Characterization of Ross River virus tropism and virus-induced inflammation in a mouse model of viral arthritis and myositis.
ABSTRACT Mosquito-borne alphaviruses are a significant cause of both encephalitic and arthritic disease in humans worldwide. In contrast to the encephalitic alphaviruses, the pathogenesis of alphavirus-induced arthritic disease is not well understood. Utilizing a mouse model of Ross River virus (RRV) disease, we found that the primary targets of RRV infection are bone, joint, and skeletal muscle tissues of the hind limbs in both outbred CD-1 mice and adult C57BL/6J mice. Moreover, histological analyses demonstrated that RRV infection resulted in severe inflammation of these tissues. Characterization of the inflammatory infiltrate within the skeletal muscle tissue identified inflammatory macrophages, NK cells, and CD4+ and CD8+ T lymphocytes. To determine the contribution of the adaptive immune system, the outcome of RRV-induced disease was examined in C57BL/6J RAG-1(-/-) mice, which lack functional T and B lymphocytes. RAG-1(-/-) and wild-type mice developed similar disease signs, infiltration of inflammatory macrophages and NK cells, and muscle pathology, suggesting that the adaptive immune response does not play a critical role in the development of disease. These results establish the mouse model of RRV disease as a useful system for the identification of viral and host factors that contribute to alphavirus-induced arthritis and myositis.
- SourceAvailable from: Suan-Sin Foo[Show abstract] [Hide abstract]
ABSTRACT: Background Arthritogenic alphaviruses such as Ross River virus (RRV) and chikungunya virus (CHIKV) have caused widespread outbreaks of chronic polyarthritis. The inflammatory responses in alphavirus-induced arthritis and osteoarthritis (OA) share many similar features, which suggests the possibility of exacerbated alphavirus-induced bone pathology in individuals with pre-existing OA. Here, we investigated the susceptibility of osteoblasts (OBs) from OA patients to RRV infection and dissected the immune mechanisms elicited from infection.Methods Primary hOBs obtained from trabecular bone of healthy donors and OA patients were infected with RRV. Infectivity and viral replication were determined using flow cytometry and plaque assay, respectively. Real-time PCR was performed to determine expression kinetics of type I interferon (IFN)-related immune mediators and osteotropic factors.ResultsOA hOBs showed enhanced RRV infectivity and replication during infection, which was associated with delayed induction of IFN-ß and RIG-I expression. Enhanced susceptibility of OA hOBs to RRV was associated with a more pronounced increase in RANKL/OPG ratio and expression of osteotropic factors (IL-6, IL-1ß, TNF-¿ and CCL2) in comparison to RRV-infected healthy hOBs.Conclusions Delayed activation of type I IFN-signalling pathway may have contributed to enhanced susceptibility to RRV infection in hOBs from OA patients. RRV-induced increases in RANKL/OPG ratio and expression of osteotropic factors that favour bone resorption, which may be exacerbated during osteoarthritis. This study provides the novel insight that osteoarthritis may be a risk factor for exacerbated arthritogenic alphaviral infection.Virology Journal 11/2014; 11(1):189. · 2.09 Impact Factor
- [Show abstract] [Hide abstract]
ABSTRACT: The recent epidemic of the arthritogenic alphavirus, chikungunya virus (CHIKV) has prompted a quest to understand the correlates of protection against virus and disease in order to inform development of new interventions. Herein we highlight the propensity of CHIKV infections to persist long term, both as persistent, steady-state, viraemias in multiple B cell deficient mouse strains, and as persistent RNA (including negative-strand RNA) in wild-type mice. The knockout mouse studies provided evidence for a role for T cells (but not NK cells) in viraemia suppression, and confirmed the role of T cells in arthritis promotion, with vaccine-induced T cells also shown to be arthritogenic in the absence of antibody responses. However, MHC class II-restricted T cells were not required for production of anti-viral IgG2c responses post CHIKV infection. The anti-viral cytokines, TNF and IFNγ, were persistently elevated in persistently infected B and T cell deficient mice, with adoptive transfer of anti-CHIKV antibodies unable to clear permanently the viraemia from these, or B cell deficient, mice. The NOD background increased viraemia and promoted arthritis, with B, T and NK deficient NOD mice showing high-levels of persistent viraemia and ultimately succumbing to encephalitic disease. In wild-type mice persistent CHIKV RNA and negative strand RNA (detected for up to 100 days post infection) was associated with persistence of cellular infiltrates, CHIKV antigen and stimulation of IFNα/β and T cell responses. These studies highlight that, secondary to antibodies, several factors are involved in virus control, and suggest that chronic arthritic disease is a consequence of persistent, replicating and transcriptionally active CHIKV RNA.PLoS Neglected Tropical Diseases 12/2014; 8(12):e3354. · 4.49 Impact Factor
- [Show abstract] [Hide abstract]
ABSTRACT: Part of the Togaviridae family, alphaviruses are arthropod-borne viruses that are widely distributed throughout the globe. Alphaviruses are able to infect a variety of vertebrate hosts, but in humans infection can result in extensive morbidity and mortality. Symptomatic infection can manifest as fever, an erythematous rash and/or significant inflammatory pathologies such as arthritis and encephalitis. Recent overwhelming outbreaks of alphaviral disease have highlighted the void in our understanding of alphavirus pathogenesis and the re-emergence of alphaviruses has given new impetus to anti-alphaviral drug design. In this review, the development of viable mouse models of Old Word and New World alphaviruses is examined. How mouse models that best replicate human disease have been used to elucidate the immunopathology of alphavirus pathogenesis and trial novel therapeutic discoveries is also discussed.Journal of General Virology 10/2014; · 3.53 Impact Factor
JOURNAL OF VIROLOGY, Jan. 2006, p. 737–749
Copyright © 2006, American Society for Microbiology. All Rights Reserved.
Vol. 80, No. 2
Characterization of Ross River Virus Tropism and Virus-Induced
Inflammation in a Mouse Model of Viral Arthritis and Myositis
Thomas E. Morrison,1,2,3Alan C. Whitmore,3Reed S. Shabman,1,2,3Brett A. Lidbury,4
Suresh Mahalingam,4and Mark T. Heise1,2,3*
Department of Genetics,1Department of Microbiology and Immunology,2and Carolina Vaccine Institute,3University of
North Carolina at Chapel Hill, Chapel Hill, North Carolina 27599, and Viral Arthritis/Asthma Research Group,
School of Health Sciences, University of Canberra, Canberra, ACT 2601, Australia4
Received 25 August 2005/Accepted 22 October 2005
Mosquito-borne alphaviruses are a significant cause of both encephalitic and arthritic disease in humans
worldwide. In contrast to the encephalitic alphaviruses, the pathogenesis of alphavirus-induced arthritic
disease is not well understood. Utilizing a mouse model of Ross River virus (RRV) disease, we found that the
primary targets of RRV infection are bone, joint, and skeletal muscle tissues of the hind limbs in both outbred
CD-1 mice and adult C57BL/6J mice. Moreover, histological analyses demonstrated that RRV infection
resulted in severe inflammation of these tissues. Characterization of the inflammatory infiltrate within the
skeletal muscle tissue identified inflammatory macrophages, NK cells, and CD4?and CD8?T lymphocytes. To
determine the contribution of the adaptive immune system, the outcome of RRV-induced disease was examined
in C57BL/6J RAG-1?/?mice, which lack functional T and B lymphocytes. RAG-1?/?and wild-type mice
developed similar disease signs, infiltration of inflammatory macrophages and NK cells, and muscle pathology,
suggesting that the adaptive immune response does not play a critical role in the development of disease. These
results establish the mouse model of RRV disease as a useful system for the identification of viral and host
factors that contribute to alphavirus-induced arthritis and myositis.
