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Although xenotropic murine leukemia virus-related virus (XMRV) has been previously linked to prostate cancer and myalgic encephalomyelitis/chronic fatigue syndrome, recent data indicate that results interpreted as evidence of human XMRV infection reflect laboratory contamination rather than authentic in vivo infection. Nevertheless, XMRV is a retrovirus of undefined pathogenic potential that is able to replicate in human cells. Here we describe a comprehensive analysis of two male pigtailed macaques (Macaca nemestrina) experimentally infected with XMRV. Following intravenous inoculation with >1010 RNA copy equivalents of XMRV, viral replication was limited and transient, peaking at ≤2,200 viral RNA (vRNA) copies/ml plasma and becoming undetectable by 4 weeks postinfection, though viral DNA (vDNA) in peripheral blood mononuclear cells remained detectable through 119 days of follow-up. Similarly, vRNA was not detectable in lymph nodes by in situ hybridization despite detectable vDNA. Sequencing of cell-associated vDNA revealed extensive G-to-A hypermutation, suggestive of APOBEC-mediated viral restriction. Consistent with limited viral replication, we found transient upregulation of type I interferon responses that returned to baseline by 2 weeks postinfection, no detectable cellular immune responses, and limited or no spread to prostate tissue. Antibody responses, including neutralizing antibodies, however, were detectable by 2 weeks postinfection and maintained throughout the study. Both animals were healthy for the duration of follow-up. These findings indicate that XMRV replication and spread were limited in pigtailed macaques, predominantly by APOBEC-mediated hypermutation. Given that human APOBEC proteins restrict XMRV infection in vitro, human XMRV infection, if it occurred, would be expected to be characterized by similarly limited viral replication and spread.
In 2006, Urisman and coworkers identified sequences from a novel gammaretrovirus in an analysis of human prostate tumor tissues using a Virochip DNA microarray and named the new virus xenotropic murine leukemia virus (X-MLV)-related virus (XMRV) due to its high sequence identity with X-MLVs (64). Although several follow up studies described findings interpreted as evidence of XMRV infection in up to 27% of prostate cancer patients and up to 4% of healthy prostate controls using PCR (3, 9, 11, 54), immunohistochemistry (IHC) (54), and fluorescence in situ hybridization (ISH) (3) on prostate tissues, as well as anti-XMRV serum neutralization assays (3), the majority of studies detected little or no evidence of XMRV infection in either prostatic tumors or healthy controls, raising questions about the authenticity of human XMRV infection and what role, if any, XMRV might play in prostate cancer (1, 2, 15, 23, 50, 56, 61).
Following the initial suggestion of an association with prostate cancer, XMRV was also implicated as a potential factor in myalgic encephalomyelitis/chronic fatigue syndrome (ME/CFS) based on a study by Lombardi and coworkers that reported evidence of XMRV infection in nearly 70% of ME/CFS patients, compared with <4% of healthy controls from the United States, using PCR, serological testing, and virus isolation (34). Although a subsequent study by a different lab identified MLV-related sequences in 86.5% of ME/CFS patient and in 6.8% of healthy controls, all but one of the viral sequences were distinct from XMRV and X-MLV and were instead more closely related to polytropic and modified polytropic MLVs (27, 33, 63). The preponderance of subsequent analyses, however, reported no evidence of XMRV infection in samples from patients with ME/CFS in the United States (22, 29, 53, 55, 60), the United Kingdom (13, 18), Germany (24), The Netherlands (65), or China (25), and reexaminations of samples from patients previously identified as XMRV positive in the original Lombardi et al. publication found no consistent evidence of XMRV infection (29, 58). In addition, though XMRV infection has been proposed as a possible contributor to many other human diseases and disorders, efforts to detect the virus in samples from people with systemic lupus erythematosus (4), fibromyalgia (36), multiple sclerosis (24), amyotrophic lateral sclerosis (38), or autism (32, 52) have thus far yielded negative results, and the virus has not been found in HIV-1-positive or immunosuppressed individuals or in people at high risk of infection with blood-borne pathogens (5, 8, 10, 17, 30, 35, 37, 62).
Several possible explanations have been proposed for the noncongruent detection of XMRV by different laboratories, including potential differences in the geographical distribution of XMRV, the use of different assay methodologies and techniques, and the use of different patient selection criteria. An alternative explanation emerged, however, when it was shown that XMRV sequences identified in human samples formed a monophyletic clade with XMRV produced from the 22Rv1 cell line and lacked the diversity typical of circulating retroviruses (26). This finding underscored uncertainties about the replicative capacity and sequence evolution of XMRV in vivo while raising the possibility that sample or reagent contamination was responsible for the identification of XMRV in human samples. Indeed, several studies subsequently showed that mouse DNA and MLV sequences contaminate a number of commonly used PCR and nucleic acid extraction reagents (12, 42, 49, 51, 63), and a recent partial retraction of the Lombardi et al. report has indicated XMRV plasmid DNA contamination in some of the ME/CFS patient samples in that study (57). Moreover, a recent study has shown that XMRV almost certainly originated in the lab during the derivation of the 22Rv1 prostate cancer cell line, during which a recombination event between two different MLVs was facilitated by serial passage of a human prostate tumor xenograft in nude mice, indicating that widespread human XMRV infection is highly unlikely (46).
Although there is a growing consensus that evidence of XMRV infection in human samples is more likely the result of contamination than genuine in vivo infection, the fact remains that XMRV is a novel, replication-competent retrovirus of unknown pathogenic potential once implicated in the etiology of several human diseases. Given uncertainties about the capacity of XMRV to cause human disease, as well as the potential target cell tropism, tissue distribution, in vivo replication capacity, and sequence evolution of the virus and elicited antiviral immune responses, we infected two pigtailed macaques (Macaca nemestrina) with a well-characterized 22Rv1-produced XMRV stock. We conducted these studies with two primary goals. First, we sought to generate bona fide in vivo-derived positive-control samples for PCR, reverse transcriptase PCR (RT-PCR), and serology assays performed on clinical samples (10, 28). Second, we aimed to comprehensively examine the natural history of in vivo XMRV infection in a primate host, evaluating levels and kinetics of replication, viral sequence changes, cell and tissue tropism, and cellular and humoral antiviral immune responses. We show here that XMRV replication in pigtailed macaques is restricted, with limited, transient viremia associated with the accumulation of extensive G-to-A hypermutation in cell-associated viral DNA (vDNA). In spite of limited viral replication, humoral immune responses to the virus were relatively robust and stable, while innate immune responses were transient and adaptive cellular immune responses were negligible.
