Regulatory T Cells Promote Myositis and Muscle Damage in Toxoplasma gondii Infection

Richard M. Jin, Sarah J. Blair, Jordan Warunek, Reid R. Heffner, Ira J. Blader and Elizabeth A. Wohlfert
J Immunol January 1, 2017, 198 (1) 352-362; DOI: https://doi.org/10.4049/jimmunol.1600914

Abstract

The coordination of macrophage polarization is essential for the robust regenerative potential of skeletal muscle. Repair begins with a phase mediated by inflammatory monocytes (IM) and proinflammatory macrophages (M1), followed by polarization to a proregenerative macrophage (M2) phenotype. Recently, regulatory T cells (Tregs) were described as necessary for this M1 to M2 transition. We report that chronic infection with the protozoan parasite Toxoplasma gondii causes a nonresolving Th1 myositis with prolonged tissue damage associated with persistent M1 accumulation. Surprisingly, Treg ablation during chronic infection rescues macrophage homeostasis and skeletal muscle fiber regeneration, showing that Tregs can directly contribute to muscle damage. This study provides evidence that the tissue environment established by the parasite could lead to a paradoxical pathogenic role for Tregs. As such, these findings should be considered when tailoring therapies directed at Tregs in inflammatory settings.

Introduction

The current paradigm for muscle repair is based on the sequential and discrete actions of inflammatory monocytes (IM)/proinflammatory macrophages (M1) and proregenerative macrophages (M2) (1–3). IM/M1 mediate the acute inflammatory phase of repair by production of lytic factors to break down necrotic cells, phagocytosis of cellular debris, and satellite cell activation, a population of myofiber progenitor cells (2, 4, 5). M2 facilitate a regenerative phase through suppression of inflammatory cells and production of tissue remodeling products, such as collagen matrix enzymes. Critically, the IM/M1 to M2 balance in the muscle environment drives both the phase and efficiency of repair (1, 2, 6). Prolonged IM/M1 presence during muscle injury may lead to delayed or aberrant repair (7–13), whereas exuberant M2 activity may cause muscle fibrosis (6). However, it remains unclear how muscles are repaired in the context of chronic inflammation, and the factors that prevent repair from proceeding remain unidentified. Recent studies implicate a tissue-specific role for resident regulatory T cells (Tregs) in the transition from IM/M1 to M2 during tissue repair in acute sterile injury and dystrophy models (12, 13). Still, greater insight into how repair proceeds in the face of immune dysfunction and chronic inflammation is needed because these are hallmark features of a variety of debilitating inflammatory myopathies, such as polymyositis, dermatomyositis, and idiopathic inflammatory myositis (6, 14–16).

Chronic infections of muscle can represent a source of chronic inflammation that may have detrimental consequences for muscle physiology. In this article, we examine skeletal muscle injury and potential tissue repair in the context of infection with Toxoplasma gondii in mice. T. gondii persists as a life-long chronic infection characterized by parasite-containing tissue cysts in the brain and skeletal muscle (17, 18). Ingestion of tissue cysts or oocysts shed in feline feces results in a robust Th1 immune response to the parasite in the gastrointestinal tract that controls the fast-growing tachyzoite life stage. Parasites that escape this response disseminate throughout the host, and IFN-γ production by innate and adaptive cells is required for control of parasite growth (19). The mechanisms by which IFN-γ operates to limit tachyzoite proliferation include induction of immunity-related GTPases and inducible NO synthase (iNOS) in a number of host cell types, including M1 (20, 21). Immune pressure induces the transition of the parasite to slow-growing tissue cysts (bradyzoites) that reside in the skeletal muscle and CNS. Because contaminated meats represent a major parasitic reservoir and source of food-borne transmission, tight regulation of tissue integrity and immunity in skeletal muscle during chronic infection is important for survival of the host, as well as the parasite (22).