Mosquito-borne arthritogenic alphaviruses, such as Ross
River virus (RRV), Chikungunya virus, O’nyong-nyong virus,
and Mayaro virus, are a significant cause of infectious rheu-
matic disease worldwide (10, 36). For example, RRV, which is
endemic to Australia, causes several thousand cases of infec-
tious polyarthritis per year. Furthermore, these viruses are a
major concern due to their ability to emerge and cause major
epidemics. This is illustrated by an epidemic of Ross River
virus disease in the South Pacific which involved greater than
60,000 patients in 1979 to 1980 (13) and a 1959 to 1962 epi-
demic of O’nyong-nyong fever in Africa which involved at least
2 million patients (39). This is further reinforced by the reports
that RRV has reemerged in Fiji, where the virus had appar-
ently been absent since the 1979–1980 epidemic (19), and by
recent Chikungunya and O’nyong-nyong fever outbreaks in
Africa and Asia (18, 21, 27, 30).
Severe arthritis/arthralgia is a shared symptom of many of
the alphavirus-induced diseases. The clinical course of RRV
infection, which causes severe acute polyarthritis, is one of the
best-characterized alphavirus-induced arthritic diseases. RRV-
induced disease symptoms include fever, rash, myalgia, and
pain and stiffness in the joints (13). Muscle and joint pain in
afflicted individuals may persist for weeks to months, and anti-
inflammatory drugs are the best current treatment for RRV
disease (12, 13). The arthritic disease is thought to be initiated
by viral replication and inflammatory infiltrates in the affected
joints (7, 35). This is largely based on the detection of RRV
RNA in the synovia from the knees of patients infected with
RRV (35) and the detection of RRV antigen in synovial infil-
trates from affected joints (6). RRV-induced arthritis is char-
acterized by inflammatory infiltrates comprised largely of
mononuclear cells. Characterization of these infiltrates sug-
gests that monocytes/macrophages are a major constituent of
the infiltrate (6, 14), while immunohistological studies of sy-
novial biopsy samples have also identified CD4?and CD8?T
lymphocytes within the inflammatory infiltrates (35).
Although a large number of studies have focused on the
pathogenesis of alphavirus-induced encephalitis, the mecha-
nisms by which arthritogenic alphaviruses cause disease are
largely unknown. Recently, Lidbury et al. reported that RRV-
infected 17- to 21-day-old outbred mice developed severe dis-
ease characterized by inflammation of muscle tissue and mus-
cle damage (22). That study demonstrated that disease signs,
such as hind limb dragging and muscle pathology, were ame-
liorated following treatment of mice with macrophage-toxic
agents, suggesting a critical role for host immunity and mac-
rophages in mediating RRV-induced disease.
In this study, we utilized the mouse model of RRV disease
described by Lidbury et al. (22) to identify the primary sites of
RRV replication. In addition, we report that RRV-infected
24-day-old C57BL/6J (B6) mice developed severe disease char-
acterized by inflammation of hind limb bone and joint-associ-
ated tissues as well as skeletal muscle tissue. Finally, to deter-
mine the contribution of adaptive immune responses to the
development of RRV disease, we examined the outcome of
RRV infection in C57BL/6J RAG-1?/?mice, which lack func-
tional T and B lymphocytes. In addition to elucidating mech-
anisms by which arthritogenic alphaviruses cause disease, these
findings indicate that RRV infection of B6 mice represents a
* Corresponding author. Mailing address: The Carolina Vaccine
Institute, University of North Carolina at Chapel Hill, 827 Mary Ellen
Jones Bldg., CB no. 7292, Chapel Hill, NC 27599. Phone: (919) 843-
1492. Fax: (919) 843-6924. E-mail: email@example.com.
powerful animal model for studying virus-induced inflamma-
tion and immunopathology.
MATERIALS AND METHODS
Viruses and cells. Viral stocks of the wild-type T48 strain of RRV were
generated by in vitro transcription of SacI-linearized plasmid pRR64 (generously
provided by Richard Kuhn, Purdue University), which encodes the full-length
T48 cDNA clone (20), by using SP6-specific mMessage mMachine in vitro tran-
scription kits (Ambion). The T48 stain of RRV was initially isolated from Aedes
vigilax mosquitoes in Queensland, Australia (5). Prior to cDNA cloning, the virus
was passaged 10 times in suckling mouse brain, followed by two passages on Vero
cells (3). Full-length transcripts were electroporated into BHK-21 cells (ATCC
CRL 8544) using a Bio-Rad electroporator as described previously (16). Culture
supernatants were harvested at 24 h after electroporation, centrifuged for 20 min
at 3,000 rpm, aliquoted, and stored at ?80°C. Virus was titrated by plaque assay
on BHK-21 cells as described previously (34).
To generate an RRV that expresses the enhanced green fluorescent protein
(EGFP), a second RRV 26S promoter sequence was inserted at the 3? end of the
viral genome, followed by the coding sequence for EGFP. In brief, the 26S
subgenomic promoter region of Ross River virus from nucleotides 7300 to 7507
of pRR64 was PCR amplified. The resulting PCR product had engineered
sequential NotI and SpeI restriction sites 46 nucleotides downstream of the 26S
RNA start site and was flanked by HindIII sites. This PCR product was intro-
duced into position 11,330 in the pRR64 sequence to produce the plasmid
pRR64(26S). The EGFP coding sequence was removed from plasmid
pREP91Egfp (16) and introduced into pRR64(26S) using the NotI and SpeI
restriction sites downstream of the second 26S subgenomic promoter to generate
plasmid pRR64-Egfp. pRR64-Egfp was linearized with SacI and used as a tem-
plate for in vitro transcription and virus production as described above for
pRR64. EGFP expression in virus-infected cells was confirmed by fluorescence
BHK-21 cells were grown in ?-minimal essential medium (Gibco) supple-
mented with 10% donor calf serum, 10% tryptose phosphate broth, and 0.29
Mice. Specific-pathogen-free pregnant female outbred CD-1 mice at 13 to 15
days of gestation were obtained from Charles River Breeding Laboratories
(Raleigh, North Carolina). C57BL/6J mice, C57BL/6J RAG-1?/?mice, and
?MT mice were obtained from The Jackson Laboratory (Bar Harbor, Maine)
and bred in-house. ?MT mice carry a stop codon and the neomycin gene cassette
in the first transmembrane exon of the ? chain, resulting in B-lymphocyte
deficiency (17). In some experiments, mice were obtained from the animal
breeding establishment in the John Curtin School of Medical Research, Australia.
Animal housing and care at UNC were in accordance with all UNC-CH Insti-
tutional Animal Care and Use Committee guidelines. Although RRV is classified
as a biosafety level 2 pathogen, due to its exotic nature, all mouse studies in the
United States were performed in a biosafety level 3 laboratory.
Mice were inoculated in the left rear footpad with 103PFU of virus in diluent
(phosphate-buffered saline [PBS]–1% donor calf serum) in a 10-?l volume.
Alternatively, in a subset of studies, mice received the virus subcutaneously in the
thorax below the right forelimb in a 50-?l volume. No significant differences in
viral replication or development of disease were detected in mice inoculated
from either route. Mock-infected animals received diluent alone. Mice were
monitored for disease signs and weighed at 24 h intervals. The clinical signs of
disease were determined by assessing grip strength and altered gait. Grip
strength and hind limb weakness were assessed by testing the ability of each
mouse to support itself while suspended from a wire cage. Mice were scored as
follows: 0, no disease signs; 1, ruffled fur; 2, very mild hind limb weakness; 3, mild
hind limb weakness; 4, moderate hind limb weakness and dragging of hind limbs;
5, severe hind limb weakness/dragging; 6, complete loss of hind limb function; 7,
moribund; and 8, death. To determine viral titers in tissues, mice were sacrificed
by exsanguination and perfused with 1? PBS. The popliteal lymph node, right
and left ankles, right and left quadriceps muscles, spleen, brain, and spinal cord
(divided into thoracic and lumbar regions) were removed by dissection and
weighed. Tissues were homogenized in 1? PBS supplemented with 1% donor
calf serum, Ca2?, and Mg2?and stored at ?80°C until viral load was assessed by
a standard plaque assay on BHK-21 cells.
