demonstrated that the VP2 protein is not essential for
the production of infectious RHDV virions, which is in
contrast to previous suggestions that VP2 is involved in the
maturation and assembly of calicivirus particles (Wirblich
et al., 1995; Glass et al., 2000; Sosnovtsev & Green, 2000;
Oehmig et al., 2003).
We have systematically studied the effects of the VP2
deletion on viral replication, capsid expression and RHDV
infectivity. The results clearly demonstrated that RHDV
retains its infectivity in the absence of VP2, but the titre of
the mutant virus is slightly lower than that of the parental
virus. We hypothesize that RHDV VP2 might not
participate in the production of progeny virions but may
in fact represent an important virulence gene. In agreement
with this, we have found that the RHDV VP2 protein can
induce apoptosis when transfected into host cells (data not
shown) and that deletion of the protein decreases the
ability of RHDV to induce apoptosis and cell death in
tissue culture cells (data not shown). Of course, to
determine more precisely the role of VP2 in the RHDV
life cycle, additional experimental evidence is required and
will be the focus of our future research.
Our results also showed that deletion of VP2 decreased the
levels of the RHDV subgenomic RNA up to 60 h post-
transfection, but expression of the RHDV mutant increased
gradually in line with culture time and its expression
exceeded that of parental virus at 72 h post-infection.
Quantification of the RHDV genome during virus
replication by qRT-PCR demonstrated that the replication
level of the VP2 deletion mutant was reproducibly higher
than that of the parental virus. This increased level of
genome replication in the absence of VP2 is in agreement
with our hypothesis that VP2 may be involved in the
induction of apoptosis during infection, as VP2 expression
would result in cell death at an earlier stage of the virus life
In summary, we have constructed a convenient and robust
system to allow manipulation of the RHDV genome and
utilized this system to investigate the effects of VP2
deletion on viral replication. These data and the reverse
genetics system described herein will undoubtedly aid in
the study of the molecular mechanisms of RHDV
replication and pathogenesis.
We would like to thank Dr Ian G. Goodfellow, Department of
Virology, Faculty of Medicine, Imperial College London, UK, for
critical reading of the manuscript. This work was funded by grants
from the National Basic Research (973) Program (2006CB708209),
the Zhejiang Natural Sciences Foundation (Y305047 and Y307582),
the Chinese Natural Sciences Foundation (30670074) and the
Graveness Technology Program of the Science and Technology
Department of Zhejiang Province (2007C12G4010009).
Bertolotti-Ciarlet, A., Crawford, S. E., Hutson, A. M. & Estes, M. K.
(2003). The 39 end of Norwalk virus mRNA contains determinants
that regulate the expression and stability of the viral capsid protein
VP1: a novel function for the VP2 protein. J Virol 77, 11603–11615.
G. Liu and others
3084 Journal of General Virology 89
Boga, J. A., Martin Alonso, J. M., Marin, M. S., Casais, R, Lo ´pez
Va ´zquez, A., Machin, A. & Parra, F. (1999). Genome organisation and
gene expression strategies of rabbit haemorrhagic disease virus. Recent
Res Devel Virol 1, 107–119.
Chang, K.-O., Sosnovtsev, S. S., Belliot, G., Wang, Q., Saif, L. J. &
Green, K. Y. (2005). Reverse genetics system for porcine enteric
calicivirus, a prototype sapovirus in the Caliciviridae. J Virol 79, 1409–
Chaudhry, Y., Skinner, M. A. & Goodfellow, I. G. (2007). Recovery of
genetically defined murine norovirus in tissue culture using a fowlpox
virus expressing T7 RNA polymerase. J Gen Virol 88, 2091–2100.
Glass, P. J., White, L. J., Ball, J. M., Leparc-Goffart, I., Hardy, M. E. &
Estes, M. K. (2000). Norwalk virus open reading frame 3 encodes a
minor structural protein. J Virol 74, 6581–6591.
Glass, P. J., Zeng, C. Q. & Estes, M. K. (2003). Two non-overlapping
domains on the Norwalk virus ORF3 protein are involved in the
formation of the phosphorylated 35K protein and ORF3–capsid
protein interactions. J Virol 77, 3569–3577.
Gregg, D. A., House, C., Meyer, R. & Berninger, M. (1991). Viral
haemorrhagic disease of rabbits in Mexico: epidemiology and viral
characterization. Rev Sci Tech 10, 435–451.
Kaiser, W. J., Chaudhry, Y., Sosnovtsev, S. V. & Goodfellow, I. G.