In this study, we asked how tissue integrity is maintained during chronic infection. Specifically, we investigated the status of macrophage-mediated tissue repair mechanisms in the tissue environment established by chronic infection. We find chronic infection in mice with T. gondii elaborates extensive muscle damage and prolonged disruption of macrophage homeostasis. In the setting of a highly Th1-polarized environment established during chronic skeletal muscle infection, Tregs are altered and, strikingly, hinder skeletal muscle repair by promoting accumulation of M1. Our results provide evidence for the previously uncharacterized detrimental effects of chronic inflammation on muscle Treg function and their role in skeletal muscle tissue repair.

Materials and Methods

Mice

Female C57BL/6 and B6.SJL (CD45.1) mice were obtained from Taconic Farms (Germantown, NY). C57BL/6-Tg(Foxp3-DTR/EGFP)23.2Spar/Mmjax DEREG mice were obtained from the Jackson Laboratory (Bar Harbor, ME). Foxp3eGFP knock-in reporter mice were obtained from M. Oukka (Seattle Children’s Research Institute). All procedures involving mice were reviewed and approved by the Institutional Animal Care and Use Committee at the University at Buffalo. Mice used were 8–10 wk of age and were matched for age and sex.

Oral infection with T. gondii

Brains from chronically infected mice (30–60 d postinfection [dpi]) were isolated and homogenized in PBS (pH 7.2) for oral infection (23). The total number of RFP-expressing ME49 cysts (graciously provided by Michael Grigg) within the brain homogenate was counted in at least three 20-μl aliquots using a fluorescence microscope. Mice were orally gavaged with five ME49 cysts and monitored by weight and posture daily for the first 14 d of infection and then weekly thereafter.

Muscle histopathology

Skeletal hindlimb muscles were fixed in 10% paraformaldehyde (Sigma-Aldrich, St. Louis, MO). Fixed tissue samples were embedded in paraffin and sectioned at 5 μm for H&E staining. H&E-stained skeletal muscle sections were blindly scored by an independent pathologist for muscle damage based on the degree of immune infiltration by location, necrotic fibers, and centralized nuclei (0 = normal to 3 = severe). Images were taken with an Aperio ScanScope slide scanner. Centralized nuclei per square millimeter were calculated by counting the number of centrally nucleated myofibers on skeletal muscle sections. Muscle tissue cross-sectional area was calculated using ImageScope software (Leica, Wetzlar, Germany).

Muscle functional strength testing

Mice were placed on a screen consisting of 1 × 1-cm squares. The screen was slowly inverted and held two feet above a padded container. Muscle strength was scored based on the time elapsed between full inversion and maximum duration of hanging (maximum 60 s).

Screening of musculoskeletal disease pathways

RNA from naive (n = 5) and chronically infected (n = 5) skeletal muscle tissue was isolated by TRIzol extraction (Thermo Fisher Scientific, Waltham, MA). Reverse transcriptase (iScript; Bio-Rad, Hercules, CA) was used to convert extracted RNA to cDNA. Pooled samples of cDNA from naive mice or chronically infected mice were used as a template (100 ng per sample) for a SYBR Green (SsoAdvanced Universal SYBR Green Supermix; Bio-Rad) quantitative PCR–based screen of 379 unique targets implicated in musculoskeletal disease pathways (PrimePCR Musculoskeletal Diseases Tier 1–4; Bio-Rad). Three hundred and eighty-four–well plates were run on a CFX384 Touch Real-Time PCR Detection System and analyzed using CFX Manager software (both from Bio-Rad).

Isolation of tissue lymphocytes from organ tissues

Mice were euthanized and immediately perfused with PBS (pH 7.2). Skeletal muscle was harvested and minced in digestion media (RPMI 1640, 1% penicillin-streptomycin, 1 mM sodium pyruvate, 0.1% 2-ME, 25 mM HEPES, 150 μg/ml DNase I [Sigma-Aldrich], 50 μg/ml Liberase TL [Roche]). Tissues were digested at 37°C for 55 min and subsequently passed through a 70-μm filter. Mononuclear cells were purified by a Percoll gradient (37.5% Percoll [GE Healthcare, Chicago, IL]/62.5% HBSS) and resuspended in 10% media (RPMI 1640 with 10% FBS, 1% penicillin-streptomycin, 1 mM sodium pyruvate, 0.1% 2-ME, 25 mM HEPES) to create a single-cell suspension.