For histological analysis, mice were sacrificed by exsanguinations and perfused
with PBS–4% paraformaldehyde, pH 7.3. Following fixation of tissues and fur-
ther decalcification of bone-associated tissues, all tissues were embedded in
paraffin and 5-?m sections were prepared by the UNC histopathology core
facility. To determine the extent of inflammation, tissues were stained with
hematoxylin and eosin (H & E). Myelin was stained with luxol fast blue followed
by a periodic acid-Schiff counterstain. Sections were analyzed using a Nikon
Microphot-FXA microscope fitted with an Optronics DEI 750 three-chip charge-
coupled-device camera for digital imaging.
In situ hybridization. In situ hybridization was performed as described previ-
ously (15). Briefly, a35S-labeled RRV-specific riboprobe (complementary for
RRV nucleotides 7300 to 7775) was generated with an SP6-specific MAXIscript
in vitro transcription kit (Ambion) from a NotI-linearized plasmid. A riboprobe
complementary for the EBER2 gene from Epstein-Barr virus was used as a
negative control. Deparaffinized tissue sections were hybridized with 5 ? 104
cpm/?l of35S-labeled riboprobes overnight. Tissues were washed, dehydrated
through graded ethanol, and immersed in Nitro Blue Tetrazolium autoradiog-
raphy emulsion (Kodak). Following development, sections were counterstained
with hematoxylin and silver grain deposition was analyzed by light microscopy.
Flow cytometry. Mice were inoculated as described above, sacrificed by exsan-
guination at 5 and 7 days postinfection (dpi), and perfused for 10 min with 1?
PBS. Quadriceps muscles and spleens were dissected, minced, and incubated for
2 h with vigorous shaking at 37°C in digestion buffer (RPMI 1640, 10% fetal
bovine serum, 15 mM HEPES, 2.5 mg/ml collagenase A [Roche], 1.7 mg/ml
DNase I [Sigma]). Following digestion, cells were passed through a 40-?m cell
strainer, red blood cells were lysed (spleens only), cells were washed in wash
buffer (1? Hanks balanced salt solution, 15 mM HEPES), and total viable cells
were determined by trypan blue exclusion. Cells were incubated with anti-mouse
Fc?RII/III (2.4G2; BD Pharmingen) for 20 min on ice to block nonspecific
antibody binding and then stained in fluorescence-activated cell sorting staining
buffer (1? Hanks balanced salt solution, 1% fetal bovine serum, 2% normal
rabbit serum) with the following antibodies from eBioscience: F4/80-fluorescein
FIG. 1. Ross River virus-induced disease in 15-day-old CD-1 mice.
Fifteen-day-old CD-1 mice were infected with 103PFU of RRV (?) by
subcutaneous injection in the left rear footpad. Mock-infected mice
(?) were injected with diluent alone. (A) Mice were scored for devel-
opment of hind limb dysfunction and disease based on the following
scale: 0, no disease signs; 1, ruffled fur; 2, very mild hind limb weak-
ness; 3, mild hind limb weakness; 4, moderate hind limb weakness and
dragging of hind limbs; 5, severe hind limb weakness/dragging; 6,
complete loss of hind limb function; 7, moribund; and 8, death.
(B) Mice were monitored for weight gain or loss at 24-h intervals. Each
data point represents the arithmetic mean ? standard deviation (SD)
for 7 (mock-infected) or 25 (RRV-infected) mice.
738 MORRISON ET AL. J. VIROL.
isothiocyanate, NK1.1-phycoerythrin, Ly6C/G (Gr1)-phycoerythrin, CD3-fluo-
rescein isothiocyanate, CD4-biotin, CD19–allophycocyanin (APC), CD8?-APC,
CD11b-APC, and B220-APC. Biotin conjugates were detected with streptavidin-
peridinin chlorophyll protein (eBioscience). Cells were fixed overnight in 2%
paraformaldehyde and analyzed on a FACSCalibur (Becton Dickinson) using
Ross river virus replicates in bone and joint-associated tis-
sues and skeletal muscle. Infection of 14- to 21-day-old out-
bred mice with the T48 strain of RRV has previously been
demonstrated to induce inflammation and destruction of hind
limb skeletal muscle (22). To further evaluate this model for its
utility in investigating the pathogenesis of RRV-induced dis-
ease, we sought to identify the sites of viral replication in vivo.
As a first step, studies were performed to ensure that CD-1
mice exhibited the same phenotype upon RRV infection as the
Swiss outbred mice used in previous studies (22). Fourteen- to
15-day-old CD-1 outbred mice were inoculated with 103PFU
of RRV by subcutaneous injection in the left rear footpad and
monitored for virus-induced morbidity and mortality. Infected
mice developed severe disease signs characterized by loss of
hind limb gripping, hind limb dragging (Fig. 1A), and lack of
weight gain (Fig. 1B). These disease signs became apparent 4
to 5 dpi, peaked at 10 dpi, and resolved by 20 dpi (Fig. 1A and
B). These findings are consistent with previous observations of
Swiss outbred mice. However, up to 75% of Swiss outbred mice
were reported to have succumbed to infection (22), whereas
only 16% of CD-1 outbred mice used in this study succumbed
To begin to identify the sites of RRV replication in vivo,
viral replication in tissues at various times postinfection was
assessed by plaque assay. Peak titers of infectious virus were
detected at 24 to 48 h postinfection (hpi) (Fig. 2). Viral titers
of up to 109PFU/gram were detected in ankle-associated tis-
sue (Fig. 2A) and quadriceps muscle tissue (Fig. 2B). In con-
trast, peak viral titers in the brain (Fig. 2D) and spleen (Fig.
2E) were markedly lower than those observed in both ankle
and skeletal muscle tissues.
In order to identify specific sites of RRV replication, in
situ hybridization using an35S-labeled riboprobe specific for
RRV was performed on sections derived from hind limb
tissues of 14- to 15-day-old CD-1 mice infected with RRV.
FIG. 2. Ross River virus tissue titers in 15-day-old CD-1 mice. Fifteen-day-old CD-1 mice were infected with 103PFU of RRV by subcutaneous
injection in the left rear footpad. At 12, 24, 48, 72, 96, and 120 hpi, ankle (A), quadriceps muscle (B), serum (C), brain (D), and spleen (E) were
harvested and homogenized, and the amount of infectious virus present was quantified by plaque assay on BHK-21 cells. Each data point represents
the arithmetic mean ? SD for three mice.
VOL. 80, 2006 ROSS RIVER VIRUS TROPISM AND INFLAMMATORY DISEASE739
At 24 hpi, abundant RRV-specific signal was detected
within cells of the periosteum lining bones of the hind limbs
(Fig. 3A) and cells within synovial tissue of the knee joint
(Fig. 3B). In addition, RRV-specific signal was observed in
cells within tendons and ligaments and associated with ar-
ticular surfaces (data not shown). A similar distribution of
RRV replication within bone- and joint-associated tissues of
the hind limbs was also detected at both 48 hpi (Fig. 3D and
E) and 72 hpi (Fig. 3G and H). Interestingly, although RRV
replication was not observed within myofibers of hind limb
skeletal muscle tissue at 24 hpi (Fig. 3C), RRV-specific, in
situ signal was detected within connective tissues, such as
the perimysium and tendons, that are associated with skel-
etal muscle (data not shown). By 48 and 72 hpi, extensive
signal was detected within the myofibers of skeletal muscle
tissue (Fig. 3F and I). No signal was detected in tissue sections
from infected animals hybridized with a riboprobe specific for
the EBER2 gene from Epstein-Barr virus or in tissues from
mock-infected animals hybridized with the RRV-specific ribo-
probe (data not shown).
To confirm the in situ hybridization studies, an RRV that
expressed enhanced green fluorescent protein (RRV-EGFP)
in infected cells was constructed by inserting the EGFP coding
sequence downstream of a second 26S subgenomic promoter
placed at the 3? end of the viral genome (Fig. 4A). Initial
studies demonstrated that RRV-EGFP was attenuated in 14-
to 15-day-old CD-1 mice; however, infection of 10- to 12-day-
old mice resulted in similar disease signs (data not shown).
