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. Author manuscript; available in PMC: 2019 Jan 15.
Published in final edited form as: J Immunol. 2017 Dec 6;200(2):632–642. doi: 10.4049/jimmunol.1700927

Osteoblasts are rapidly ablated by virus-induced systemic inflammation following LCMV or PVM infection in mice

Steven Maltby *, Alyssa J Lochrin *, Bianca Bartlett *, Hock L Tay *, Jessica Weaver *, Ingrid J Poulton †, Maximilian W Plank *, Helene F Rosenberg ‡, Natalie A Sims †, Paul S Foster *
PMCID: PMC5760340  NIHMSID: NIHMS919823  PMID: 29212906

Abstract

A link between inflammatory disease and bone loss is now recognized. However, limited data exist on the impact of virus infection on bone loss and regeneration. Bone loss results from an imbalance in remodeling, the physiological process whereby the skeleton undergoes continual cycles of formation and resorption. The specific molecular and cellular mechanisms linking virus-induced inflammation to bone loss remain unclear. In the current study, we provide evidence that infection of mice with either lymphocytic choriomeningitis virus (LCMV) or pneumonia virus of mice (PVM) resulted in rapid and substantial loss of osteoblasts from the bone surface. Osteoblast ablation was associated with elevated levels of circulating inflammatory cytokines, including TNFα, IFNγ, IL-6 and CCL2. Both LCMV and PVM infections resulted in reduced osteoblast-specific gene expression in bone, loss of osteoblasts and reduced serum markers of bone formation, including osteocalcin and procollagen type 1 N propeptide. Infection of Rag-1-deficient mice (which lack adaptive immune cells) or specific depletion of CD8+ T lymphocytes limited osteoblast loss associated with LCMV infection. By contrast, CD8+ T cell depletion had no apparent impact on osteoblast ablation in association with PVM infection. In summary, our data demonstrate dramatic loss of osteoblasts in response to virus infection and associated systemic inflammation. Further, the inflammatory mechanisms mediating viral infection-induced bone loss depend on the specific inflammatory condition.

Introduction

Inflammatory and infectious diseases are major causes of morbidity, may be difficult to treat, and their incidence is high worldwide. Inflammation caused by virus infection is often well characterized at the site of infection, but our knowledge of the impact of infection and resulting inflammation systemically is comparatively limited. Virus infections and associated systemic inflammation have been linked to bone loss under certain specific conditions. For example, Chikungunya virus (family Togaviridae) infection induces progressive bone loss, which responds to treatment with the CCL2 inhibitor, bindarit (1). Similarly, HIV infection results in bone loss through a combination of elevated bone resorption and reduced formation (2). However, there is limited understanding of the impact of acute viral infection on the skeleton.

To assess the effects of severe, acute virus infection on bone remodeling, we performed infections with pneumonia virus of mice (PVM) and lymphocytic choriomeningitis virus (LCMV). PVM (Family, Paramyxoviridae) is a natural mouse pathogen that infects both alveolar macrophages and the respiratory epithelium, resulting in impaired respiratory function in association with localized pro-inflammatory chemokine production (3) and myeloid cell recruitment to the lung (4, 5). LCMV (Family Arenaviridae) is a non-lytic virus that replicates systemically within stromal cells and has been extensively studied as a model of T cell exhaustion and chronic infection (6–8). Most studies linking viral infections to bone metabolism focus on altered hematopoiesis (reviewed in (9, 10)). For example, we found that PVM infection resulted in an increase in myeloid cell production in the bone marrow (11). LCMV infection induced immune-mediated anemia and, in perforin-deficient animals, induced progressive bone marrow failure driven by CD8+ T cells (12, 13).

One potential link between virus infection and altered bone structure is through the induction of systemic inflammation. Inflammation and bone loss, including osteoporosis, are correlated in a range of inflammatory conditions, including rheumatoid arthritis, inflammatory bowel disease and chronic obstructive pulmonary disease (COPD) (14). Such bone loss occurs due to an imbalance in bone remodeling, the normal physiological process by which the skeleton is continually renewed by cycles of bone resorption followed by an equivalent level of bone formation (15). However, the mechanism whereby inflammation leads to bone loss can vary. Arthritogenic alphavirus infection (16), COPD (17) and rheumatoid arthritis (18) are all associated with bone loss that is caused by increased bone resorption, with no effect on osteoblast numbers or activity. Conversely, colitis-induced bone loss has been attributed to impaired bone formation (19). Still other inflammatory conditions associated with bone loss either have unknown etiology (e.g. cystic fibrosis (20)), or the effects of the underlying condition are difficult to discern due to the inhibition of bone formation by long-term corticosteroid treatment (e.g. asthma (21)).

Many immune factors and immune cell types regulate the differentiation and activity of both bone-forming osteoblasts and bone-resorbing osteoclasts. These include interferon-γ (IFNγ), tumor necrosis factor-α (TNFα), CCL2 and IL-6 (reviewed in (22, 23)), as well as macrophages, NK cells, and T cells (24). The relative roles of each of these cytokines and cell types in different inflammatory conditions remain unclear. In order to treat the bone loss that occurs in inflammatory conditions, an improved understanding of how inflammation causes bone loss under different inflammatory settings is required, as it may provide insight into pathogenic mechanisms and new approaches for therapy.