Spleens were harvested and passed through a 70-μm filter to obtain a single-cell suspension. RBCs were lysed in ACK Lysing Buffer (Lonza, Basel, Switzerland) and resuspended in 10% media. Blood was collected by cardiac puncture and immediately suspended in RPMI 1640 with 10% FBS, 1% penicillin-streptomycin, 1 mM sodium pyruvate, 0.1% 2-ME, 25 mM HEPES, and 5 mM EDTA on ice. RBCs were lysed in ACK Lysing Buffer for 3 min on ice two times. To obtain single-cell suspensions of bone marrow cells, femurs were collected, cleared of muscle and connective tissue, flushed with PBS, and filtered through a 70-μm filter.

Flow cytometric analysis

Single-cell suspensions were stained with HBSS containing extracellular Ab stains and LIVE/DEAD Fixable Aqua Dead Cell Stain (Thermo Fischer Scientific). After extracellular staining, cells were fixed and permeabilized (Intracellular Fixation & Permeabilization Buffer Set; eBioscience, Santa Clara, CA). Then cells were rinsed and stained with Permeabilization Buffer containing intracellular Ab stains (eBioscience). For intracellular stains containing biotinylated Abs, streptavidin staining was performed separately in Permeabilization Buffer (eBioscience) after intracellular staining. Samples were washed and resuspended in flow cytometry buffer/FACS buffer (PBS, 1% BSA [Sigma-Aldrich], 2 mM EDTA [Thermo Fischer Scientific]) for acquisition. Absolute numbers were calculated using CountBright Absolute Counting Beads (Life Technologies).

In vitro restimulation

Lymphocyte single-cell suspensions from tissue were prepared as above and stimulated with 50 ng/ml PMA and 5 μg/ml ionomycin (both from Sigma-Aldrich) in the presence of brefeldin A (GolgiPlug; BD Biosciences, Franklin Lakes, NJ) for 3 h at 37°C in 5% CO2. Extracellular staining was performed as above, and cells were fixed in 2% paraformaldehyde and permeabilized with 0.5% saponin in FACS buffer. Cells were then stained with intracellular Abs and FACS buffer with 0.5% saponin, washed, and resuspended in FACS buffer for flow cytometric acquisition.

T. gondii–specific tetramer staining

Lymphocyte single-cell suspensions from tissue were prepared as above. Allophycocyanin-conjugated MHC class II tetramers bound to T. gondii ME49 hypothetical antigenic peptide I-A(b) (AVEIHRPVPGTAPPS) was obtained through the National Institutes of Health Tetramer Facility (Atlanta, GA). Cells were incubated with the T. gondii–specific tetramer or CLIP tetramer for 1 h, followed by extracellular and intracellular staining as described above.

Abs

The following Abs were used: anti-TCRβ–allophycocyanin–Cy7 (clone H57-597; BD Pharmingen), anti-CD44–BV605 (clone IM7; BD Horizon), anti-CD4–PE–Cy7 (clone RM4-5; BD Pharmingen), anti-CD8β–PE (clone H35-17.2; BD Pharmingen), anti-CD8β–PerCP–Cy5.5 (clone YTS156.7.7; BioLegend, San Diego, CA), anti-Foxp3–FITC (clone FJK-16s; eBioscience), anti-CD25–PerCP–Cy5.5 (clone PC61; BD Pharmingen), anti-CD278–PE–Cy5 (clone 7E.17G9; eBioscience), anti-Tbet–ef660 (clone eBio4B10; eBioscience), anti-Ki67–Alexa Fluor 700 (clone B56; BD Pharmingen), anti-IL-10–allophycocyanin (BD Pharmingen), anti–TNF-α (BD Pharmingen), anti–IFN-γ BV650 (clone XMG1.2; BD Horizon), anti-mAmphiregulin–Biotin (R&D Systems, Minneapolis, MN), anti-Ly6C–PerCp–Cy5.5 (clone HK1.4; eBioscience), anti-CD11b–BV605 (clone M1/70; BD Horizon), anti-Ly6G–PECF94 (clone 1A8; BD Horizon), anti-CD45–V450 (clone 30-F11; BD Horizon), anti-Nos2–Alexa Fluor 488 (clone CXNFT; eBioscience), anti-CD68–PE–Cy7 (clone FA-11; eBioscience), anti-CD206–allophycocyanin (clone C068C2; BioLegend), anti-TCR-β–FITC (clone H57-597; eBioscience), anti-Ly6G–FITC (clone 1A8; BioLegend), anti-NK1.1–FITC (clone PK136; eBioscience), anti-SiglecF–BB515 (clone E50-2440; BD Horizon), anti-TER119–FITC (clone TER-119; BD Pharmingen), anti-CD45–allophycocyanin–eFluor 710 (clone 30-F11; eBioscience), and anti-CD115–BV421 (clone AFS98; BioLegend). Flow cytometry data were acquired using a BD LSRFortessa Cell Analyzer and analyzed using FlowJo version 10.0.8 (TreeStar, Ashland, OR).