Therefore, 10- to 12-day-old CD-1 outbred mice were inocu-
lated with 103PFU of RRV-EGFP in the ventral thorax, mice
were sacrificed at the times postinfection indicated in Fig. 4,
and tissue sections were analyzed for GFP expression. At 24 to
48 hpi, extensive viral replication, as indicated by GFP expres-
sion, was observed in synovial tissues of hind limb joints. Major
sites of replication included cells lining the synovial cavity of
the ankle joint (Fig. 4A and B), cells within the persiosteum
(Fig. 4C), and cells within the tendons at junctions with the
skeletal muscle (data not shown). Though high viral titers were
detectable by plaque assay within the skeletal muscle at 24 to
48 hpi (Fig. 2B), few GFP-positive cells were detectable in the
FIG. 3. Ross River virus replication within bone and joint-associated connective tissues and skeletal muscle tissue. Fifteen-day-old CD-1 mice
were infected with 103PFU of RRV by subcutaneous injection in the ventral thorax. Mock-infected mice were injected with diluent alone. Mice
were sacrificed at 24 (A, B, C), 48 (D, E, F), and 72 (G, H, I) hours postinfection and perfused with 4% paraformaldehyde. Following
decalcification, 5-?m-thick paraffin-embedded sections derived from the hind limbs were probed with35S-labeled riboprobes complementary for
RRV (A-I) or the EBER2 gene from Epstein-Barr virus (data not shown). M, muscle; B, bone. (A) RRV-specific in situ signal in tarsal bone
periosteum. (B) RRV-specific in situ signal in synovial connective tissue of the knee joint. (C) Absence of RRV-specific in situ signal in hind limb
skeletal muscle. (D) RRV-specific in situ signal in tarsal bone periosteum and associated skeletal muscle. (E) RRV-specific in situ signal in synovial
tissue of a tarsal joint. (F) RRV-specific in situ signal in hind limb skeletal muscle tissue. (G) RRV-specific in situ signal in metatarsal bone
periosteum and associated skeletal muscle. (H) RRV-specific in situ signal in hind limb tendon. (I) RRV-specific in situ signal in hind limb skeletal
740 MORRISON ET AL.J. VIROL.
skeletal muscle at these time points (data not shown). Consis-
tent with the in situ hybridization studies, at 72 hpi, high levels
of GFP expression were observed in large areas of hind limb
skeletal muscle tissue (Fig. 4D). That GFP expression accu-
rately represented the distribution of virally infected cells was
confirmed by staining sections from RRV-GFP-infected mice
with RRV-specific hyperimmune sera, which demonstrated
that GFP-positive cells exhibited RRV-specific staining (data
Taken together, these data indicate that hind limb bone and
joint-associated tissues and skeletal muscle tissue are primary
targets for RRV infection. Furthermore, the in situ hybridiza-
tion (Fig. 3) and GFP expression studies (Fig. 4) indicate that
the initial viral titers in the skeletal muscle at 12 to 24 hpi may
reflect either localized replication in connective tissue associ-
ated with skeletal muscle, such as the perimysium, or contam-
ination of the muscle preparations with infected tendon that
exhibited high levels of replication at early times. These find-
ings also suggest that the virus may initially infect joint-asso-
ciated connective tissues and subsequently spread into skeletal
muscle tissue, although additional studies are required to truly
address this possibility.
Ross river virus infection causes severe hind limb disease in
C57BL/6J mice. Previous studies of mice have suggested that
RRV infection induces an immunopathological inflammatory
disease (22). To understand the components of the host im-
mune response and other host determinants that contribute to
RRV-induced disease, we sought to characterize the outcomes
of RRV infection in inbred strains of mice as a starting point
for studies of mice deficient for specific host factors. RRV
infection of 15-day-old C57BL/6J (B6) mice resulted in severe
hind limb dysfunction similar to that observed following infec-
tion of 15-day-old outbred CD-1 mice. However, disease be-
came increasingly severe, and 100% of 15-day-old B6 mice
succumbed to infection (Table 1). Therefore, older mice were
evaluated for susceptibility to RRV disease. Infection of 24-
day-old B6 mice resulted in severe morbidity similar to that
observed in 14- to 15-day-old CD-1 mice (Fig. 5 and Table 1).
Disease signs following RRV infection of 24-day-old B6 mice
included failure to gain weight (Fig. 5B) and progressive sym-
metrical hind limb dysfunction ranging from loss of hind limb
gripping ability to very severe hind limb dragging (Fig. 5A).
FIG. 4. Ross River virus double-promoter virus targets cells within joint, bone, and skeletal muscle tissues. (A) Schematic diagram of Ross
River virus that was engineered to express EGFP by inserting a second RRV 26S subgenomic promoter at the 3? end of the viral genome followed
by the EGFP coding sequence. (B) Ten- to 12-day-old CD-1 mice were infected with 103PFU of RRV-EGFP by subcutaneous injection in the
ventral thorax. At 48 and 72 hpi, tissues were harvested and sections were analyzed for EGFP expression. SC, synovial cavity; M, muscle; P,
periosteum. The white arrows indicate the articular surface. (Subpanel A) Synovial cavity in the foot at 48 hpi. (Subpanel B) Synovial cavity in the
ankle at 48 hpi. (Subpanel C) Periosteum in the hind limb at 48 hpi. (Subpanel D) Hind limb skeletal muscle at 72 hpi.
TABLE 1. Morbidity and mortality following RRV infection
25 10016 10 ? 0.8
7 ? 0.717 100 100
C57BL/6 24 Mock
aAST, average survival time; NA, not applicable.
VOL. 80, 2006 ROSS RIVER VIRUS TROPISM AND INFLAMMATORY DISEASE741
Disease signs, such as hunched posture, tremulousness, or
spontaneous circling and falling, were not observed in RRV-
infected mice. In addition, RRV-infected 24-day-old B6 mice
appeared to completely recover from the disease by 25 to 30
dpi, as indicated by resumption of weight gain and an absence
of observable disease signs (Fig. 5).
Ross river virus replicates in bone and joint-associated tis-
sues and skeletal muscles of adult C57BL/6J mice. Plaque
assays were performed to measure the amount of infectious
virus within tissues of 24-day-old B6 mice at 12, 24, 48, 72, 96,
and 120 hpi with RRV. Peak viral titers were detected at 24 to
48 hpi in all tissues examined (Fig. 6). Similar to findings for
CD-1 mice, the highest viral titers detected were from ankle
tissue, quadriceps muscle tissue, and serum (Fig. 6A, B, and C,
respectively). In contrast, viral titers in the spleen (Fig. 6D),
brain (Fig. 6E), and upper and lower spinal cord (Fig. 6F) were
10- to 1,000-fold lower on a PFU/gram basis. Consistent with
the symmetrical signs of disease, viral titers detected in ankle
and skeletal muscle tissue harvested from the injected limb and
the contralateral limb (noninjected) were similar at each time
point (Fig. 6A and B), indicating that the observed replication
pattern is not limited to the injected limb.
RRV infection induces inflammation of bone and joint-as-
sociated tissues and skeletal muscle of adult C57BL/6J mice.
Comprehensive histological analyses of hind limb bone, joint,
and skeletal muscle tissue, as well as tissues of the central
nervous system, were performed to determine whether RRV
infection of B6 mice resulted in inflammation and pathology at
any of these sites. Little to no inflammation was observed at
times earlier than 5 dpi. By 5 dpi, in contrast to what occurred
with mock-infected mice (Fig. 7A), evaluation of bone and
joints within the feet of RRV-infected mice revealed numerous
inflammatory cells within the periosteum, synovial tissue, and
associated skeletal muscle tissue (Fig. 7B). At 7 dpi, inflam-
mation was absent in synovial tissue of mock-infected mice
(Fig. 7C) but was again detected in synovial tissue of RRV-
infected mice (Fig. 7D). Similarly, no inflammation was ob-
served in quadriceps skeletal muscle tissue in mock-infected
animals (Fig. 8A) or RRV-infected animals (Fig. 8B) at 3 dpi.