We report here that infection with either LCMV or PVM induced a rapid loss of osteoblasts from the bone surface and thereby reduced bone formation, but did not increase bone resorption. Further, we demonstrate that osteoblast loss was mediated by CD8+ T cells following LCMV infection, but not after PVM infection. Our findings also show that inhibition of osteoblasts occurs very early after viral infection. The difference in etiology between these infections and the other reported infections suggest that treatment of viral infection-induced bone loss may require specific targeting for each condition.

Materials & Methods

Ethics statement

Animal experiments were conducted in accordance with New South Wales, Australia Animal Research legislation. The experimental protocol A-2011-139 has been reviewed and approved by the University of Newcastle Animal Ethics Committee.

Mice, virus infections and interventions

C57Bl/6 and RAG-1-deficient (recombination activating gene 1; Rag-1−/−) mice on a C57Bl/6 background were received from University of Newcastle Animal Services Unit and experiments were performed in the Hunter Medical Research Institute (HMRI) animal facility, under specific pathogen-free conditions.

Mice were infected by intravenous (i.v.) injection of 2×106 pfu LCMV clone 13, to achieve chronic systemic infection, or sham administered DMEM + 10% FCS, as previously described (25). Virus levels were quantified by plaque-forming assay in Vero cells (25). Alternatively, mice were infected by intranasal (i.n.) instillation of 100 pfu PVM strain J3666 in DMEM + 10% FCS, or sham administered DMEM + 10% FCS, as previously described (11). All mice were then monitored daily and weighed as indicated in the text. PVM-infected animals were also assessed based on symptoms as follows: 1 = no signs of illness, 2 = consistently ruffled fur, 3 = piloerection, deeper breathing and decreased alertness. Animals were euthanized by sodium pentobarbitone injection (Virbac, Australia) at the timepoints indicated in the text, and all efforts were made to minimize suffering in treated mice.

Where indicated, mice were injected intraperitoneally (i.p.) with calcein (20 mg/kg in sterile saline) on days -1 and +5 post-infection, for bone formation assessments. To deplete CD8+ T cells, mice were injected i.p. with anti-CD8 antibody (500μg/dose; clone YTS 169.4) or isotype control (clone LTF-2) on days -1, +2 (and also +5 for LCMV) post-infection. CD8+ T cell depletion was confirmed by flow cytometry.

Flow cytometry processing and antibody staining

Tissue samples were collected and processed to single cell suspensions prior to staining. Lung tissue was digested in HEPES buffer containing collagenase D (Sigma-Aldrich; St Louis, MO) and DNAse for 1 hour, then forced through a 70μm strainer. Bone marrow was flushed in sterile PBS +2% FCS. To assess osteoblasts, flushed femurs were crushed using a mortar and pestle, digested with collagenase D, DNAse and dispase, then rinsed through a 70μm strainer.

Following isolation, red blood cell lysis was performed using ammonium chloride solution. All cells were incubated with anti-FcγRIII/II (Fc block) prior to staining with combinations of the following fluorochrome conjugated antibodies as indicated in the text (all antibodies from BD Biosciences unless otherwise indicated; San Jose, CA): Osteoblast Panel: Lineage-APC cocktail (containing CD3e (145-2C11); CD11b (M1/70); B220 (RA3-6B2); Ly-76 (TER-119); Gr-1 (RB6-8C5)), CD45-FITC (30-F11), CD31-PerCP-Cy5.5 (390; BioLegend), ScaI-PE-Cy7 (D7), CD51-PE (RMV-7; BioLegend), Lymphocyte Panel: CD3e-PE (145-2C11), B220-FITC (RA3-6B2), CD8a-PerCP (56-6.7) and CD4-APC (RM4-5). Samples were fixed overnight in PBS/2% FCS + 0.1% PFA, collected on a BD FACSCanto II flow cytometer and analyzed using FlowJo software (FlowJo; Ashland, OR).

Protein assessment

Lung tissue was disrupted in radioimmunoprecipitation assay buffer (RIPA; Sigma-Aldrich) with protease/phosphatase inhibitor cocktail (Cell Signaling Technology; Danvers, MA), on a Tissuelyser LT tissue disruptor (Qiagen; Valencia, CA) at 50 Hz for 5 minutes and stored at -80°C. Serum was collected by cardiac puncture and centrifugation, after allowing blood to clot. Inflammatory cytokine levels for IL-6, CCL2, IFNγ and TNFα were assessed using the mouse inflammation cytometric bead array kit (BD BioSciences), according to manufacturer’s specifications on a BD FACSCanto II flow cytometer and analyzed using FCAP Array v3 software (BD BioSciences). Enzyme-linked immunosorbent assay (ELISA) was used to quantify osteocalcin and P1NP using commercial kits (OSC, Immutopics, CA; P1NP, Immunodiagnostic Systems) according to the manufacturer’s instructions, and measured on a SpectraMax M5 Microplate Reader (Molecular Devices) with SoftMax Pro v5.4.3 software.