The human prostate carcinoma cell line 22Rv1 (59) (American Type Culture Collection catalog number CRL-2505) and the human T lymphoblastoid cell lines SupT1-CCR5 (39), Jurkat-TAT-CCR5 (41), and H9 were cultured in Roswell Park Memorial Institute (RPMI) 1640 medium supplemented with 10% fetal bovine serum (FBS), 2 mM l-glutamine, and 100 U/ml penicillin–100 μg/ml streptomycin. DERSE.LiG-puro indicator cells (K. Lee, unpublished data) were maintained in RPMI 1640 medium supplemented with 15% FBS, 2 mM l-glutamine, 100 U/ml penicillin–100 μg/ml streptomycin, and 1 μg/ml puromycin (Calbiochem). Human 293T cells were cultured in Dulbecco's modified Eagle's medium supplemented with 10% FBS, 2 mM l-glutamine, and 100 U/ml penicillin–100 μg/ml streptomycin.
For neutralization studies, XMRV clone VP62 and VSV-G-pseudotyped Moloney MLV (Mo-MLV) were produced in 293T cells transiently transfected with pVP62 (kindly provided by Robert Silverman ) or cotransfected with pMX-RFP, pJK3, pCMV-tat, and pL-VSV-G (for long terminal repeat [LTR]-driven expression of VSV-G) at a 4:2:0.3:1 ratio (6). The pMX-RFP vector is derived from Mo-MLV (43) and has been modified through insertion of the dsRed gene into the multicloning site (31). Virus-containing supernatants were collected 2 days posttransfection, clarified by centrifugation, filtered through a 0.45-μm filter (Millipore), aliquoted, and stored at −80°C. For all other analyses, including inoculation into macaques, XMRV was produced in 22Rv1 cells. Virus-containing 22Rv1 culture supernatants were harvested 3 days after 100% culture medium change (6 days after cell line passage), clarified by centrifugation at 4,000 × g for 15 min, filtered using a 0.45-μm filter (Millipore), aliquoted, and stored in liquid nitrogen vapor. To purify and concentrate XMRV for protein analysis, 30 ml of filtered 22Rv1 culture supernatant was first centrifuged through 20% sucrose in TNE (0.01 M Tris-HCl [pH 7.2], 0.1 M NaCl, 1 mM EDTA) using a Beckman SW32 rotor at 25,000 rpm (~100,000 × g, maximum) for 1 h at 4°C. The resultant virus pellet was washed by resuspension in phosphate-buffered saline (PBS) and centrifugation in a Beckman TLA100.3 rotor at 60,000 rpm (~200,000 × g, maximum) for 6 min at 4°C, and the washed pellet was resuspended in 30 μl PBS for a final concentration factor of 1,000×.
Two adult male pigtailed macaques (M. nemestrina) were each intravenously inoculated (saphenous vein) with 2 ml of infectious XMRV generated in 22Rv1 cells. The animals were 9 to 10 years old, weighed 13 to 14 kg, and were free of cercopithicine herpesvirus 1, D-type simian retrovirus, simian T-lymphotrophic virus type 1, and simian immunodeficiency virus (SIV) at the beginning of the study. Plasma for virus neutralization, virus rescue, and viral RNA (vRNA) quantification, and sequencing analyses and peripheral blood mononuclear cells (PBMC) for virus rescue, cell-associated DNA quantification, and flow cytometry assays were prepared from blood collected in EDTA Vacutainer tubes (BD). PBMC were isolated from whole EDTA blood by Ficoll-Paque Plus (GE Healthcare) gradient centrifugation. Sera for immunoblot assays were prepared from blood collected in Serum Vacutainer tubes (BD). Lymph node (LN) mononuclear cells (LNMC) and splenocytes for cell-associated vDNA quantification were prepared by mincing tissues in RPMI medium supplemented with 10% FBS and then passing the cell-medium slurry through a 70-μm cell strainer (BD). Whole prostate tissue pieces for cell-associated vDNA quantification were collected at necropsy, immediately snap-frozen in liquid nitrogen, and stored at −80°C. For ISH and IHC, prostate and LN tissues were immediately fixed in paraformaldehyde (PFA). The animals were housed and cared for in accordance with American Association for Accreditation of Laboratory Animal Care (AAALAC) guidelines in an AAALAC-accredited facility, and all animal procedures were performed according to protocols approved by the Institutional Animal Care and Use Committee of the National Cancer Institute.
Equal volumes of purified XMRV sample and 4% glutaraldehyde was centrifuged at 100,000 × g for 1 h at 4°C. The virus pellet were postfixed in 1% osmium tetroxide in cacodylate buffer (0.1 M, pH 7.2) for 1 h at room temperature. The pellet was washed in the same buffer and then in acetate buffer (0.1 M, pH 4.2) and stained in uranyl acetate (0.5% aqueous solution) for 1 h. The pellet was then washed in acetate buffer and dehydrated in ethanol solutions (e.g., 35%, 50%, 75%, 95%, and 100%), followed by 100% propylene oxide. The pellet was infiltrated in an equal volume of epoxy resin and 100% propylene oxide for 18 h at room temperature, embedded in fresh resin, and then cured at 55°C for 48 h. Thin sections (80 to 90 nm) were made and mounted on a naked copper grid, stained with uranyl acetate and lead citrate, and examined in an electron microscope (Hitachi 7600) operated at 80 kV, and images were taken with a digital camera (AMT).
Viral samples were disrupted in 8 M guanidine-HCl (Pierce) with or without 50 mM dithiothreitol (Calbiochem) and fractionated by high-performance liquid chromatography (HPLC) to isolate viral proteins. HPLC was performed at a flow rate of 300 μl/min on a Poros R2/H narrow-bore column (2.1 by 100 mm; Boehringer Mannheim GmbH) using aqueous acetonitrile-trifluoroacetic acid solvents and a Shimadzu HPLC system equipped with LC-10AD pumps, an SCL-10A system controller, a CTO-10AC oven, an FRC-10A fraction collector, and an SPD-M10AV diode array detector. The gradient of buffer B (0.1% trifluoroacetic acid in acetonitrile) was 10% to 36.5% for 8 min; 36.5% for 12 min; 36.5% to 60% for 10 min; 60% to 80% for 5 min; and 80% for 5 min. A temperature of 55°C was maintained during HPLC separation. Peaks were detected by UV absorption at 206 and 280 nm and analyzed by sequencing using an automated Applied Biosystems Inc. 477 Protein Sequencer, by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), and by immunoblot analysis using Millipore Luminata Forte Western HRP Substrate (catalog number WBLUF0500). Quantification of total viral and purified proteins was performed by amino acid analysis using a Hitachi L-8800 amino acid analyzer.