The presence of inflammatory infiltrates was first observed in
quadriceps muscle tissue at 5 dpi (Fig. 8C). Inflammation in
skeletal muscle tissue reached peak severity by 7 and 10 dpi
(Fig. 8D and E). Obvious signs of myofiber destruction, con-
sistent with previous findings (22, 31), were observed at 7 and
10 dpi (Fig. 8D and E). Both the RRV-induced inflammation
and pathology within hind limb tissues resolved by 25 to 30 dpi
(Fig. 8F). This resolution included the presence of centralized
nuclei within myofibers (Fig. 8F), indicating that muscle fiber
regeneration was occurring within RRV-infected animals (2,
38). Unlike with hind limb bone, joint, and skeletal muscle
tissues, we did not observe inflammatory infiltrates, demyelin-
ation, or other virus-induced pathology within brain and spinal
cord tissues, including the spinal root (data not shown). Focal
areas of hypercellularity and activated microglia were observed
within the spinal cords of RRV-infected mice; however, these
observations were quite subtle and were not associated with
any overt pathology. Taken together, these findings indicate
that the severe hind limb weakness and dysfunction that de-
velops in B6 mice following RRV infection is most likely due
to the severe inflammation and pathology observed in hind
limb bone, joint, and skeletal muscle tissues.
Identification of inflammatory infiltrates in vivo. To begin
to identify mechanisms involved in RRV-induced inflamma-
tion and pathology, the composition of inflammatory infiltrates
within the hind limb skeletal muscle was analyzed by flow
cytometry. The hind limb quadriceps muscle was chosen over
joint tissues for these analyses due to the larger tissue volume
resulting in greater cell yield. At 5 and 7 dpi, quadriceps mus-
cle tissue from each of the hind limbs of mock-infected or
RRV-infected mice were dissected, minced, and digested with
collagenase A and DNase I to generate single-cell suspensions.
To control for effects of the digestion protocol on cell surface
antigen expression, splenocytes from mock-infected and RRV-
infected animals were isolated in similar fashions. Isolated cells
were then stained with various antibodies directed against cell
surface antigens. Significant increases in both the percentages
and total numbers of natural killer (NK) cells (NK1.1?/CD3?)
and inflammatory macrophages (F4/80?/Gr-1?) (8, 25, 29)
within the skeletal muscle tissue were detected at 5 dpi com-
pared to those in mock-infected controls (Fig. 9A and C).
FIG. 5. Ross River virus infection induces severe hind limb disease
in 24-day-old C57BL/6J mice. Twenty-four-day-old C57BL/6J mice
were inoculated with 103PFU of RRV (?) by subcutaneous injection
in the left rear footpad. Mock-infected mice (?) were injected with
diluent alone. (A) Mice were scored for development of hind limb
dysfunction and disease based on the following scale: 0, no disease
signs; 1, ruffled fur; 2, very mild hind limb weakness; 3, mild hind limb
weakness; 4, moderate hind limb weakness and dragging of hind limbs;
5, severe hind limb weakness/dragging; 6, complete loss of hind limb
function; 7, moribund; and 8, death. (B) Mice were monitored for
weight gain or loss at 24-h intervals. Each data point represents the
arithmetic mean ? SD for three (mock-infected) or six (RRV-in-
fected) animals. Data are representative of four independent experi-
742MORRISON ET AL.J. VIROL.
Further analyses demonstrated that the F4/80?/Gr-1?popu-
lation was also CD11b?/CD11c?/B220?, confirming that these
cells are inflammatory macrophages (data not shown). In-
creased numbers of both CD4?(CD3?CD4?) and CD8?
(CD3?CD8?) T lymphocytes were also detected by 5 dpi,
albeit to a much lesser extent than with both NK cells and
inflammatory macrophages (Fig. 9A and C). By 7 dpi, the
number of NK cells and inflammatory macrophages detected
was similar to that observed at 5 dpi; however, a further in-
crease in the total number of both CD4?and CD8?T lym-
phocytes was detected within the skeletal muscle tissues of
infected mice (Fig. 9B and C). No significant CD19 staining
was detected at 7 dpi, suggesting a lack of B lymphocytes (data
not shown). In addition to identifying the above-mentioned
cell types, our analyses indicated that additional cell types were
also present within the inflammatory infiltrate, such as den-
dritic cells. However, further studies will be required to iden-
tify the exact nature of these additional cell types.
RRV induces inflammation and disease in RAG-1?/?mice.
Our analysis of the inflammatory infiltrates detected cells of
both the innate, such as NK cells and macrophages, and adap-
tive, such as CD4?and CD8?T lymphocytes, arms of the
immune response. However, previous studies have suggested
that the RRV-induced inflammatory disease is independent of
the adaptive immune response (22, 31). Therefore, to defini-
tively determine the contribution of the adaptive immune
system in the development of RRV-induced disease, 24-day-
old C57BL/6J RAG-1?/?mice, which lack functional T and B
lymphocytes (24, 33), were inoculated with 103PFU of RRV in
the left rear footpad and monitored for virus-induced disease.
FIG. 6. Ross River virus tissue titers in 24-day-old C57BL/6J mice. Twenty-four-day-old C57BL/6J mice were infected with 103PFU of RRV
by subcutaneous injection in the left rear footpad. At 12, 24, 48, 72, 96, and 120 hpi, the following tissues were harvested and homogenized and
the amount of infectious virus present was quantified by plaque assay on BHK-21 cells. (A) ?, ankle of injected leg; ?, ankle of noninjected leg.
(B) ?, quadriceps muscle of injected leg; ?, quadriceps muscle of noninjected leg. (C) Serum. (D) Spleen. (E) Brain. (F) ?, lower spinal cord;
?, upper spinal cord. Each data point represents the arithmetic mean ? SD for three mice.
VOL. 80, 2006ROSS RIVER VIRUS TROPISM AND INFLAMMATORY DISEASE 743
FIG. 7. Ross River virus induces inflammation in hind limb bone and joint tissues of C57BL/6J mice. Twenty-four-day-old C57BL/6J mice
were infected with 103PFU of RRV by subcutaneous injection in the left rear footpad. Mock-infected mice were injected with diluent alone.
M, muscle; B, bone; P, periosteum; ST, synovial tissue. At 5 days (A and B) and 7 days (C and D) postinfection, mice were perfused with
4% paraformaldehyde. Following decalcification, 5-?m-thick paraffin-embedded sections generated from ankle and foot tissues of mock-
infected (A and C) and RRV-infected (B and D) mice were H & E stained. Images (magnification, ?200) are representative of at least six
mice per group.
FIG. 8. Ross River virus induces inflammation in hind limb skeletal muscle tissue of C57BL/6J mice. Twenty-four-day-old C57BL/6J mice were
infected with 103PFU of RRV by subcutaneous injection in the left rear footpad. Mock-infected mice were injected with diluent alone. At 3, 5,
7, 10, and 30 dpi, mice were perfused with 4% paraformaldehyde and 5-?m-thick paraffin-embedded sections generated from the quadriceps
muscle were H & E stained. (A) Mock infection. (B) RRV infection at 3 dpi. (C) RRV infection at 5 dpi. (D) RRV infection at 7 dpi. (E) RRV
infection at 10 dpi. (F) RRV infection at 30 dpi. Images (magnification, ?200) are representative of three to six mice per group.