RNA isolation, reverse-transcription (RT)-PCR and quantitative PCR (qPCR) assessment

Lung tissue was placed in RNALater (Invitrogen; Carlsbad, CA) and stored at -80°C. Unflushed femurs were disrupted in Trizol Reagent (Invitrogen), on a Tissuelyser LT tissue disruptor (Qiagen) at 50 Hz for 5 minutes and stored at -80°C. Total RNA was isolated by phenol-chloroform separation and isopropanol precipitation and quantified on a Nanodrop 1000 spectrophotometer (Nanodrop; Wilmington, DE). cDNA was prepared by RT-PCR using random hexamer primers (Invitrogen) and MMLV reverse transcriptase (Invitrogen), on a T100 thermal cycler (Bio-Rad; Hercules, CA). Relative qRT-PCR quantification was performed on a ViiA7 real-time PCR machine (Life Technologies; Carlsbad, CA), using SYBR reagents. Measured cDNA levels were normalized to the housekeeping gene Hprt1. Primer sets (Table I) were designed across exon boundaries to specifically amplify mRNA products.

Table I.

Primer sequences used for qPCR analysis

Gene Forward Primer Reverse Primer
Bglap1 ttc tgc tca ctc tgc tga ccc t ccc tcc tgc ttg gac atg aa
Alpl cgg atc ctg acc aaa aac c tca tga tgt ccg tgg tca at
Osx ctg ctt gag gaa gaa gct cac ta cct ttc ccc agg gtt gtt ga
Runx2 cgt gtc agc aaa gct tct ttt ggc tca cgt cgc tca tct
PVM SH gcc tgc atc aac aca gtg tgt gcc tga tgt ggc agt gct t
LCMV GP tgc ctg acc aaa tgg atg att ctg ctg tgt tcc cga aac act
Hprt1 agg cca gac ttt gtt gga ttt gaa caa ctt gcg ctc atc tta ggc ttt
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Note: All primer sequences are indicated in 5′ – 3′ orientation.

Bone histomorphometry

Bones for histomorphometry were embedded in methyl methacrylate resin, as previously described (26). Five-micron thick sections were stained with toluidine blue to visualize osteoblasts and osteoclasts, or with xylenol orange to visualize calcein labels. Histomorphometry was performed on undecalcified methyl methacrylate-embedded sections of the tibial distal metaphysis as previously described (26) using the Osteomeasure Image analysis system (Decatur, GA).

Statistical analysis

Statistical analysis was performed using GraphPad Prism v6.01 software. For comparisons between 2 groups, an unpaired two-way Student’s t test was used. For comparisons between multiple groups, a one-way ANOVA was used, corrected for multiple comparisons. P values <0.05 were considered statistically significant.

Results

Chronic LCMV infection induces systemic inflammation and prolonged reduction in bone marrow cellularity and osteoblasts

We assessed the impact of chronic, systemic virus infection on the bone marrow, using the well-established LCMV (clone 13) intravenous infection model (25). Weight loss was observed from day 8 through day 10 post-infection, which then stabilized through day 20 (Fig. 1A). Following infection, total bone marrow cellularity progressively decreased throughout the course of the experiment (Fig. 1B). LCMV persistence was confirmed in the serum by plaque-forming assay, and in bone and spleen tissue by qPCR (Fig. 1C–E). Serum concentrations of the inflammatory cytokines TNFα, IFNγ, IL-6 and CCL2 were all markedly increased on day 8 post-infection (Fig. 1F). TNFα, IFNγ and IL-6 levels then subsided, while serum CCL2 concentrations remained elevated at all endpoints assessed (Fig. 1F).

Figure 1. Chronic LCMV Infection Reduces Bone Marrow Cell Count and Induces Systemic Inflammation.

Figure 1

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C57Bl/6 mice were infected with 2×106 pfu LCMV (i.v.). Animals were monitored for (A) weight loss and assessed at days 8, 14 and 20 post-infection for (B) bone marrow cell count, viral load in (C) serum by plaque-forming assay and (D) bone and (E) spleen by qPCR, normalized to Hprt1. (F) Levels of TNFα, IFNγ, IL-6 and CCL2 quantified in serum by cytometric bead array assay. White bars = sham samples pooled from each endpoint, black bars = LCMV-infected (Data = mean +/− SEM, >2 replicate experiments, n = 4 - 6 animals/group. * represents p<0.05, ** p<0.01, *** p<0.001, **** p<0.0001 compared to sham-infected controls; Sh = Sham).