Viral samples were loaded into a 4 to 20% gradient gel, which was calibrated with a preparation of p30 purified from Mo-MLV by HPLC and quantified by amino acid analysis. After staining with Coomassie blue, densitometry was performed and the p30 in the virus was quantified by interpolation onto standard curves produced from scanning known input amounts of p30 on the same gel. A Scan Ace III from Pacific Image Electronics was used to digitally scan the data in grayscale transparency mode from an immunoblot assay film in TIFF format. After scanning of gels, band intensities were quantified using Nonlinear Dynamics TotalLab v 2.0. No image enhancement options were used that might have altered the relative intensity ratios of the bands being analyzed. Care was taken to use series of dilutions of sample which gave band intensities within the nonsaturated, linear response range of the optical hardware. For Western blot analyses, 100 ng of p30 was loaded per lane of a 4 to 20% gradient gel and, after separation, proteins were transferred onto a polyvinylidene difluoride (PVDF) membrane (Immobilon-P IPVH00010; Millipore) using a semidry electroblotter (Ellard Instrumentation) for 1 to 1.5 h. Detection of p30 and gp70 was performed using in-house rabbit polyclonal antibodies DJ-39462 and DJ-3946, respectively, each used at a 1:10,000 dilution, followed by a goat anti-rabbit secondary antibody (Sigma catalog number A6154) at 1:10,000.
Lysed whole XMRV, HPLC-fractionated XMRV, or microvesicles from human cell lines were loaded onto a 4 to 20% gradient gel, and after separation, proteins were transferred onto a PVDF membrane (Immobilon-P IPVH00010; Millipore) using a semidry electroblotter (Ellard Instrumentation) for 1 to 1.5 h. Membranes were probed with macaque serum, followed by a goat anti-human secondary antibody (Sigma catalog number A8667) diluted 1:10,000. Microvesicles were prepared from the human cell lines H9, SupT1-CCR5, and Jurkat-TAT-CCR5 following the same protocol described for the purification of XMRV from 22Rv1 cells.
Pigtailed macaque (M. nemestrina) PBMC were isolated from whole blood by Ficoll-Paque Plus (GE Healthcare) gradient centrifugation, stimulated with plate-bound anti-CD3 (BD), and cultured in 100 IU/ml interleukin-2 (IL-2; BD) for 3 days prior to infection. We inoculated 6 × 106 PBMC with 1010, 109, 108, or 106 RNA copy equivalents of XMRV and incubated them for 6 h at 37°C. The cells were then washed 3 times in PBS and subsequently maintained in 6 ml RPMI-complete medium with 100 IU/ml IL-2 (BD). Cell-free supernatants were collected at 1, 3, 6, 9, 12, and 15 days postinfection (dpi) from each culture and stored at −80°C for later quantification of virus by quantitative RT-PCR (qRT-PCR) (see below). Cells collected at day 15 were washed once in PBS, resuspended at 107/ml in PBS, and spotted onto glass slides (Fisher) at 20 μl/spot for ISH.
XMRV RNA was extracted from in vitro-infected cell supernatants and quantified using qRT-PCR as described previously (7). For in vivo-derived samples, XMRV RNA in macaque plasma and XMRV DNA in whole PBMC, LNMC, and splenocytes were extracted and quantified using a single-copy qRT-PCR and quantitative PCR (qPCR) assays (X-SCA) as described previously (28). Nucleic acid was extracted from snap-frozen prostate tissue pieces harvested at necropsy as described previously (21), and XMRV DNA was quantified by X-SCA.
vRNA from macaque plasma and vDNA from macaque PBMC and splenocytes were sequenced using XMRV single-genome sequencing (X-SGS) as described elsewhere (28a). Briefly, to obtain PCR products amplified from a single viral genome, extracted DNA or cDNA generated by reverse transcription of vRNA was serially diluted to achieve a dilution that yields products from ~30% of subsequent PCRs. The primers used generated ~1.4-kb sequences in XMRV gag.
Antibodies were from BD Biosciences unless indicated otherwise. All data were acquired on an LSR-II cytometer (BD Immunocytometry Systems), and subsequent data analysis was performed using FCS Express, version 4.0 (De Novo Software). Absolute cell counts were determined from whole blood using BD TruCOUNT absolute cell counting tubes (BD Biosciences) according to the manufacturer's recommendations. Briefly, 50 μl whole blood was added to TruCOUNT tubes containing a panel of the following antibodies: CD45-fluorescein isothiocyanate (FITC) (clone DO58-1283), CD3-phycoerythrin (PE) (clone SP34-2), CD4-allophycocyanin (APC) (clone L200), CD14-APC-Cy7 (clone M5E2; BioLegend), CD8α–PE-Cy7 (clone SK1), and CD20 Pacific Blue (clone 2H7; BioLegend). Following a 20-min incubation at room temperature in the dark, red blood cells were lysed with FACS Lysing Solution (BD Biosciences) and 10,000 gated bead events were acquired for each sample. For immunophenotyping, freshly isolated cells were surface labeled with a panel of the following antibodies: CD4 Pacific Blue (clone OKT4; BioLegend), CCR5-PE (clone 3A9), CD28 ECD (clone CD28.2; Beckman Coulter), CD95–PE-Cy5 (clone DX2), CD8α–PE-Cy7, CD38-APC (clone OK10; NIH Nonhuman Primate Reagent Resource), and CD3–APC-Cy7. After surface staining, cells were fixed, permeabilized (BD Cytofix/Cytoperm Kit), and labeled with Ki67-FITC (clone B56). Approximately 200,000 gated CD3+ T cells were acquired for each sample. For intracellular cytokine staining (ICS), viably cryopreserved PBMC were quick-thawed and stimulated with whole purified XMRV/22Rv1 and purified recombinant XMRV proteins for CA, MA, p12, NC, PR, RT, IN, and TM in the presence of 1 μg/ml anti-CD49d and anti-CD28 costimulatory antibodies for 9 h at 37°C with 7% CO2. Medium alone and 10 μg/ml tetanus toxoid protein (List Biological Laboratories, Inc.) were used as negative controls, and 100 ng/ml phorbol myristate acetate with 1 μM ionomycin (Sigma) was used as a positive control. After 1 h of culture initiation, 10 μg/ml of brefeldin A (Sigma) with 0.14 μl/well of GolgiStop (monensin; BD Biosciences) were added to all wells. Cells were then surface stained with LIVE/DEAD Fixable Yellow Dead Cell Stain (Invitrogen; to exclude dead cells), CD4 Pacific Blue (OKT4; BioLegend), and CD8–PerCP-Cy5.5 (SK1; BD Biosciences). Cells were then fixed, permeabilized (BD Cytofix/Cytoperm Kit), and intracellularly stained with CD3–APC-Cy7 (SP34-2), gamma interferon (IFN-γ)–FITC (B27), MIP-1b–PE (D21-1351), tumor necrosis factor alpha–PE-Cy7 (MAb11), and IL-2–APC (MQ1-17H12). Approximately 200,000 viable CD3+ T cells were acquired for each sample.