Treg adoptive transfer

CD4+CD44–Foxp3– cells were FACS sorted from Foxp3-eGFP knock-in reporter mice. Isolated cells were stimulated with 1 μg/ml plate bound anti-CD3/anti-CD28 (eBioscience) for 48 h in conditioning media. After the initial 24 h, 100 U/ml rIL-2 (PeproTech, Rocky Hill, NJ) was added to the wells/conditioning media. To generate nonpolarized Tregs, conditioning media contained 10 ng/ml TGF-β and 5 μg/ml anti–IL-4–blocking Abs and anti–IFN-γ–blocking Abs (BD Pharmingen). To generate Th1-polarized Tregs (Th1-Tregs), conditioning media contained 10 ng/ml TGF-β (PeproTech), 5 μg/ml anti-IL-4–blocking Abs, and 50 ng/ml IFN-γ (PeproTech). Cells were expanded in vitro for a total of 8 d. HBSS, 7.5 × 105 nonpolarized Tregs, or Th1-Tregs were adoptively transferred i.v. into CD45+ B6.SJL mice at 23 dpi. Organs were harvested for analysis 6 d following adoptive transfer.

Treg-specific depletion in DEREG mice

DEREG mice were orally infected with ME49 cysts. Starting at 23 dpi, DTR+ and DTR− mice were treated i.p. with 1 μg of diphtheria toxin (DT) for a total of three injections every other day. Organs were harvested for analysis 1 d following the last injection.

Quantitative RT-PCR of muscle differentiation factors

Tissues from experimental mice were isolated and preserved in RNAlater (QIAGEN, Venlo, the Netherlands) for RNA isolation by TRIzol extraction. Isolated RNA was converted to cDNA (iScript; BD Biosciences) and assayed for muscle differentiation factors MyoD and myogenin by real-time PCR (iTaq Universal SYBR Green Supermix; BD Biosciences) using primers purchased from Bio-Rad (PrimePCR assay). Ct values were normalized to the housekeeping gene 18s rRNA (forward: 5′-GCAATTATTCCCCATGAACG-3′, reverse: 5′-GGGACTTAATCAACGCAAGC-3′). A total of 100 ng of skeletal muscle RNA was used for quantitative RT-PCR (qRT-PCR) analysis.

Quantification of parasite burden

Tissues from experimental mice were isolated and preserved in RNAlater (QIAGEN) for DNA extraction with the QIAGEN DNeasy Blood & Tissue Kit or RNA isolation with TRIzol Reagent (Life Technologies). Total parasite burden was quantified by PCR amplification of a T. gondii–specific gene, B1, from DNA isolated from target tissues (forward: 5′-TCCCCTCTGCTGGCGAAAAGT-3′, reverse: 5′-AGCGTTCGTGGTCAACTATCGATTG-3′). Ct values were compared with a standard curve constructed from B1 amplification of known T. gondii genomic DNA concentrations. Tachyzoite and bradyzoite parasite burden was quantified by qRT-PCR of Eno2 (forward: 5′-CCGTTACTCAACTTCCAACA-3′, reverse: 5′-CCATCGGTCAACAAGTCAA-3′) and Bag1 (forward: 5′-GGGATGTACCAAGCATCCTG-3′, reverse: 5′-AGGGTAGTACGCCAGAGCAA-3′), respectively. Ct values were compared with a standard curve constructed from Eno2 or Bag1 amplification of known T. gondii RNA concentrations. A total of 1 μg of skeletal muscle DNA or RNA was used for PCR analysis of parasite burden.