LCMV infection was associated with marked reduction in mRNA levels of the osteoblast-specific genes osteocalcin (bone gamma carboxyglutamate; Bglap1), alkaline phosphatase (Alpl), osterix (Osx) and runt-related transcription factor 2 (Runx2) (Fig. 2A) detected in unflushed femur bone samples evaluated at day 8 post-infection. While osteoblast-specific mRNA expression increased somewhat over time, from day 8 through day 20 post-infection, levels remained lower than sham-infected controls at all time points evaluated (Fig. 2A). Osteoblasts (defined as Lineage−CD45−CD31−Sca-I−CD51+ (27–30)) were also markedly reduced at all timepoints, from 0.56 +/− 0.07% to 0.03 +/− 0.01% of total cells isolated from bone by enzymatic digestion at day 8 of infection (Fig. 2B; ****p < 0.0001). Serum osteocalcin and procollagen type 1 N propeptide (P1NP) levels were also decreased in response to LCMV infection and, in parallel with the aforementioned mRNA marker genes, serum levels began to recover over the course of the experiment (Fig. 2C). Based on the striking reduction in osteoblast levels following LCMV infection, we also assessed early timepoints post-infection. Diminished levels of osteoblast-specific gene expression (Fig. 2D) and reduced osteoblasts assessed by flow cytometry (Fig. 2E) were observed as early as day 4 post-LCMV infection.

Figure 2. Chronic LCMV Infection Reduces Osteoblast Numbers and Circulating Markers of Bone Formation.

Figure 2

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C57Bl/6 mice were infected with 2×106 pfu LCMV (i.v.) and assessed at days 2, 4, 6, 8, 14 and 20 post-infection. (A/D) Levels of osteoblast-specific gene expression for Bglap1, Alpl, Osx and Runx2 in bone by qPCR, normalized to Hprt1. (B/E) Osteoblasts (Lineage−CD45−CD31−Sca-I−CD51+), expressed as the percentage of total isolated cells from digested bone samples, quantified by flow cytometry. (C) Levels of serum osteocalcin and P1NP quantified by ELISA. White bars = pooled sham samples, black bars = LCMV-infected (Data = mean +/− SEM, >2 replicate experiments, n=4-6 animals/group. * represents p<0.05, ** p<0.01, *** p<0.001, **** p<0.0001 compared to sham-infected controls; Sh = Sham).

Acute PVM infection reduces bone osteoblast numbers and circulating markers of bone formation

To determine whether osteoblast loss was unique to LCMV infection, we also characterized responses to infection with the mouse pneumovirus, PVM. After initiation of PVM infection, reduction in osteoblast-specific gene expression was also observed, at day 8 post-infection, compared to sham-infected controls by qRT-PCR (Fig. 3A), in association with a reduction in osteoblasts quantified by flow cytometry (Fig. 3B); serum osteocalcin and P1NP quantified by ELISA were likewise diminished (Fig. 3C). We previously demonstrated that TNFα, IFNγ, IL-6 and CCL2 were all increased in lung tissue during PVM infection, with comparable elevations in serum IFNγ, IL-6 and CCL2 (although serum TNFα levels were typically below the limit of detection) (11).

Figure 3. Acute PVM Infection Reduces Osteoblast Numbers and Circulating Markers of Bone Formation.

Figure 3

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C57Bl/6 mice were infected with 100 pfu PVM (i.n.) and assessed at days 3, 6 and 8 post-infection. (A) Levels of osteoblast-specific gene expression for Bglap1, Alpl, Osx and Runx2 in bone by qPCR, normalized to Hprt1. (B) Osteoblasts (Lineage−CD45−CD31−Sca-I−CD51+) in digested bone samples quantified by flow cytometry. (C) Levels of serum osteocalcin and P1NP quantified by ELISA. White bars = sham, black bars = PVM-infected (Data = mean +/− SEM, >2 replicate experiments, n=4-6 animals/group. * represents p<0.05, *** p<0.001, **** p<0.0001 compared to sham-infected controls).

Bone histomorphometry confirms osteoblast loss and reduced bone formation

The reductions in osteoblast-associated genes, serum biochemistry and flow cytometric indices of osteoblasts led us to perform histomorphometry of trabecular bone at day 8 post-infection in mice infected with LCMV or PVM. Newly formed un-mineralized bone matrix (osteoid) volume and surface were markedly lower in infected mice compared to sham-infected controls (Fig. 4A), and in some samples osteoid was completely absent. Loss of osteoblasts in response to LCMV and PVM infections was also confirmed by this independent method (Fig. 4B).

Figure 4. Histomorphometry Assessment Identifies Osteoblast Loss and Reduced Bone Formation.

Figure 4

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C57Bl/6 mice were infected with 2×106 pfu LCMV (i.v.) or 100 pfu PVM (i.n.), injected with calcein (20 mg/kg i.p.) on day -1 and +5 and assessed 8-days post-infection. Bone histomorphometry assessments performed on femur sections to quantify (A) osteoid volume and surface and (B) osteoblast numbers; BS = bone surface. (C) Representative images for each infection; with 1st (1) and 2nd (2) calcein labels and new bone between the 2nd label and endocortical surface (arrow) indicated; m = marrow, c = cortical bone, t = trabecular bone; scale bar = 100 micron. (D) Calcein deposition quantified based on areas of single and double-labeling and (E) osteoclast numbers and surface area. White bars = sham, black bars = infected (Data = mean +/− SEM, n=4-6 animals/group. * represents p<0.05, *** p<0.001, **** p<0.0001 compared to sham-infected controls).