Digoxigenin-labeled XMRV riboprobes were developed for ISH using 5′XMRV and 3′XMRV plasmids (47). XMRV riboprobes were generated by PCR-based cloning of target regions in gag, pro, pol, and env (4 riboprobes) of roughly equal sizes (~600 bp; allowing for equal molar stoichiometric representation in the riboprobe cocktail) using primers with phage T7 (antisense) promoter sequences cloned upstream of the viral sequence (see Table S1 in the supplemental material). XMRV ISH was performed using previously published lentiviral ISH procedures (20, 45). In brief, 5-μm tissue sections were dewaxed in xylenes, rehydrated through graded ethanol, and placed in HyPure molecular biology grade H2O (HyClone; Thermo Scientific). Cell spots were fixed in 4% PFA for 20 min and placed in HyPure molecular biology grade H2O. All slides were immersed in 0.2 N HCI for 30 min, 0.15 M triethanolamine (pH 7.4) for 15 min, and 0.005% digitonin for 5 min, all at room temperature. The slides were then incubated for 15 to 30 min at 37°C in a Tris-buffered solution containing 2 mM CaCI2 and proteinase K (5 μg/ml). After the slides were removed and washed in HyPure molecular biology grade H2O, they were acetylated (0.25% acetic anhydride) for 20 min and then placed in 0.1 M triethanolamine (pH 8.0) until hybridization. The sections were then covered with hybridization solution (50% deionized formamide, 10% dextran sulfate, 0.6 M NaCl, 0.4 mg/ml yeast RNA [Ambion, Inc.], and 1× Denhardt's medium in 20 mM HEPES buffer [pH 7.2] with 1 mM EDTA) containing 0.4 ng/μl pooled XMRV riboprobes and hybridized for 18 h at 48°C. After hybridization, slides were washed and developed as previously described (45).
Immunohistochemistry was performed using a biotin-free polymer approach (mouse and rabbit Polink-2; Golden Bridge International, Inc.) as previously described (14). Representative images (magnifications, ×200 and ×400) were acquired using a Nikon 80i bright-field microscope. The primary antibody used was a mouse monoclonal anti-myxovirus resistance protein A antibody (MxA; clone M143 from Otto Haller and Georg Kochs, Department of Virology, Universitätsklinikum Freiburg, Freiburg, Germany).
Virus rescue assays were performed using DERSE.LiG-puro cells under optimized conditions (28). DERSE.LiG-puro cells are a subclone of the LNCaP human prostate cancer cell line that has been stably transfected with an intron-interrupted green fluorescent protein (GFP) reporter construct flanked by MLV LTRs that will be packaged into XMRV particles and expressed upon the second and subsequent rounds of replication, allowing the detection of infectious XMRV. Briefly, DERSE.LiG-puro cells were plated at 1 × 105/well in 24-well tissue culture plates 1 day before infection. On the next day, macaque plasma samples were diluted in RPMI medium containing CaCl2 and heparin and then applied to DERSE.LiG-puro cells. As a positive control, 22Rv1-produced XMRV was spiked into preinfection plasma samples and added to DERSE.LiG-puro cells. At 1 dpi, plasma was removed and replaced with fresh medium. After being maintained for 18 days, cells were collected and resuspended in a 2% PFA solution for the measurement of GFP-positive cells by FACS analysis (FACSCalibur; BD). To rescue virus from different PBMC subsets, whole PBMC were stimulated for 2 to 3 days with lipopolysaccharide, with anti-CD3 antibody, or by culturing on a monolayer of irradiated CD40-L-expressing NIH 3T3 cells and then cocultured with DERSE.LiG-puro cells.
Plasma was heat inactivated for 30 min at 55°C, serially diluted, and added to XMRV clone VP62, 22Rv1-produced XMRV, or VSV-G-pseudotyped MLV-containing samples. These virus samples were then applied to DERSE.LiG-puro cells seeded at 1 × 105/well in 24-well tissue culture plates 1 day earlier. After 10 h, virus-plasma inocula were replaced with fresh medium. At 4 dpi, cells were collected in a 2% PFA solution and GFP-positive cells were measured by FACS analysis (FACSCalibur; BD).
Prior to initiating our XMRV infection studies, we first generated and characterized an infectious XMRV stock from the human prostate carcinoma line 22Rv1. This cell line, the derivation of which was recently implicated as the original source of XMRV (46), constitutively releases infectious XMRV into culture supernatants. To evaluate the composition and concentration of 22Rv1-derived XMRV, a portion of the infectious virus stock was purified, concentrated 1,000×, and examined by SDS-PAGE (Fig. 1A). Coomassie staining revealed a limited number of protein bands in the purified XMRV material, all of which approximately corresponded in size to the protein constituents of similarly purified and concentrated Mo-MLV, suggestive of a virus stock relatively free of contaminating host cell proteins. The viral identities of several of the protein bands were verified using antisera raised against the Mo-MLV p30CA and gp70SU proteins (Fig. 1C). In addition, purified, concentrated XMRV was fractionated by HPLC and polypeptide microsequencing was performed on the fractions, demonstrating XMRV sequences as the constituents of each major peak (see Fig. 5B and C). To determine the virion content of our 22Rv1-derived XMRV stock, we first determined the concentration of CA protein by generating a densitometry standard curve using a dilution series of purified Mo-MLV p30CA of known concentrations (Fig. 1A). Using linear regression analysis (Fig. 1B), we calculated a concentration of 2.6 mg/ml CA protein in 1,000×-concentrated XMRV or 2.6 μg/ml p30CA in the 1× infectious virus stock. Using real-time qRT-PCR, we also determined the viral genome copy number in our 1× infectious virus stock to be 2.4 × 1010 RNA copies/ml, consistent with prior data suggesting a rough rule of thumb of approximately 1010 viral particles/μg of CA protein for other retroviruses (48). Transmission electron microscopy of pelleted, 22Rv1-derived material verified the presence of highly abundant and highly homogeneous virions, with a mean diameter of approximately110 nm (Fig. 1D). Our further work on XMRV was conducted using this well-characterized stock of virus generated from 22Rv1 cells.