744 MORRISON ET AL.J. VIROL.
FIG. 9. CharacterizationofRossRivervirus-inducedinflammatoryinfiltrates.Twenty-four-day-oldC57BL/6Jmicewereinfectedwith103PFUofRRVby
muscle at 5 dpi. Dot plots shown for each stain are representative of three mice. Two independent experiments gave similar results. (B) Cell surface staining of
cells isolated from the quadriceps muscle at 7 dpi. Dot plots shown for each stain are representative of three mice per group. Three independent experiments
and CD8 T lymphocytes (CD3?/CD8?) from the quadriceps muscle of mock- and RRV-infected animals at 5 and 7 days postinfection. Data presented are the
means ? standard errors of the means for three to four mice per group and are representative of at least two independent experiments.
VOL. 80, 2006ROSS RIVER VIRUS TROPISM AND INFLAMMATORY DISEASE745
FIG. 10. Ross River virus-induced inflammation and disease in RAG-1?/?. Twenty-four-day-old C57BL/6J RAG-1?/?mice were infected with
103PFU of RRV (?) by subcutaneous injection in the left rear footpad. Mock-infected mice (?) were injected with diluent alone. (A) Mice were
scored for development of hind limb dysfunction and disease based on the following scale: 0, no disease signs; 1, ruffled fur; 2, very mild hind limb
weakness; 3, mild hind limb weakness; 4, moderate hind limb weakness and dragging of hind limbs; 5, severe hind limb weakness/dragging; 6,
complete loss of hind limb function; 7, moribund; and 8, death. Each data point represents the arithmetic mean ? SD for two (mock-infected) or
six (RRV-infected) mice and are representative of two independent experiments. (B) Mice were monitored for weight gain or loss at 24-h intervals.
Each data point represents the arithmetic mean ? SD for four (mock-infected) or six (RRV-infected) mice and are representative of two
746MORRISON ET AL. J. VIROL.
RAG-1?/?mice developed disease signs similar to those ob-
served in B6 mice, including failure to gain weight (Fig. 10B)
and severe symmetrical hind limb dysfunction (Fig. 10A). The
average clinical score at the peak of observable disease (day
12) in RAG-1?/?mice was 4.5 ? 0.38, compared to 5.9 ? 0.25
for wild-type B6 mice. Interestingly, RAG-1?/?mice recov-
ered from RRV-induced disease with kinetics similar to that
observed in wild-type B6 mice (Fig. 5 and 10A and B). In
addition, both ?MT and ?MT mice depleted of T lymphocytes
with anti-CD3 antibody also developed disease similar to that
of wild-type mice following RRV infection (data not shown).
Histological analyses demonstrated severe inflammation in the
hind limb skeletal muscle (Fig. 10C) and joint-associated tis-
sues (data not shown) of RAG-1?/?mice, which peaked in
severity from 7 to 10 dpi. The degree of inflammation and
severity of tissue destruction appeared very similar to that
observed in wild-type B6 mice. We next analyzed the compo-
sition of the inflammatory infiltrates in the skeletal muscle by
flow cytometry. Similar to what was observed in wild-type B6
mice, increases in the percentage and total number of inflam-
matory macrophages (F4/80?/Gr-1?) and NK cells (NK1.1?/
CD3?) in the quadriceps muscle of RRV-infected RAG-1?/?
mice compared to those in mock-infected controls were de-
tected (Fig. 10E). CD3?T lymphocytes were not detected
within the skeletal muscle tissues or spleens of RAG-1?/?
mice, confirming the lack of T lymphocytes in these mice (data
not shown). As was observed for the wild-type B6 mice, there
were additional cell populations present, including dendritic
cells (data not shown), that will require additional analysis to
The pathogenesis of alphavirus-induced arthritides, which
represent a significant worldwide disease threat, is poorly un-
derstood. Therefore, the recently described mouse model of
RRV-induced inflammation was used to study viral tropism
and the composition of the virus-induced inflammatory re-
sponse. This report extends earlier work with the RRV mouse
model by demonstrating that the primary targets of Ross River
virus infection in outbred CD-1 mice and inbred adult B6 mice
are bone, joint, and skeletal muscle tissues. Significantly, fur-
ther analyses demonstrated that RRV infection of adult B6
mice resulted in severe inflammation within these same tissues.
In addition to inflammatory macrophages and NK cells, CD4?
and CD8?T lymphocytes were found to be major components of
the virus-induced inflammatory response. However, RAG-1?/?
mice developed disease signs and infiltration of NK cells and
macrophages similar to those of wild-type B6 mice, suggesting
that the adaptive immune response does not play a critical role
in the development of disease. These studies demonstrate that
the mouse model of RRV disease will greatly facilitate the
identification of viral and host factors which contribute to the
induction and resolution of this severe inflammatory disease.
The detection of RRV in multiple bone and joint-associated
tissues, such as synovial tissue, periosteum, tendons, and liga-
ments, has not previously been described. These findings are
consistent with observations of RRV-infected humans in
whom both viral antigen and viral RNA have been detected
from synovial effusions and synovial biopsy samples (6). Addi-
tionally, infectious virus was detectable in the ankle joints of
RRV-infected mice by plaque assay by 12 hpi; however, infec-
tious RRV has not yet been recovered from the joints of
RRV-infected patients (13). Similar to previous reports, high
titers of RRV were also detected within skeletal muscle tissues
of infected mice (22, 26, 31). However, our targeting studies
demonstrated that there were very few RRV-infected muscle
fibers in the hind limbs at early times postinfection. By 48 to 72
hpi, large areas of RRV-infected muscle fibers were observed
in hind limb skeletal muscle tissue. These findings raise the
possibility that RRV may initially infect joint- or skeletal mus-
cle-associated connective tissues and subsequently spread into
skeletal muscle myofibers. Direct infection of skeletal muscle
tissue by RRV in humans has not been demonstrated, although
60% of patients diagnosed with Ross River virus disease ex-
perience myalgia (12). The route and spread of RRV from the
initial site of infection to joint and skeletal muscle tissue have
not been characterized fully. Studies performed with other
alphaviruses, such as Venezuelan equine encephalitis virus,
have suggested that this group of viruses may initially infect
skin dendritic cells (23). The infected dendritic cells migrate to
the draining lymph node, where the virus undergoes additional
rounds of replication and seeds a high-titer serum viremia,
resulting in viral spread to target tissues.
Histological analyses of tissues from adult B6 mice revealed
that RRV infection induced inflammation within joint-associ-
ated tissues, such as periosteum, tendons, and synovial tissue,
as well as skeletal muscle tissue. Inflammatory infiltrates were
first observed within these hind limb tissues at 5 dpi, peaked at
7 to 10 dpi, and were dramatically decreased in number by 20
to 30 dpi. The occurrence of inflammation within these tissues
occurred well after peak viral titers and correlated with the
observed hind limb weakness, which was first detectable at 4 to
5 dpi, became increasingly severe from 7 to 12 dpi, and re-
solved between 20 to 30 dpi. Although RRV has been reported
to induce encephalomyelitis characterized by central nervous
system demyelination in 1-week-old BALB/c mice (32), we
found no evidence of virus-induced inflammation, demyelina-
tion, or other pathology in the brains or spinal cords of 24-
day-old adult B6 mice. RRV infection of 7-day-old BALB/c
mice was also proposed to cause muscle destruction in the
absence of a host immune response (31), while immune pa-
thology clearly contributes to virus-induced pathology in our
system. The differences in mouse age and strain used in the two
independent experiments. (C) At 10 days postinfection, mice were perfused with 4% paraformaldehyde and 5-?m-thick paraffin-embedded
sections generated from the quadriceps muscle were H & E stained. Images (magnification, ?200) are representative of three mice per group. (D)
Total numbers and percentages of natural killer cells (NK1.1?/CD3?) and inflammatory macrophages (F4/80?/Gr-1?) from the quadriceps
muscles of mock-infected (black bars) or RRV-infected (gray bars) RAG-1?/?mice at 7 days postinfection. Data presented are the means ?
standard errors of the means for three mice per group and are representative of two independent experiments.