Quantification of fluorescent calcein label incorporated on bone-forming surfaces revealed no significant changes in mineral appositional rate or bone formation rate. Inspection of calcein labels at the endocortical surface confirmed that bone formation had occurred up until the time of the injection of the 2nd label (day 5 post-infection), in both infection models (Fig. 4C). There was no consistent difference in the thickness of the calcein label. However, the thickness of new bone deposited after that 2nd label was noticeably reduced in the bones from LCMV and PVM-infected mice, and osteoblasts were not observed on endocortical surfaces (Fig. 4C). Further, when quantified in the trabecular bone following LCMV infection, there was less bone surface with two layers of calcein fluorescence, and a corresponding increase in single-labeled bone surface, indicating a rapid reduction in active bone-forming surfaces (Fig. 4D). In parallel, an ~50% decrease in osteoclasts was observed after LCMV infection (Fig. 4E). A population of large multinucleated cells that were morphologically similar to osteoclasts was observed in PVM-infected mice, but not in sham or LCMV-infected mice (Suppl. Fig. 1). While these cells were large and multinucleated, they were not attached firmly to the bone surface, and did not stain positively for tartrate-resistant acid phosphatase.

Rag-1-deficiency suppresses LCMV-induced inflammation and protects from osteoblast loss

T cells, notably CD8+ T cells, play a prominent role in anti-viral responses, and have been extensively characterized in LCMV infection (31). However, limited data are available on the effect of CD8+ T cells on osteoclast and osteoblast function (24). To determine the role of adaptive immune cells in LCMV-induced loss of osteoblasts, Rag-1−/− mice, which lack adaptive immune cells (T and B cells) were employed. At day 8 of infection, LCMV-infected wild-type mice demonstrated significant weight loss, in contrast to their LCMV-infected Rag-1−/− counterparts (Fig. 5A). Levels of LCMV detected in bone tissue were significantly higher in Rag-1−/− mice, compared to wildtype controls (Fig. 5B), consistent with the requirement for adaptive immune cells for virus clearance. There was a trend towards decreased bone marrow cellularity in wildtype mice following LCMV infection, which was not statistically significant (Fig. 5C). In LCMV-infected wild-type mice, substantial increases in CD8+ T cells were detected in bone marrow; this was not observed in Rag1−/− mice (Fig. 5D). Levels of circulating pro-inflammatory cytokines (TNFα, IFNγ, CCL2) were significantly diminished in Rag-1−/− mice, and IFNγ was nearly undetectable (Fig. 5E). Serum IL-6 levels were not affected by Rag-1-deficiency. Overall, we found that systemic inflammation is significantly limited in LCMV-infected Rag-1−/− mice compared to wildtype controls.

Figure 5. Rag-1-deficiency Dampens Inflammation and Suppresses Osteoblast Loss During LCMV Infection.

Figure 5

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Rag-1−/− and wildtype control mice were infected with 2×106 pfu LCMV (i.v.). Animals were monitored for (A) weight loss and assessed at day 8 post-infection. (B) LCMV viral levels in bone quantified by qPCR, normalized to Hprt1. (C) Total bone marrow cell count and (D) CD8+ T cell count (CD3+), assessed by flow cytometry. (E) Levels of TNFα, IFNγ and CCL2 quantified in serum by cytometric bead array. (F) Levels of osteoblast-specific gene expression for Bglap1, Alpl, Osx and Runx2 in bone by qPCR, normalized to Hprt1. (G) Levels of serum P1NP, quantified by ELISA. (H) Osteoblasts (Lineage−CD45−CD31−Sca-I−CD51+) in digested bone samples quantified by flow cytometry, also presented as normalized data to respective sham-infected strain controls. (Data = mean +/− SEM, 3 replicate experiments, n=4-6 animals/group. * represents p<0.05, ** p<0.01, *** p<0.001, **** p<0.0001, compared to sham-infected controls or as indicated).

Rag-1−/− mice were also protected from osteoblast ablation. Specifically, osteoblast-specific gene expression in LCMV-infected Rag1−/− mice was comparable to that detected in sham-infected Rag-1−/− controls, while, as shown earlier, gene-expression was significantly diminished in LCMV-infected wildtype mice (Fig. 5F). Circulating levels of serum P1NP likewise remained undiminished in LCMV-infected Rag-1−/− mice (Fig. 5G). Osteoblasts identified by flow cytometry were diminished dramatically in response to LCMV infection in wild-type mice (to 7.4% of sham-infected control) (Fig. 5H). This reduction was blunted in LCMV-infected Rag-1−/− mice (32.7% of sham-infected Rag-1−/− controls) (Fig. 5H).

CD8+ T cell depletion protects from osteoblast loss during LCMV Infection

To assess the role of CD8+ T cells, we administered anti-CD8 depleting antibodies before virus infection. Administration of CD8-depleting antibodies to wild-type mice prior to and during LCMV infection protected from LCMV-induced weight loss and resulted in higher virus titers in bone tissue, compared to isotype-treated controls (Fig. 6A/B). Similar, albeit slight decreases in total bone marrow cellularity were observed after LCMV infection, regardless of antibody treatment (Fig. 6C). We observed effective CD8+ T cell depletion following anti-CD8 antibody administration in the bone marrow (Fig. 6D). Anti-CD8 treatment reduced circulating TNFα levels, but increased IFNγ and did not appreciably alter circulating CCL2 levels compared to isotype-treated controls (Fig. 6E). Importantly, CD8+ T cell depletion protected from both loss of osteoblast-specific gene expression (Fig. 6F) and reductions in circulating osteocalcin and P1NP levels (Fig. 6G). These findings provide evidence that CD8+ T cells are critical for osteoblast loss following LCMV infection.