Prior to infecting pigtailed macaques (M. nemestrina) with XMRV, we first sought to determine if pigtailed macaque cells could support XMRV replication (defined as at least a single round whereby a de novo infection event leads to the production of progeny virions). In addition to human prostate cancer cell lines, XMRV has been shown to replicate in human T cell lines and primary human PBMC (7, 47). We therefore evaluated XMRV replication in pigtailed macaque PBMC in vitro. Stimulated pigtailed macaque PBMC were inoculated with various input amounts of 22Rv1-derived XMRV (1010, 109, 108, or 106 RNA copy equivalents) and monitored by real-time qRT-PCR for vRNA output in the culture supernatants (Fig. 2A). Although the virus replicated in the PBMC cultures, showing increases in vRNA levels between 1 dpi and peak viral output at 3 to 6 dpi, we observed a plateau effect, with maximum virus production levels dependent upon the size of the input inoculum, suggestive of restricted viral replication similar to results previously described for XMRV replication in human PBMC (7). In the 1010, 109, and 108 RNA copy input cultures, viral replication peaked between days 3 and 6 and was maintained through 15 dpi (Fig. 2A), with no apparent cytopathicity up to 15 dpi. Virus was not detected in the 106 RNA copy input culture (<102 RNA copies/8.4 μl). To confirm the presence of productively infected cells in the PBMC cultures, we performed ISH on day 15 cells using a cocktail of XMRV riboprobes. We identified vRNA-positive cells in the infected culture, confirming the presence of productively infected cells (Fig. 2B). Importantly, our ISH protocol detected RNA-positive cells only in the infected cell culture and only in the presence of the XMRV riboprobe cocktail, demonstrating the specificity of our ISH riboprobes and the absence of nonspecific reactivity with our secondary detection antibody. Although XMRV replication appeared to be somewhat restricted in pigtailed macaque PBMC, these results indicate that XMRV is clearly infectious for pigtailed macaque cells.
To evaluate the in vivo replication and evolution of XMRV, as well as the elicited antiviral immune responses in primates, we intravenously inoculated two adult male pigtailed macaques (animal designations 8242 and 14232) with 4.8 × 1010 RNA copy equivalents of XMRV and monitored infection for 119 days. Both animals were healthy throughout the course of the study with no clinical indications of disease. Viral replication was longitudinally monitored using X-SCA to quantify vRNA in plasma and DNA in cells and tissues, including PBMC and LNMC (Fig. 3). In both animals, XMRV replicated transiently, with peak plasma viral loads (VLs) of 2,200 (Fig. 3A) and 530 (Fig. 3B) RNA copies/ml measured at 13 and 5 dpi, respectively. Following peak viremia, plasma VLs rapidly declined to undetectable levels (<1.1 RNA copies/ml) ~2 weeks postpeak. Except for a single small blip (7 copies/ml) detected at 63 dpi in the animal with higher peak VLs (animal 8242), plasma viremia remained undetectable in both animals throughout the subsequent duration of follow-up. vDNA levels in PBMC also peaked within the first 2 weeks of infection but remained at detectable levels in both animals throughout the experiment, maintaining a relatively stable pool of 300 to 700 DNA copies/106 PBMC in animal 8242 (Fig. 3A), while showing a gradual ~12-fold decline from 270 to 23 copies/106 PBMC between 13 and 119 dpi in animal 14232 (Fig. 3B). The initial appearance of vDNA in peripheral LNMC was slightly delayed with respect to plasma RNA and PBMC DNA but peaked with kinetics similar to those seen in PBMC. Although peripheral LNs could not be obtained from animal 14232 at later time points, peripheral LNMC from animal 8242 remained vDNA positive with 190 DNA copies/106 cells at the study's conclusion (Fig. 3A).
At 119 dpi, both infected animals were euthanized and necropsies were performed with extensive tissue sampling, including the collection of additional LN, spleen, and prostate tissues. Of the lymphoid organs tested over the course of this study, the highest levels of vDNA were detected in the spleen, with 1,700 (Fig. 3A) and 280 (Fig. 3B) DNA copies/106 cells for 8242 and 14232, respectively. While comparable levels of vDNA were measured in bronchial LNMC (Bron-LN) in both animals (150 to 250 DNA copies/106 cells), vDNA was detected in mesenteric LNMC (Mes-LN) only from animal 8242 (Fig. 3A). Because the prostate has been previously implicated as a possible target organ of XMRV infection, we extracted DNA from whole prostate tissue pieces and quantified vDNA by X-SCA. Detectable but very low levels of vDNA were measured in prostate tissue from both animals, with 14 (Fig. 3A) and 5 (Fig. 3B) copies/106 cell equivalents detected in animals 8242 and 14232, respectively. Despite the presence of XMRV DNA in LN and prostate tissues, we did not detect any XMRV RNA-positive cells in either tissue type using our ISH protocol (see Fig. S1 in the supplemental material; data not shown), which detected XMRV RNA in in vitro-infected pigtailed macaque PBMC with good specificity (Fig. 2B).
Although replication-competent virus could be cultured from preinfection plasma samples spiked with infectious XMRV using DERSE.LiG-puro indicator cells, virus could not be successfully rescued from plasma isolated at multiple postinfection time points, including peak viremia (see Fig. S2 in the supplemental material), or from differentially stimulated PBMC (see Materials and Methods) isolated at necropsy (data not shown).
Given questions about the apparent lack of diversity of XMRV sequences identified in human samples (26), and because XMRV plasma viremia in our infected monkeys was only transient despite the continued presence of vDNA-positive cells in the blood, LNs, and spleen (Fig. 3), we used a limiting dilution single-genome sequencing method (X-SGS) (28a) to sequence vRNA in plasma and vDNA in PBMC to assess sequence diversification and evolution (Fig. 4). For animal 8242, plasma RNA was sequenced at days 5, 13, and 19, representing ramp-up, peak, and postpeak viremia, respectively. For animal 14232, plasma viremia peaked earlier and resolved more rapidly (Fig. 3), so plasma RNA sequences could be obtained only from day 5 samples. While day 5 RNA from both animals showed the acquisition of random point mutations in multiple viral clones, 36.8% (7/19) of the sequences from 8242 were 100% identical to each other (Fig. 4A), whereas 75.0% (12/16) of the sequences from 14232 were 100% identical to each other (Fig. 4B), consistent with the higher levels of replication and associated mutations in animal 8242. Day 13 and day 19 RNA sequences from 8242 showed similar levels of diversity, with 42.9% (9/21) and 25.0% (1/4) identical sequences, respectively (Fig. 4A). Using longitudinal RNA sequences from animal 8242, we generated a neighbor-joining phylogeny rooted on the predominant 22Rv1-produced inoculum sequence. As shown in Fig. 4C, the virus acquired a limited number of random mutations, with no apparent selection in the sequenced region over the first 19 days of infection. The acquisition of a small number of mutations in plasma RNA was consistent with transient, low-level viral replication.