VOL. 80, 2006ROSS RIVER VIRUS TROPISM AND INFLAMMATORY DISEASE 747
studies may explain the differences between our results and
those previously reported. In addition, it has also been re-
ported that, in contrast to 1-week-old BALB/c mice, 4-week-
old BALB/c mice infected with RRV developed little to no
disease signs (31), indicating there are age- and strain-depen-
dent effects of RRV infection. Taken together, our findings
suggest that the observed disease signs in 24-day-old B6 mice
are most likely due to virus-induced pathology in hind limb
Consistent with previous work, our studies identified inflam-
matory macrophages as major constituents of the inflammatory
infiltrate in hind limb skeletal muscles of RRV-infected mice.
The presence of inflammatory macrophages, as well as a large
increase of NK cells, was readily detectable by 5 dpi. By 7 dpi,
increased numbers of both CD4?and CD8?T lymphocytes
were also found within hind limb skeletal muscle tissue. Al-
though our studies focused on the skeletal muscle, the cellular
reported following RRV infection of humans. Both NK cells and
macrophages have been detected within synovial exudates from
and CD8?T lymphocytes were detected in synovial tissue sec-
tions generated from knee joint biopsies (35).
A recent study demonstrated that treatment of mice with
macrophage-toxic agents prior to infection completely pre-
vented RRV-induced muscle inflammation (22), suggesting an
important role for innate immune responses in the develop-
ment of RRV-induced disease. Our studies of RAG-1?/?
mice, which develop inflammation and disease similar to those
of wild-type B6, underscore the importance of the innate im-
mune response. However, we cannot rule out the possibility of
a limited contribution of the adaptive response to the severity
of disease. In addition, RAG-1?/?mice recovered from RRV-
induced disease with kinetics similar to that observed in wild-
type B6 mice. Future studies will be aimed at determining
whether innate immune mechanisms are sufficient to control
RRV infection or whether other mechanisms are involved in
the resolution of disease in RAG-1?/?mice.
Infection of neonatal mice with the Tucson strain of cox-
sackievirus B1 (CVB1T) or 10- to 14-day-old mice with the DA
strain of Theiler’s murine encephalomyelitis virus has also
been shown to induce severe myositis in hind limb skeletal
muscle tissue (9, 28). Whereas the role of the immune re-
sponse in Theiler’s murine encephalomyelitis virus-induced
disease is not well understood, T lymphocytes have been dem-
onstrated to play a major role in CVB1T-induced disease (41),
suggesting that RRV and CVB1Tmay promote muscle patho-
logy by distinct mechanisms.
The mechanisms by which RRV or other arthritogenic al-
phaviruses trigger inflammatory responses are not understood.
Interestingly, when injected directly into murine knee joints,
double-stranded RNA (dsRNA) (which is formed during the
replication and transcription of RNA viruses such as RRV) is
itself arthritogenic (42). In addition, similarly to the critical
role of macrophages in the development of RRV-induced dis-
ease, depletion of monocytes completely prevented dsRNA-
induced arthritis. T and B lymphocytes were also found to be
dispensable for the development of dsRNA-induced arthritis
(42). A number of different molecules have been demonstrated
to contribute to the host’s detection of viral dsRNA. Toll-like
receptor 3 was not required for dsRNA-induced arthritis (42);
therefore, it will be interesting to determine the role of other
sensors of dsRNA in the development of RRV-induced dis-
ease, such as protein kinase R (4), or the newly identified
caspase activation and recruitment domain-containing RNA
helicases retinoic acid-inducible gene I (40) and mda-5 (1).
In addition to RRV, Chikungunya virus, O’nyong-nyong vi-
rus, and Mayaro virus, Sindbis group alphaviruses are also
associated with arthritis and arthralgia in humans (10). How-
ever, studies of adult mice have demonstrated that infection
with most Sindbis group alphaviruses results in encephalitic
disease (11, 37). Recent work has demonstrated that Sindbis
group alphaviruses, such as S.A.AR86 and TR339, replicated
in bone and joint-associated tissues of adult CD-1 mice; how-
ever, virus-induced inflammation or other pathology was not
observed within these tissues (15). Therefore, the RRV mouse
model represents a unique system for studying the pathogen-
esis of alphavirus-induced inflammatory disease in bone, joint,
and skeletal muscle tissue.
This research was supported by NIH research grant R01 AR47190.
Work was also funded by an Australian NHMRC Project grant
(303404). T.E.M. was supported by NIH postdoctoral fellowship F32
AR052600-01. S.M. is a recipient of an NHMRC R. D. Wright Fel-
We thank members of the Carolina Vaccine Institute and the
Johnston laboratory for helpful scientific discussions. We thank Nancy
Davis for critical reading of the manuscript. We also thank Dwayne
Muhammed and Kenya Madric for assistance with tissue culture; Jan-
ice Weaver, Robin Smith, and Wuhan Jiang at the LCCC/DLAM
UNC histopathology core facility; and Kinuko I. Suzuki for histological
analysis of brain and spinal cord tissue sections.
1. Andrejeva, J., K. S. Childs, D. F. Young, T. S. Carlos, N. Stock, S. Good-
bourn, and R. E. Randall. 2004. The V proteins of paramyxoviruses bind the
IFN-inducible RNA helicase, mda-5, and inhibit its activation of the IFN-
beta promoter. Proc. Natl. Acad. Sci. USA 101:17264–17269.
2. Charge, S. B., and M. A. Rudnicki. 2004. Cellular and molecular regulation
of muscle regeneration. Physiol. Rev. 84:209–238.
3. Dalgarno, L., C. M. Rice, and J. H. Strauss. 1983. Ross River virus 26 s
RNA: complete nucleotide sequence and deduced sequence of the encoded
structural proteins. Virology 129:170–187.
4. Diebold, S. S., M. Montoya, H. Unger, L. Alexopoulou, P. Roy, L. E. Haswell,
A. Al-Shamkhani, R. Flavell, P. Borrow, and C. Reise Sousa. 2003. Viral
infection switches non-plasmacytoid dendritic cells into high interferon pro-
ducers. Nature 424:324–328.
5. Doherty, R. L., R. H. Whitehead, B. M. Gorman, and A. K. O’Gower. 1963.
The isolation of a third group A arbovirus in Australia, with preliminary
observations on its relationship to epidemic polyarthritis. Aust. J. Sci. 26:
6. Fraser, J. R., A. L. Cunningham, B. J. Clarris, J. G. Aaskov, and R. Leach.
1981. Cytology of synovial effusions in epidemic polyarthritis. Aust. N. Z. J.
7. Fraser, J. R., V. M. Ratnamohan, J. P. Dowling, G. J. Becker, and G. A.
Varigos. 1983. The exanthem of Ross River virus infection: histology, loca-
tion of virus antigen and nature of inflammatory infiltrate. J. Clin. Pathol.
8. Geissmann, F., S. Jung, and D. R. Littman. 2003. Blood monocytes consist of
two principal subsets with distinct migratory properties. Immunity 19:71–82.
9. Gomez, R. M., J. E. Rinehart, R. Wollmann, and R. P. Roos. 1996. Theiler’s
murine encephalomyelitis virus-induced cardiac and skeletal muscle disease.
J. Virol. 70:8926–8933.
10. Griffin, D. E. 2001. Alphaviruses, p. 917–962. In P. M. Howley, P. M. Howley,
et al. (ed.), Fields virology, 4th ed. Lippincott, Williams, & Wilkins, Phila-
11. Griffin, D. E., and R. T. Johnson. 1977. Role of the immune response in
recovery from Sindbis virus encephalitis in mice. J. Immunol. 118:1070–1075.
12. Harley, D., D. Bossingham, D. M. Purdie, N. Pandeya, and A. C. Sleigh.
2002. Ross River virus disease in tropical Queensland: evolution of rheu-
matic manifestations in an inception cohort followed for six months. Med. J.
748 MORRISON ET AL. J. VIROL.
13. Harley, D., A. Sleigh, and S. Ritchie. 2001. Ross River virus transmission,
infection, and disease: a cross-disciplinary review. Clin. Microbiol. Rev. 14:
14. Hazelton, R. A., C. Hughes, and J. G. Aaskov. 1985. The inflammatory
response in the synovium of a patient with Ross River arbovirus infection.