Statistics

All statistics were generated using GraphPad Prism v6.0c (GraphPad, La Jolla, CA).

Results

T. gondii infection causes myositis and loss of function in skeletal muscle

Following oral infection with five type II ME49 T. gondii cysts, C57BL/6 mice experience acute significant weight loss at 9–12 dpi and fail to recover weight, similar to that of their naive counterparts as chronic infection (day 30) is established (Fig. 1A). We speculated that the inability to efficiently regain weight was due, in part, to ongoing skeletal muscle damage following acute infection. To this end, we compared inflammatory cell infiltration and muscle fiber damage in mock- and parasite-infected mice by H&E staining (Fig. 1B, 1C). Specifically, analysis of cellular damage elaborated during infection shows little inflammatory infiltrate in muscle at day 12; however, by day 30 postinfection, extensive inflammatory infiltrates were observed in perivascular, perimysial, and endomysial locations (Fig. 1B, 1C). Prenecrotic hyaline and necrotic fibers were comparably increased at days 12 and 30 postinfection (Fig. 1B, 1C). Additionally, there were more myofibers with central nuclei and prominent nucleoli, indicators of regenerating fibers, at 30 dpi (Fig. 1B, 1C). To determine whether this histologic damage resulted in physiological consequences and alterations in muscle function, we measured muscle strength in infected mice using an inverted screen test. Chronically infected mice (>day 30) displayed significant muscle weakness compared with their naive counterparts (Fig. 1D).

FIGURE 1.
FIGURE 1.

T. gondii causes myositis and skeletal muscle damage. (A) C57BL/6 mice were orally infected with five ME49 T. gondii cysts. Body weight was monitored for up to 35 dpi. (B) Representative images of H&E-stained skeletal muscle sections from naive and infected (30 dpi) mice. (C) Blinded histopathological scoring of muscle damage and inflammation in naive and infected (12 and 30 dpi) skeletal muscle. (D) Functional muscle strength was quantified in naive and infected (>30 dpi) mice by measuring hang time using Kondziela’s inverted screen test. Al results are representative of at least two experiments with n = 4 mice per group per experiment; error bars represent SD. ****p < 0.0001, Student t test.

To investigate myositis elicited by T. gondii in skeletal muscle, we next characterized the immunological landscape of infected skeletal muscle. To this end, we analyzed whole-tissue RNA from naive and chronically infected skeletal muscle for differential expression of gene targets and pathways previously implicated in musculoskeletal diseases. A total of 379 unique targets was screened by qRT-PCR using a predesigned musculoskeletal disease pathway panel (Bio-Rad). A Th1-driven immune response is protective during acute and chronic stages (19, 24–27). As expected, we found a robust increase in whole-tissue expression of Th1 cytokines, including Il12a, il1a, il2, il12b, and Ifng, in day-30 infected muscle that was below the level of detection in naive skeletal muscle (Fig. 2A, left panel). Of note, transcripts encoding proteins involved in immunomodulatory pathways, such as il1rn, il10, and il10ra, in day-30 infected muscle are expressed at levels several fold greater than those seen in naive skeletal muscle (Fig. 2A, right panel). The data from this screen describe an environment in which Th1-inflammatory pathways are active despite a concurrent increase in immunomodulatory mechanism, ultimately propagating unresolved inflammation. Furthermore, at the cellular level, total CD4+ and CD8+ T cells dramatically increase in infected muscle, whereas muscle from naive controls shows few infiltrating T cells (Fig. 2B, 2C). Muscle CD4+ and CD8+ effector T cells highly express the Th1 canonical transcription factor Tbet (Fig. 2D). Upon restimulation, both CD4+ and CD8+ effectors from day-24 infected muscle produce IFN-γ but not IL-17A (Fig. 2E, 2F). Notably, these populations persist in the infected muscle, as evidenced by the continued expression of high levels of Tbet and the production of IFN-γ that was detectable directly ex vivo at 60 dpi (Fig. 2G).