Although early plasma RNA sequences from both animals showed the acquisition of point mutations consistent with productive replication, plasma RNA fell to undetectable levels in both animals despite the persistent presence of XMRV DNA-positive cells in blood and LNs. We therefore performed X-SGS on DNA extracted from PBMC at days 13 and 40, representing viremic and aviremic time points, respectively, for both animals (Fig. 4A and B). X-SGS of vDNA revealed extensive G-to-A hypermutation in samples derived from both animals, consistent with APOBEC-mediated hypermutation. Although multiple hypermutated sequences were identified at both time points, the proportion of sequences containing multiple G-to-A mutations increased from the viremic to the aviremic time points, from ~38% to ~83% in animal 8242 and from ~59% to ~78% in animal 14232, indicating a loss of cells containing nonhypermutated viral genomes and/or an increase in the number of hypermutated genomes over time. These results likely underestimate the proportion of genomes that are hypermutated due to inefficient priming and amplification of hypermutated sequences relative to nonhypermutated sequences. Similar results were obtained from sequence analysis of splenocyte-associated vDNA at necropsy, with 16/18 sequences containing ≥10 G-to-A mutations and 18/18 sequences containing at least 1 G-to-A mutation for animal 8242 (data not shown).
Studies assessing the prevalence of XMRV have frequently utilized serological analyses to identify potential human XMRV infection (3, 13, 16, 18, 23, 24, 29, 34, 40, 50, 52, 53, 60, 61); however, the potential magnitude and kinetics of antibody responses to XMRV infection are not known. To evaluate the kinetics of antibody responses elicited in our XMRV-infected pigtailed macaques, we performed immunoblot analyses using longitudinal serum samples to probe blots containing whole, lysed XMRV (Fig. 5A). Importantly, preinfection sera did not show background reactivity with XMRV, indicating that any reactivity in subsequent samples reflected specificity for XMRV infection. For both animals, anti-XMRV reactivity was first detected at 2 weeks postinfection, with both animals showing reactivity to the viral envelope TM protein p15E. Serum obtained at 2 weeks postinfection from animal 8242 also detected the SU subunit of envelope, gp70, whereas serum from animal 14232 did not bind gp70 but instead detected p30CA. Peak antibody response breadth was first detected at 4 weeks postinfection, with serum samples from both animals showing reactivity with p15E, p30CA, gp70SU, and p15MA (Fig. 5C). Anti-XMRV antibody responses were relatively stable, remaining detectable in serum at necropsy at 17 weeks postinfection, despite undetectable plasma VLs for up to 100 days. Although antibody reactivity had started to wane at this time point, p15E and p30CA antibodies were still detectable by immunoblot assay at a 1:2,000 serum dilution. Sera were obtained by terminal blood collection at euthanasia and stored for future use as positive-control reagents.
To confirm the identities of viral proteins detected by the macaque antisera, we HPLC fractionated 22Rv1-derived XMRV and determined the identities of the constituent proteins of each peak by amino acid sequencing (Fig. 5B). Fifty percent of the total protein from each fraction was run on a polyacrylamide gel and blotted using week 7 macaque antiserum. As shown in Fig. 5C, we confirmed antibody reactivity to p15E, p30CA, gp70SU, and p15MA. We did not detect any antibody responses to p10NC or pp12. All of the protein bands identified on the immunoblot assay of whole XMRV corresponded to HPLC-purified viral proteins (Fig. 5A and C), suggesting minimal or no antibody reactivity to nonviral proteins. In addition, we did not detect any serum reactivity with purified microvesicles derived from several human cell lines (see Fig. S3 in the supplemental material), validating that the macaque antiserum reactivity was specific for XMRV and not any human proteins that might have been present in the 22Rv1 cell line-derived virus stock.
To evaluate the titer of anti-XMRV antibodies in serum harvested at necropsy, equivalent amounts of lysed XMRV were blotted with 10-fold serial dilutions of day 119 serum (Fig. 6). The two animals showed indistinguishable antibody titers, with strong XMRV detection at 1:200 and 1:2,000 serum dilutions. At 1:20,000, p15E was still readily detectable; however, other viral proteins were no longer detectable. All immunoblot assay reactivity was lost at a 1:200,000 dilution, indicating a serum immunoblot assay endpoint dilution between 1:20,000 and 1:200,000.
We next examined the neutralization activity in antibody responses elicited by XMRV infection by assessing the neutralization titer in heat-inactivated plasma collected at 2 weeks (peak viremia), 7 weeks, and 17 weeks (necropsy) postinfection, as well as in preinfection control plasma samples (Fig. 7). Neutralization activity was determined using a multicycle infection assay with DERSE.LiG-puro reporter cells. To control for the specificity of the neutralizing antibody response and any potential neutralizing activity directed against nonviral proteins, we compared the plasma neutralization of an infectious XMRV clone, VP62, generated in 293T cells with that of an infectious 293T-produced Mo-MLV clone pseudotyped with VSV-G. Importantly, preinfection plasma samples did not substantially neutralize XMRV or Mo-MLV, with a modest, nonspecific reduction (<40%) in infectivity for both viruses observed only at the lowest plasma dilution (50% effective dilution [ED50], <10) (Fig. 7A). By 2 weeks postinfection, the time point at which peak plasma viremia was detected, XMRV-specific neutralizing activity was measured in plasma samples from both animals, with plasma dilution-dependent reductions in XMRV infectivity and no neutralization of VSV-G-pseudotyped Mo-MLV. Week 2 neutralizing antibody potency was comparable in the two monkeys, with ED50s of 5,240 and 1,083 for 8242 and 14232, respectively, and complete inhibition of XMRV infectivity achieved at lower plasma dilutions. Both animals maintained similar levels of XMRV-neutralizing activity throughout the study, with week 7 and week 17 plasma neutralization titers within 4-fold of those measured for week 2. Nonclonal XMRV produced in 22Rv1 cells had plasma neutralization profiles virtually identical to those of the 293T-produced clone VP62 (data not shown). Taken together with the immunoblotting data, these results indicate that the humoral immune responses elicited by XMRV infection in pigtailed macaques are relatively stable for at least 4 months after infection, despite transient, low-level viremia.
To evaluate the innate antiviral immune response in LNs, longitudinal LN biopsy samples were stained for myxovirus resistance protein A (MxA), an antiviral resistance gene product that is upregulated by type I IFN, commonly used as a marker for innate immune activation. As shown in Fig. 8, MxA expression was transiently upregulated during the first week of XMRV infection, showing expression levels approaching those observed in chronically SIV-infected macaques. However, MxA expression returned to preinfection levels by 14 dpi and remained at baseline levels up to 28 dpi, indicating a transient innate immune response to XMRV infection that is rapidly downregulated concurrent with the resolution of plasma viremia.
We longitudinally monitored changes in immune cell subsets to assess immune reactivity and any potential immunopathology associated with XMRV infection. Within the first 3 weeks of infection, we noted an increase in both animals in the overall frequency of proliferating CD8+ T cells in peripheral blood and LNs, as defined by Ki67 positivity. This increase in Ki67 staining was specific to memory CD8+ T cells (CD95+) and was most pronounced in the central memory (Tcm) subset (CD95+ CD28+) (Fig. 9). Ki67 positivity in CD8 cells in peripheral blood peaked at 3 weeks postinfection and returned to near baseline levels by week 4. In LNs, peak Ki67 staining for CD8 Tcm cells was measured at 2 weeks postinfection (LNs were not sampled at 3 weeks postinfection) but remained elevated at 4 weeks relative to preinfection time points. Changes in the frequency of Ki67 staining for CD4+ T cells were minimal (not shown).