(2006). Analysis of protein–protein interactions in the feline
calicivirus replication complex. J Gen Virol 87, 363–368.
Liu, S. J., Xue, H. P., Pu, B. Q. & Qian, S. H. (1984). A new viral disease
in rabbits. Anim Husb Vet Med 16, 253–255.
Liu, G. Q., Zhang, Y. Y., Ni, Z., Yun, T., Sheng, Z. T., Liang, H. L., Hua,
J. G., Li, S. M., Du, Q. Y. & Chen, J. P. (2006). Recovery of infectious
rabbit hemorrhagic disease virus from rabbit after direct inoculation
with in vitro-transcribed RNA. J Virol 80, 6597–6602.
Meyers, G. (2003). Translation of the minor capsid protein of a
calicivirus is initiated by a novel termination-dependent reinitiation
mechanism. J Biol Chem 278, 34051–34060.
Meyers, G., Wirblich, C. & Thiel, H.-J. (1991). Genomic and
subgenomic RNAs of rabbit hemorrhagic disease virus are both
protein linked and packaged into particles. Virology 184, 677–686.
Meyers, G., Wirblich, C., Thiel, H. J. & Thumfart, J. O. (2000). Rabbit
hemorrhagic disease virus: genome organization and polyprotein
processing of a calicivirus studied after transient expression of cDNA
constructs. Virology 276, 349–363.
Mitro, S. & Krauss, H. (1993). Rabbit hemorrhagic disease: a review
with special reference to its epizootiology. Eur J Epidemiol 9, 70–78.
Morales, M., Barcena, J., Ramirez, M. A., Boga, J. A., Parra, F. &
Torres, J. M. (2004). Synthesis in vitro of rabbit hemorrhagic disease
virus subgenomic RNA by internal initiation on(2)sense genomic
RNA: mapping of a subgenomic promoter. J Biol Chem 279, 17013–
Mutze, G., Cooke, B. & Alexander, P. (1998). The initial impact of
rabbit hemorrhagic disease on European rabbit populations in South
Australia. J Wildl Dis 34, 221–227.
Nowotny, N., Bascunana, C. R., Ballagi-Pordany, A., Gavier-Widen, D.,
Uhlen, M. & Belak, S. (1997). Phylogenetic analysis of rabbit
haemorrhagic disease and European brown hare syndrome viruses by
comparison of sequences from the capsid protein gene. Arch Virol 142,
Oehmig, A., Buttner, M., Weiland, F., Werz, W., Bergemann, K. &
Pfaff, E. (2003). Identification of a calicivirus isolate of unknown
origin. J Gen Virol 84, 2837–2845.
Ohlinger, V. F., Haas, B., Meyers, G., Weiland, F. & Thiel, H.-J. (1990).
Identification and characterization of the virus causing rabbit
hemorrhagic disease. J Virol 64, 3331–3336.
Parra, F., Boga, J. A., Marin, M. S. & Casais, R. (1993). The amino
terminal sequence of VP60 from rabbit hemorrhagic disease virus
supports its putative subgenomic origin. Virus Res 27, 219–228.
Sosnovtsev, S. V. & Green, K. Y. (2000). Identification and genomic
mapping of the ORF3 and VPg proteins in feline calicivirus virions.
Virology 277, 193–203.
Sosnovtsev, S. V., Belliot, G., Chang, K. O., Onwudiwe, O. & Green,
K. Y. (2005). Feline calicivirus VP2 is essential for the production of
infectious virions. J Virol 79, 4012–4024.
Thumfart, J. O. & Meyers, G. (2002). Feline calicivirus: recovery of
wild-type and recombinant viruses after transfection of cRNA or
cDNA constructs. J Virol 76, 6398–6407.
Wirblich, C., Meyers, G., Ohlinger, V. F., Capucci, L., Eskens, U.,
Haas, B. & Thiel, H. J. (1994). European brown hare syndrome virus:
relationship to rabbit hemorrhagic disease virus and other calicivirus.
J Virol 68, 5164–5173.
Wirblich, C., Sibilia, M., Boniotti, M. B., Rossi, C., Thiel, H. J. &
Meyers, G. (1995). 3C-like protease of rabbit hemorrhagic disease
virus: identification of cleavage sites in the ORF1 polyprotein and
analysis of cleavage specificity. J Virol 69, 7159–7168.
Wirblich, C., Thiel, H. J. & Meyers, G. (1996). Genetic map of the
calicivirus rabbit hemorrhagic disease virus as deduced from in vitro
translation studies. J Virol 70, 7974–7983.
A reverse genetics system of RHDV