Figure 6. CD8+ T cell Depletion Suppresses Osteoblast Loss During LCMV Infection.

Figure 6

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C57Bl/6 mice were infected with 2×106 pfu LCMV (i.v.) and treated with anti-CD8 antibodies on days -1, +2 and +5 post-infection. Animals were monitored for (A) weight loss and assessed at day 8 post-infection. (B) LCMV viral levels in bone quantified by qPCR, normalized to Hprt1. (C) Total bone marrow cell count and (D) CD8+ T cell count (CD3+), assessed by flow cytometry. (E) Levels of TNFα, IFNγ and CCL2 quantified in serum by cytometric bead array. (F) Levels of osteoblast-specific gene expression for Bglap1, Alpl, Osx and Runx2 in bone by qPCR, normalized to Hprt1. (G) Levels of serum osteocalcin and P1NP by ELISA. (Data = mean +/− SEM, 2 replicate experiments, n=4-6 animals/group. * represents p<0.05, ** p<0.01, *** p<0.001, **** p<0.0001, compared to sham-infected controls or as indicated).

Rag-1-deficiency does not protect from osteoblast loss after PVM infection

In contrast to the observations in LCMV infected mice, Rag-1-deficiency had no impact on weight loss or clinical scores after PVM infection, when compared to wildtype controls (Fig. 7A). Virus titers in lung tissue were increased in PVM-infected Rag-1−/− mice, compared to wildtype controls (Fig. 7B). Similar to our findings with LCMV infection, CD8+ T cell numbers were increased in the bone marrow and lung tissue of wildtype mice on day 8 post PVM infection, but were absent in Rag-1−/− mice (Fig. 7C/D). IFNγ induction was nearly completely suppressed in serum and lung tissue; CCL2 was partially suppressed in lung tissue in Rag-1−/− mice, compared to wildtype controls (Fig. 7E/F). TNFα and IL-6 levels were not affected by Rag-1-deficiency. Interestingly, Rag-1-deficiency had no impact on diminished expression of osteoblast genes osteocalcin (Bglap1), alkaline phosphatase (Alpl) or osterix (Osx) (Fig. 7G). These findings demonstrate that adaptive immune cells are not required for inflammation or osteoblast loss following PVM infection. Similarly, antibody-mediated CD8+ T cell depletion had no effect on osteoblast loss following PVM infection (Suppl. Fig. 2). Thus, while both virus infections decrease osteoblast numbers, loss only required CD8+ T cells following LCMV infection.

Figure 7. Rag-1-deficiency Fails to Suppress Osteoblast Loss During PVM Infection.

Figure 7

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Rag-1−/− and wildtype control mice were infected with 100 pfu PVM (i.n.) and assessed at day 8 post-infection. Animals were monitored daily for (A) weight loss and clinical symptoms. (B) PVM viral levels in lung by qPCR, as copies PVM/copies Hprt1. CD8+ T cell counts (CD3+), by flow cytometry in (C) bone marrow and (D) lung tissue. Levels of IFNγ and CCL2 in (E) serum and (F) lung homogenates by cytometric bead array. (G) Levels of osteoblast-specific gene expression for Bglap1, Alpl, Osx and Runx2 in bone by qPCR, normalized to Hprt1. (Data = mean +/− SEM, 2 replicate experiments, n=4-6 animals/group. * represents p<0.05, ** p<0.01, *** p<0.001, **** p<0.0001, compared to sham-infected controls or as indicated).

Discussion

Our findings demonstrate that both LCMV and PVM infection cause a rapid loss of osteoblasts and reduce bone formation and indicate that, in LCMV infection, this response is dependent on the actions of CD8+ T cells. Our findings provide evidence that acute loss of osteoblasts may be a common feature of severe viral infection, which has not previously been recognized. We propose that reduced bone formation caused by infection-induced systemic inflammation may have a significant detrimental effect on the skeleton and may be linked to osteopenia associated with systemic inflammatory diseases.

The ability of some viruses (e.g. arthritogenic alphaviruses such as chikungunya virus) to infect osteoblasts and lead to bone damage has been recognized (32). These effects are thought to require accumulation of virus within the bone. We now demonstrate, for the first time, that viruses that do not directly infect osteoblasts, including those that are excluded from the bone microenvironment (i.e. PVM), can also rapidly ablate osteoblasts and inhibit bone formation. As a member of the genus Pneumovirus, PVM remains restricted to the lung; transcript was not detected in samples from total unflushed bone, yet infection had a rapid and substantial impact on osteoblast number and surface volume (Fig. 4). Likewise, there have been no reports of LCMV infection of osteoblasts, and furthermore the LCMV receptor (alpha-dystroglycan (33)) is not expressed by these cells. In the absence of CD8+ T cells (in Rag-1−/− mice and after anti-CD8 treatment), although LCMV is present in bone samples, osteoblast loss was suppressed despite an increase in total virus. Thus, it is unlikely that LCMV leads to cell loss via direct infection of osteoblasts.