Although the frequency of Ki67+ CD8 cells increased with XMRV infection, we did not measure any marked changes in the absolute number of CD8+ T cells in peripheral blood, nor did we detect any substantial changes in the absolute number of CD4+ T cells, CD20+ B cells, CD14+ monocytes/macrophages, or granulocytes (not shown). In addition, we did not detect any virus-specific immune responses by ICS of CD4+ and CD8+ T cells stimulated with whole XMRV or purified XMRV CA, MA, pp12, NC, PR, RT, IN, or TM protein (see Table S2 in the supplemental material; data not shown).
Although, based on accumulating results, the proposed association of XMRV infection with human disease, as well as evidence of any authentic human XMRV infection, appears increasingly unlikely, the virus itself is a replication-competent retrovirus with unknown pathogenic potential that is capable of infecting human cells (7, 11, 47). Given the lack of any confirmed positive cases of human XMRV infection (29, 58), the in vivo replication capacity, sequence evolution, tissue tropism, and elicited immune responses associated with XMRV infection are unknown and it is unclear what results using a variety of direct and indirect detection methods might be expected in the setting of authentic XMRV infection. Information about the natural history of XMRV infection and host immune responses to the virus would provide a framework to help interpret results suggesting evidence of human XMRV infection. In the absence of any known human infection, animal models must be relied upon to provide this information.
To address these questions in a nonhuman primate species, we intravenously infected two adult male pigtailed macaques with >1010 XMRV virions, an inoculum which likely exceeds by many orders of magnitude any viral inoculum that might be involved in a physiological human transmission. Despite this large viral inoculum, XMRV replicated only transiently to relatively low peak levels in both animals, achieving peak plasma VLs of ≤2,200 RNA copies/ml that declined to undetectable levels within 4 weeks of infection. This decline in viremia was associated with striking levels of G-to-A hypermutation in PBMC-associated vDNA, likely reflective of APOBEC-mediated viral restriction. Although plasma viremia was brief, within the first 2 to 4 weeks of infection, both animals raised robust anti-XMRV antibody responses primarily directed toward p15E, p30CA, and gp70SU that were largely maintained up to 119 dpi. In addition to these binding antibody responses, neutralizing antibodies were also elicited within the first 2 weeks of infection and maintained through the study's conclusion. In contrast, innate immune responses in LNs were only transiently upregulated in the first week of infection and rapidly diminished to baseline levels by 2 weeks postinfection, while adaptive T cell responses were essentially negligible in ICS format assays using whole XMRV virions or purified recombinant XMRV proteins as stimuli.
Our findings differ markedly from some of those made in a previous study by Onlamoon and coworkers, which examined XMRV infection of rhesus macaques (M. mulatta). Although Onlamoon et al. also reported low, transient levels of viremia that declined to undetectable levels within 3 weeks of infection (albeit with a less sensitive qRT-PCR assay) associated with the induction of a relatively stable antibody response with neutralizing activity, there were several key differences between the results of the rhesus macaque infection study and those reported here for pigtailed macaques. First, while we observed a relatively stable pool of PBMC-associated XMRV DNA composed predominantly of G-to-A-hypermutated viral genomes that were detectable through 119 dpi, Onlamoon et al. reported a complete loss of PBMC-associated vDNA within the first month of infection (44). Although a recent follow-up study has shown G-to-A hypermutation in PBMC-associated vDNA sequences in these infected rhesus monkeys (67), the disappearance of vDNA in PBMC seems to suggest that factors other than APOBEC-mediated restriction may contribute importantly to control of XMRV infection in rhesus macaques. It is not clear, however, if the PCR assay used here and that used by Onlamoon and coworkers are equally capable of detecting hypermutated vDNA, which are likely less efficiently amplified due to inefficient priming, and it is possible that the apparent loss of vDNA-positive PBMC in rhesus macaques simply reflects the accumulation of hypermutated genomes.
Other key differences between this study and that of Onlamoon et al. were noted when comparing analyses of various host tissues. In our pigtailed macaques, we identified vDNA in LNMC by X-SCA at early and late postinfection time points; however, longitudinal LN sections were vRNA negative when probed with our ISH riboprobe cocktail, the specificity of which was verified using in vitro-infected pigtailed macaque cells (Fig. 2B; see Fig. S1 in the supplemental material). These results were consistent with our observations on PBMC vis-à-vis plasma viremia and suggest that the vDNA-positive cells detected by X-SCA in LNs were not productively infected. Conversely, Onlamoon et al. reported that LNs from infected rhesus macaques contained productively infected cells detectable by IHC at both early and late (>140 dpi) postinfection time points (44). Since XMRV was initially linked to human prostate cancers, we also evaluated viral replication in prostate tissue. By X-SCA, we detected very low levels (<15 copies/106 cells) of vDNA in snap-frozen prostate tissue pieces collected at necropsy, and no vRNA-positive cells were detectable in prostate tissue sections by ISH. Since a small number of contaminating blood cells could be the source of this low-level vDNA, these data indicate very limited or no XMRV infection in prostate tissue with no productive infection at 119 dpi. Although Onlamoon and coworkers noted fewer productively infected cells in prostate tissue at late postinfection time points than at early postinfection time points, they reported that vRNA+ cells could still be detected by ISH in prostate tissue at >140 dpi (44). These findings on rhesus macaques suggest the establishment of a chronic viral infection, characterized by the stable presence of productively infected cells in LNs and other tissues, concomitant with the apparent loss of vDNA+ PBMC and undetectable plasma viremia, suggesting that once the virus makes it into LNs and other tissues, it continues to replicate but becomes trapped and does not reseed the peripheral blood compartment despite the apparent presence of suitable target cells. This conclusion is in stark contrast to the results we report here for pigtailed macaques, wherein early rounds of viral replication lead to the seeding of PBMC and LNMC with a stable pool of archived, hypermutated, likely nonfunctional viral genomes. In addition to the difference in macaque species, differences in the sensitivity and specificity of the assays employed could also contribute to the discordant results.