Aust. N. Z. J. Med. 15:336–339.
15. Heise, M. T., D. A. Simpson, and R. E. Johnston. 2000. Sindbis-group
alphavirus replication in periosteum and endosteum of long bones in adult
mice. J. Virol. 74:9294–9299.
16. Heise, M. T., L. J. White, D. A. Simpson, C. Leonard, K. A. Bernard, R. B.
Meeker, and R. E. Johnston. 2003. An attenuating mutation in nsP1 of the
Sindbis-group virus S.A.AR86 accelerates nonstructural protein processing
and up-regulates viral 26S RNA synthesis. J. Virol. 77:1149–1156.
17. Kitamura, D., J. Roes, R. Kuhn, and K. Rajewsky. 1991. A B cell-deficient
mouse by targeted disruption of the membrane exon of the immunoglobulin
mu chain gene. Nature 350:423–426.
18. Kiwanuka, N., E. J. Sanders, E. B. Rwaguma, J. Kawamata, F. P. Ssengooba,
R. Najjemba, W. A. Were, M. Lamunu, G. Bagambisa, T. R. Burkot, L.
Dunster, J. J. Lutwama, D. A. Martin, C. B. Cropp, N. Karabatsos, R. S.
Lanciotti, T. F. Tsai, and G. L. Campbell. 1999. O’nyong-nyong fever in
south-central Uganda, 1996–1997: clinical features and validation of a clin-
ical case definition for surveillance purposes. Clin. Infect. Dis. 29:1243–1250.
19. Klapsing, P., J. D. MacLean, S. Glaze, K. L. McClean, M. A. Drebot, R. S.
Lanciotti, and G. L. Campbell. 2005. Ross River virus disease reemergence,
Fiji, 2003–2004. Emerg. Infect. Dis. 11:613–615.
20. Kuhn, R. J., H. G. Niesters, Z. Hong, and J. H. Strauss. 1991. Infectious
RNA transcripts from Ross River virus cDNA clones and the construction
and characterization of defined chimeras with Sindbis virus. Virology 182:
21. Laras, K., N. C. Sukri, R. P. Larasati, M. J. Bangs, R. Kosim, Djauzi, T.
Wandra, J. Master, H. Kosasih, S. Hartati, C. Beckett, E. R. Sedyaningsih,
H. J. Beecham III, and A. L. Corwin. 2005. Tracking the re-emergence of
epidemic chikungunya virus in Indonesia. Trans. R. Soc. Trop. Med. Hyg.
22. Lidbury, B. A., C. Simeonovic, G. E. Maxwell, I. D. Marshall, and A. J.
Hapel. 2000. Macrophage-induced muscle pathology results in morbidity and
mortality for Ross River virus-infected mice. J. Infect. Dis. 181:27–34.
23. MacDonald, G. H., and R. E. Johnston. 2000. Role of dendritic cell targeting
in Venezuelan equine encephalitis virus pathogenesis. J. Virol. 74:914–922.
24. Mombaerts, P., J. Iacomini, R. S. Johnson, K. Herrup, S. Tonegawa, and
V. E. Papaioannou. 1992. RAG-1-deficient mice have no mature B and T
lymphocytes. Cell 68:869–877.
25. Mordue, D. G., and L. D. Sibley. 2003. A novel population of Gr-1?-acti-
vated macrophages induced during acute toxoplasmosis. J. Leukoc. Biol.
26. Murphy, F. A., W. P. Taylor, C. A. Mims, and I. D. Marshall. 1973. Patho-
genesis of Ross River virus infection in mice. II. Muscle, heart, and brown fat
lesions. J. Infect. Dis. 127:129–138.
27. Pastorino, B., J. J. Muyembe-Tamfum, M. Bessaud, F. Tock, H. Tolou, J. P.
Durand, and C. N. Peyrefitte. 2004. Epidemic resurgence of Chikungunya
virus in democratic Republic of the Congo: identification of a new central
African strain. J. Med. Virol. 74:277–282.
28. Ray, C. G., L. L. Minnich, and P. C. Johnson. 1979. Selective polymyositis
inducted by coxsackievirus B1 in mice. J. Infect. Dis. 140:239–243.
29. Robben, P. M., M. Laregina, W. A. Kuziel, and L. D. Sibley. 2005. Recruit-
ment of Gr-1? monocytes is essential for control of acute toxoplasmosis. J.
Exp. Med. 201:1761–1769.
30. Rwaguma, E. B., J. J. Lutwama, S. D. Sempala, N. Kiwanuka, J. Kamugisha,
S. Okware, G. Bagambisa, R. Lanciotti, J. T. Roehrig, and D. J. Gubler.
1997. Emergence of epidemic O’nyong-nyong fever in southwestern Uganda,
after an absence of 35 years. Emerg. Infect. Dis. 3:77.
31. Seay, A. R., D. E. Griffin, and R. T. Johnson. 1981. Experimental viral
polymyositis: age dependency and immune responses to Ross River virus
infection in mice. Neurology 31:656–660.
32. Seay, A. R., and J. S. Wolinsky. 1982. Ross River virus-induced demyelina-
tion: I. Pathogenesis and histopathology. Ann. Neurol. 12:380–389.
33. Shinkai, Y., G. Rathbun, K. P. Lam, E. M. Oltz, V. Stewart, M. Mendelsohn,
J. Charron, M. Datta, F. Young, A. M. Stall, et al. 1992. RAG-2-deficient
mice lack mature lymphocytes owing to inability to initiate V(D)J. rearrange-
ment. Cell 68:855–867.
34. Simpson, D. A., N. L. Davis, S. C. Lin, D. Russell, and R. E. Johnston. 1996.
Complete nucleotide sequence and full-length cDNA clone of S.A.AR86 a
South African alphavirus related to Sindbis. Virology 222:464–469.
35. Soden, M., H. Vasudevan, B. Roberts, R. Coelen, G. Hamlin, S. Vasudevan,
and J. La Brooy. 2000. Detection of viral ribonucleic acid and histologic
analysis of inflamed synovium in Ross River virus infection. Arthritis
36. Suhrbier, A., and M. La Linn. 2004. Clinical and pathologic aspects of
arthritis due to Ross River virus and other alphaviruses. Curr. Opin. Rheu-
37. Suthar, M. S., R. Shabman, K. Madric, C. Lambeth, and M. T. Heise. 2005.
Identification of adult mouse neurovirulence determinants of the Sindbis
virus strain AR86. J. Virol. 79:4219–4228.
38. Tidball, J. G. 2005. Inflammatory processes in muscle injury and repair.
Am. J. Physiol. Regul. Integr. Comp. Physiol. 288:R345–R353.
39. Williams, M. C., J. P. Woodall, and J. D. Gillett. 1965. O’nyong-Nyong fever:
an epidemic virus disease in East Africa. VII. Virus isolations from man and
serological studies up to July 1961. Trans. R. Soc. Trop. Med. Hyg. 59:186–
40. Yoneyama, M., M. Kikuchi, T. Natsukawa, N. Shinobu, T. Imaizumi, M.
Miyagishi, K. Taira, S. Akira, and T. Fujita. 2004. The RNA helicase RIG-I
has an essential function in double-stranded RNA-induced innate antiviral
responses. Nat. Immunol. 5:730–737.
41. Ytterberg, S. R., M. L. Mahowald, and R. P. Messner. 1987. Coxsackievirus
B 1-induced polymyositis. Lack of disease expression in nu/nu mice. J. Clin.
42. Zare, F., M. Bokarewa, N. Nenonen, T. Bergstrom, L. Alexopoulou, R. A.
Flavell, and A. Tarkowski. 2004. Arthritogenic properties of double-stranded
(viral) RNA. J. Immunol. 172:5656–5663.
VOL. 80, 2006ROSS RIVER VIRUS TROPISM AND INFLAMMATORY DISEASE749