FIGURE 2.
FIGURE 2.

Inflammatory landscape of infected skeletal muscle is highly Th1 polarized. (A) Inflammatory targets involved in known musculoskeletal disease pathways were explored by screening RNA isolated from the skeletal muscle of naive mice and mice at 30 dpi against 379 unique targets by qRT-PCR. Heat map generated from CT values of targets normalized to housekeeping (GAPDH) from naive and infected mice (left panel). Relative gene expression of infected skeletal muscle targets normalized to housekeeping and naive skeletal muscle (right panel). Statistics were calculated based on log-transformed fold change values. Quantification of TCRβ+CD4+ T cells (B) and TCRβ+CD8+ T cells (C) in spleen and skeletal muscle postinfection. (D) Representative FACS plots of Tbet expression by Tconvs (TCRβ+CD4+Foxp3−CD44+) in naive and infected muscle (30 dpi). Representative FACS plots of IL-17 and IFN-γ production by Tconvs (E) and CD44+CD8+ T cells (F) from naive and infected muscle following 3 h of PMA/ionomycin stimulation (24 dpi). (G) Representative FACS plots of Tbet and IFN-γ by Tconvs from naive and infected muscle ex vivo (60 dpi). (A–F) Results are representative of at least three experiments with n = 4 mice per group per experiment; error bars represent SD. **p < 0.01, ***p < 0.001, ****p < 0.0001, ANOVA with Tukey multiple-comparison test (A), Student t test (B and C).

Long-lasting alterations in skeletal muscle macrophage populations during infection

Because we observed such striking muscle damage and loss of function, we next investigated macrophage polarization in infected muscle, because its coordination is essential for proper muscle fiber regeneration (4). Consistent with previous reports, naive muscle consists mostly of M2-polarized macrophages (∼80%) (28) (Fig. 3A–C). By day 12 postinfection, we observe an increase in IM/M1-polarized macrophages and a dramatic reduction in M2, which markedly shifts the ratio of IM/M1 to M2 heavily in favor of IM/M1 (Fig. 3A–C). By 24 d postinfection, muscle macrophages express M2 surface markers, such as CD206, and the absolute number of M2 increases (Fig. 3A–C). Notably, they may not be fully functionally polarized to the M2 phenotype because they continue to express iNOS, an IM/M1 feature (Fig. 3D). Furthermore, this increase in M2 is not paralleled by a contraction of IM/M1, as typically seen during skeletal muscle repair from sterile injury. Rather, IM/M1 accumulation is sustained as chronic infection is established (Fig. 3C). We postulated that this altered macrophage polarization and/or function results in impairment of muscle regeneration leading to ongoing muscle damage.

FIGURE 3.
FIGURE 3.

T. gondii infection expands IM/M1 populations. (A) IM/M1 and M2 were gated on CD45+CD11b+Ly6G−CD68+ cells. Frequencies (B) and absolute numbers (C) of IM/M1 (Ly6ChiCD206lo), M2 (Ly6CloCD206hi), and resting macrophages (Ly6CloCD206lo) in naive and infected muscle. (A and B) Results are representative of at least two experiments with n ≥ 4 mice per group per experiment; error bars represent SD. (D) Representative FACS plots and graphical representation of iNOS expression by muscle IM/M1 and M2 in naive and infected mice. Results are representative of at least four experiments with n = 4 mice per group per experiment. **p < 0.01, ***p < 0.001, ****p < 0.0001, ANOVA with Tukey multiple-comparison test (C), Mann–Whitney U test (D).