In the absence of any confirmed human XMRV infection cases, it is unclear whether infection of pigtailed macaques accurately models human XMRV infection. There is, however, evidence to suggest that several features of pigtailed macaque infection may mirror what would occur in an XMRV-infected human. Previous work has shown that XMRV infection of human PBMC in vitro results in a nonspreading, restricted infection with viral production showing a dose-dependent plateau effect with extensive G-to-A hypermutation in cell-associated vDNA, likely mediated by APOBEC proteins (7). Infection of pigtailed macaque PBMC in vitro showed kinetics and replication patterns strikingly similar to those reported for human cells (Fig. 2A), and we observed extensive G-to-A hypermutation in vivo (Fig. 5). It therefore might be expected that XMRV would be similarly hypermutated and restricted during any in vivo human infection, leading to comparably transient low-level viremia. The potential importance of APOBEC-mediated hypermutation in limiting viral replication is underscored by the rapid decline of plasma VLs to undetectable levels in our infected animals in the face of persistent vDNA-positive cells in blood and LNs, demonstrating that a total elimination of infected cells, by either immunological or viral lytic mechanisms, was not responsible for the disappearance of viremia, but rather that these cells contained defective viral genomes that did not express viral gene products. In agreement with this notion, vDNA in LNMC was first detected by X-SCA just after peak viremia and was still detectable at necropsy; however, longitudinal LN sections were vRNA negative by ISH throughout this study. Although these vDNA+ cells in blood and LNs persisted at 119 dpi, innate immune responses in LNs were only transiently detectable in the first week of infection, in stark contrast to the continual LN immune activation associated with chronic SIV infection (Fig. 8), and cellular adaptive immune responses were negligible, consistent with the view that the hypermutated proviral genomes were rendered nonfunctional and did not express viral gene products. Taken together, these results suggest that APOBEC-mediated hypermutation is likely largely responsible for the limited viral replication we observed in our infected animals. These results may have broader implications concerning the potential for gammaretroviral infection of humans, where such concerns have been raised in particular regarding the transfer of porcine gammaretroviruses to immunosuppressed humans in the setting of xenotransplantation (66). Because gammaretroviruses lack the accessory genes possessed by other retroviruses, such as HIV and SIV, they are ill equipped to counteract intrinsic host restriction factors like the APOBEC3 proteins. As shown here, these primate host restriction factors can be remarkably effective at inhibiting gammaretroviral replication in vivo and may explain, at least in part, why gammaretroviruses have not been significant pathogens in humans, including immunosuppressed individuals, despite close human association with their animal hosts.
Hypermutation of viral genomes also likely explains our unsuccessful efforts to rescue virus from differentially stimulated PBMC collected at the study's conclusion using an LNCaP-based reporter cell line (DERSE.LiG-puro) (data not shown). Given the extensive G-to-A hypermutation identified by X-SGS in cell-associated vDNA (Fig. 4), it is likely that the proviral genomes were rendered incapable of producing infectious progeny. Although vRNA in plasma was not hypermutated, reflecting the fact that only nonhypermutated proviruses produced progeny, we were also unable to rescue infectious virus from plasma samples collected at several postinfection time points (see Fig. S2 in the supplemental material). While we did not expect to rescue plasma virus at time points at which plasma viremia was undetectable, we note that we were also unable to rescue virus from plasma samples taken at the time of known peak viremia. These results suggest that the isolation of replication-competent XMRV from human blood samples may be difficult or unlikely, particularly when working with samples from unknown postinfection time points that are not likely to represent peak viremia, and they are in contrast to those of previous studies that reported an ability to isolate infectious XMRV from human PBMC and plasma samples similarly using LNCaP target cells (34, 40). While these disparities might be explained by methodological differences, it seems that XMRV would have to be less susceptible to human APOBEC3-mediated hypermutation in order for virus to be isolatable from PBMC; however, previous work has shown that XMRV is highly susceptible to APOBEC3-mediated hypermutation in human cells (7, 19, 47). To successfully rescue virus from human plasma, particularly without the ability to select peak viremia time points, XMRV would likely have to replicate to higher sustained levels in humans than those observed in our pigtailed macaques or exist in human plasma in immune complexes that have a less detrimental effect on viral infectivity.
If the course of XMRV infection in pigtailed macaques does reflect the natural history of XMRV infection in humans, we would expect serological assays and PCR assays to detect vDNA in PBMC to have the greatest likelihood of detecting XMRV infection in clinical samples, due to the greater stability of these parameters following XMRV infection in vivo. In contrast, we would expect assays to detect vRNA in plasma and assays to rescue culturable virus to be less likely to detect XMRV infection due to transient positivity and lack of sensitivity, respectively. One of the potential shortcomings of previous studies designed to identify XMRV infection in human samples using PCR, serology, IHC, ISH, virus rescue, or other means has been the lack of bona fide in vivo-derived positive-control samples. Therefore, in addition to characterizing the natural history of XMRV infection in vivo, we initiated these studies with pigtailed macaques to generate and make available positive-control samples for future analyses. We have generated an infectious, 22Rv1-derived XMRV stock and matched highly purified 1,000×-concentrated stocks and determined their virus contents by qRT-PCR and quantitative p30 densitometry. Macaque antiserum raised against infection with this 22Rv1 virus was collected at necropsy and stored for future use. We show here that this serum is XMRV specific, with clear Western blot assay reactivity for p15E, p30CA, and gp70SU at a 1:2,000 dilution (Fig. 5), detectable p15E reactivity at a 1:20,000 dilution (Fig. 6), and negligible background reactivity with microvesicles prepared from human cells (see Fig. S3 in the supplemental material).
The reagents we have generated here should facilitate future studies aimed at examining XMRV infection in in vivo-derived samples. (1×- or 1,000×-concentrated XMRV produced from 22Rv1 cells and serum samples from XMRV-infected pigtailed macaques are available on request from Julian W. Bess, Jr. [vog.hin.liam@wjsseb].) Though XMRV infection of pigtailed macaques doubtless does not perfectly model human XMRV infection, these results suggest that a small, physiological viral inoculum, in the setting of functional APOBEC proteins, will likely replicate only briefly and to extremely low levels, at best, and might therefore be very unlikely to induce any significant pathogenesis.
We thank Michael Piatak for technical assistance extracting DNA from whole tissue pieces. We also thank Mercy Gathuka and the staff of the Laboratory Animal Sciences Program for expert animal care.
This project has been funded in whole or in part with federal funds from the National Cancer Institute, National Institutes of Health, under contract HHSN261200800001E, and by the Intramural Research Program of the National Institutes of Health, National Cancer Institute, Center for Cancer Research, the Office of AIDS Research, and a Bench-to-Bedside Award to Vinay K. Pathak. John M. Coffin was a Research Professor of the American Cancer Society with support from the F. M. Kirby Foundation.
The content of this publication does not necessarily reflect the views or policies of the Department of Health and Human Services, nor does mention of trade names, commercial products, or organizations imply endorsement by the U.S. Government.
Published ahead of print 11 January 2012
Supplemental material for this article may be found at http://jvi.asm.org/.