Rag-1-deficiency and CD8+ T cell depletion individually protected mice from osteoblast loss associated with LCMV infection; the same response was not observed following infection with PVM. There are multiple differences between the two infections that could explain this difference, such as local versus systemic infection and differences in tissue localization and route of infection. We propose our observation reflects intrinsic differences in the inflammatory pathogenesis underlying LCMV and PVM infections, with CD8+ T cells being secondary to the systemic inflammatory response following PVM infection. LCMV infection induces systemic inflammation through a rapid and robust adaptive immune response, including recruitment and activation of CD8+ T cells, followed by T cell exhaustion and virus persistence (6). In contrast, PVM infection-induced inflammation is primarily driven by innate immune cells (3, 4, 34, 35), although CD8+ T cells do contribute to inflammatory pathology and are a major source of IFNγ and TNFα (36). While adaptive immune cells eventually clear PVM virus, inflammation and pathology are linked to the potency of the innate response, rather than absolute levels of virus in the lung tissue (34). Thus, we propose that CD8+ T cells are key for the development of systemic inflammation induced by LCMV infection and subsequent loss of osteoblasts. In contrast, adaptive immune responses, including CD8+ T cells, are secondary to the systemic inflammatory response following PVM infection and are not required for osteoblast ablation. In line with this hypothesis, we also noted that Rag-1−/− mice were protected from weight loss induced by LCMV infection, but not following PVM infection. We do note that we only assessed infections in C57Bl/6 mice, and that PVM infection kinetics and inflammation characteristics vary between mouse strains (e.g. BALB/c (37, 38)).

The role of specific inflammatory cytokines in osteoblast loss in our models remains to be determined. Both PVM and LCMV infection induced systemic inflammation, with increased levels of circulating TNFα, IFNγ, IL-6 and CCL2. While TNFα, IL-6 and CCL2 have previously been demonstrated to promote osteoclast differentiation (39–42), the effects of inflammatory cytokines on osteoblasts are more controversial. In vitro, TNFα impairs osteoblast differentiation (43), causes down-regulation of osteocalcin and Runx2 (44) and can directly induce osteoblast apoptosis (45). IFNγ promotes early osteoblast differentiation from mesenchymal stem cells (46) and IFNγ administration significantly increases bone mass and the ratio of bone formation/resorption in studies performed in ovariectomized mice (47). However, IFNγ also inhibits osteoclast activity and its role in the regulation of bone remodeling also remains controversial (reviewed in (48)). Osteoblasts produce CCL2 under inflammatory conditions (49) and blocking CCL2 function reduces bone loss in experimental chikungunya virus infection (1). IL-6 treatment of cultured cells promotes osteoblast activity (50) and osteoclast formation (42), however addition of soluble receptor was required for both effects. Studies in IL-6 null mice have shown that IL-6 plays a role in the increased osteoclastogenesis associated with inflammatory conditions such as rheumatoid arthritis (51) and ovariectomy (52), but that the physiological role of IL-6 may be restricted to promoting bone formation specifically on the periosteal surface that is not in contact with the bone marrow (53, 54). A role for IL-6 in inflammation-induced changes in osteoblastic bone formation has not yet been reported.

In our study, osteoblast loss was prevented both when serum levels of TNFα, IFNγ and CCL2 were reduced in LCMV-infected Rag-1−/− mice and likewise when serum TNFα alone was reduced by anti-CD8 treatment. Thus, increased serum TNFα during LCMV infection, rather than the other inflammatory cytokines assessed, may serve as a mechanism linking CD8+ T cell responses and osteoblast loss. To assess this hypothesis, we specifically blocked TNFα during LCMV infection. However, administration of anti-TNFα antibodies during LCMV infection failed to dampen the loss of osteoblast-specific gene expression (Suppl. Fig. 3A). Further, administration of blocking antibodies against TNFα, IFNγ, CCL2 or IL-6 individually failed to suppress osteoblast loss following PVM infection (Suppl. Fig. 3B–E). Of note, antibody-mediated inhibition of individual inflammatory cytokines also failed to suppress weight loss (Suppl. Fig. 3), providing evidence that targeting individual cytokines failed to suppress broader inflammatory processes. As such, we conclude that osteoblast loss may result from overlapping, compensatory mechanisms between individual cytokine signaling pathways and/or may require inflammatory cytokines not evaluated in the current study.

The dramatic decrease in osteoblast numbers caused by both LCMV and PVM infections could result from apoptosis, migration and/or de-differentiation of osteoblasts. The rapid kinetics of osteoblast ablation suggests this is an active process, rather than a cumulative effect of reduced osteoblast differentiation caused by long-term effects of virus infection, such as malnutrition. As osteoblast-specific gene mRNA levels were reduced in bone samples that had not been flushed of marrow, this suggests that osteoblasts have not relocated to the bone marrow compartment. The rapid disappearance of osteoblasts is striking in its similarity to the rapid loss of osteoblasts observed during G-CSF stimulation or cyclophosphamide treatment (55, 56), in which osteoblasts disappeared from the bone surface within days of commencing treatment. Cyclophosphamide-dependent osteoblast loss was shown to be dependent on macrophages, as protection was observed in Mafia (macrophage-deficient) mice (56). Rapid osteoblast loss has also has been reported in a mouse model of graft-versus-host disease (GVHD) via a mechanism requiring CD4+ T cells, which are a key driver of inflammation in this model (57). Osteoblast ablation was also observed in patients with chronic GVHD, and was associated with decreased B cells and an increased CD4/CD8 T cell ratio (58). Sepsis induced by cecal ligation and puncture also caused severe acute inflammation, rapid osteoblast loss and reduced bone formation, without altering bone resorption (59). The kinetics and mechanism of osteoblast recovery after acute infection also warrant further investigation.

While our data provides clear evidence that osteoblasts are lost following infection, it remains unclear whether this is a negative side effect of inflammation or is part of the functional response to infection. One possibility is that bone formation is reduced during severe acute infection to redirect energy towards short-term survival, potentially including altered immune cell production. A potential link exists between osteoblast ablation and immune cell production and differentiation. Early data identified osteoblasts as a component of the hematopoietic stem cell niche that regulates immune cell differentiation (60, 61). While recent data suggests the “osteoblast niche” may not have a primary role in hematopoietic stem cell regulation (reviewed in (62)), the precise role of osteoblasts in regulating immune progenitor differentiation remains unclear. We previously reported that PVM infection increased myeloid cell production in the bone marrow (11). We also quantified immune cell subsets in the bone marrow after LCMV infection (Suppl. Fig. 4). LCMV infection transiently increased Ly6C+ monocyte numbers in the marrow at day 8 post LCMV infection (Suppl. Fig. 4E), whereas neutrophil and eosinophil numbers were decreased at days 14 and 20 post-infection (Suppl. Fig. 4C/D). B cell numbers were dramatically decreased in the bone marrow following LCMV infection (Suppl. Fig. 4I), as has previously been reported during acute infection (63). Similarly, in the GVHD and sepsis models mentioned previously, osteoblast ablation was associated with reduced lymphoid cell production (57, 59). Thus, osteoblast ablation may shape immune cell production in response to severe infection. Alternatively, osteoblast ablation may be a negative effect of viral pathogenesis. For example, LCMV infection has previously been associated with bone marrow suppression and suppressed immunity (64, 65). Regardless of the cost/benefit of osteoblast loss during acute viral infection, in the context of chronic inflammation this process likely has a negative effect on long-term bone health. We expect that the observed osteoblast ablation would have long-term effects on bone structure and/or integrity under conditions of chronic inflammation.

Clinically, bone loss is associated with a diverse range of inflammatory conditions (2, 16–18, 20, 21). Of particular interest, a 1-year follow-up study of patients admitted to ICU and requiring mechanical ventilation revealed significant decreases in bone mineral density compared to controls, regardless of diagnosis (66). These findings support the conclusion that severe inflammation is linked to bone loss, regardless of the cause. Our findings demonstrate a significant reduction in osteoblast numbers and function following acute severe viral infection, which should be considered as a contributing factor to osteopenia in patient management.

In summary, our data demonstrate a dramatic loss of osteoblasts in association with severe viral infections; we suggest that this loss is a direct result of systemic inflammation. Translation of these observations into clinical settings suggests that serum biochemical markers of osteoblast and osteoclast activity should be assessed in susceptible populations (e.g. the elderly or patients with multiple medically-relevant conditions) after viral infection and throughout recovery. Further, the long-term impacts on bone structure and strength should be assessed under conditions of persistent, infection-induced inflammation.

Supplementary Material

1

Acknowledgments

We thank Associate Professor Scott Mueller (University of Melbourne), for providing LCMV Clone 13 stocks, Narelle E. McGregor and Brett A. Tonkin (St Vincent’s Institute of Medical Research) for analysis and the Analytical Biomolecular Research Facility (The University of Newcastle) for flow cytometry support.

Financial Support: This work was supported by project grants from the National Health and Medical Research Council (NHMRC) of Australia, the NIAID Division of Intramural Research and a start-up grant from the University of Newcastle. SM was supported by fellowships from the Canadian Institutes of Health Research (CIHR) and the University of Newcastle.

Footnotes

Author Contributions

S.M. and A.J.L. designed research, performed research, collected data, analyzed and interpreted data, performed statistical analysis and wrote the manuscript. B.B. and J.W. performed research, collected data and analyzed data. H.L.T. and M.W.P. interpreted data and wrote the manuscript. I.J.P. performed analysis. H.F.R. contributed reagents, interpreted data and wrote the manuscript. N.A.S. and P.S.F. designed research, interpreted data and wrote the manuscript.

Disclosure

The authors have no conflicts of interest to declare.

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NIHMS919823-supplement-1.pdf (461.9KB, pdf)

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