T. gondii infection results in sustained reduction of Treg frequencies throughout infection

Recent studies of models of sterile muscle injury and muscular dystrophy showed that, during proper muscle repair, Tregs are required to promote the shift in macrophage populations from proinflammatory IM/M1 to M2 (12, 13). Critically, the emergence of Tregs in muscle after sterile injury coincides with the transition from IM/M1- to M2-mediated phases of repair (12, 13). Previous studies described a collapse of the Treg compartment locally in the gastrointestinal tract, as well as systemically, during acute lethal infection with T. gondii (23, 29, 30). Decreased Treg numbers are associated with limited Treg conversion and a lack of IL-2 availability in the gut (23). However, the long-term dynamics of the Treg compartment, especially the muscle, have not been investigated. During infection, we observe a steady increase in the number of Tregs during early infection that was accompanied by increased proliferative potential at day 24 of infection (Fig. 4A). However, the substantial infiltration of CD4+Foxp3− conventional T cells (Tconvs) and CD8+ T cells into the muscle results in significantly reduced proportions of Tregs/Tconvs (<6% Tregs) during chronic infection (Fig. 4A). These findings are in striking contrast to the aforementioned sterile injury model of muscle damage in which Tregs represent ∼40% of all CD4+ T cells (12). This is significant because relatively fewer Tregs are available to regulate the expanded effector populations.

FIGURE 4.
FIGURE 4.

Treg kinetics and phenotype during chronic infection. (A) Quantification of Treg (TCRβ+CD4+Foxp3+) frequency, absolute number, and proliferation (KI67+) during infection. Representative FACS plots (B) and graphical summary (C) of CD25 and ICOS expression in Tregs from naive and infected (24 dpi) muscle. (D) Representative FACS plots and graphical summary of Tbet expression in skeletal muscle Tregs during infection (left and middle panels). Tbet mean fluorescence intensity in Tconvs and Tregs at 30 dpi (right panel). (E) Representative FACS plots and graphical summaries of Ag specific CD44+CD4+ Tconvs and Tregs in the spleen and skeletal muscle during long-term infection, as detected by staining with MHC class II tetramers loaded with T. gondii antigenic peptide. (F) FACS plots and graphical representation of Areg expression by Tregs isolated from naive and infected (24 dpi) muscle following 3 h of PMA/ionomycin (P/I) restimulation. Results are representative of at least three experiments with n ≥ 4 mice per group per experiment; error bars represent SD. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001, Kruskal–Wallis test. ns, not significant.

Tregs can be divided into central memory (CD25hi) and effector memory (ICOShi) populations based on the expression of a series of markers (31). We analyzed the expression of these proteins and found high levels of CD25 expression at day 12 that were not sustained during later time points. Instead, ICOS expression is favored by muscle Tregs (Fig. 4B, 4C). Also, phenotypically, a large proportion of these Tregs express the Th1-associated transcription factor Tbet (Fig. 4D). Of these Tbet+ Tregs, the per-cell expression of Tbet is comparable to that of Th1 (Fig. 4D). Furthermore, in contrast to Tconvs, a durable population of T. gondii Ag-specific Tregs is not detected in the spleen or skeletal muscle during infection (Fig. 4E).

As a readout of function, we assessed expression of various pro- and anti-inflammatory cytokines by muscle Tregs from day-24 infected mice by restimulating with the mitogen PMA and ionomycin. Stimulated muscle Tregs did not produce IL-10 (data not shown). Previously, it was shown muscle Tregs express high levels of the epidermal growth factor family member amphiregulin (Areg); this expression is thought to promote muscle fiber regeneration by acting on stem cells in the skeletal muscle (12). We find infection increases Areg expression, and restimulation increases that expression (Fig. 4F). We next assessed proinflammatory cytokine IFN-γ from muscle Tregs in response to restimulation and did not find skeletal muscle Tregs producing IFN-γ (Fig. 4F).

Transfer of Th1-iTregs promotes IM/M1 accumulation in infected skeletal muscle

We postulated that the reduced proportion of Tregs in infected muscle was inadequate to promote efficient transition from IM/M1 to M2 during chronic infection, resulting in the observed accumulation of IM/M1. Furthermore, chronic exposure to IFN-γ was shown to sensitize Tregs to IL-12 signaling and potentially promote IFN-γ production by Tregs (23, 32), which could hinder their ability to promote M2 differentiation. To test whether insufficient numbers of Tregs or their chronic exposure to IFN-γ results in the accumulation of IM/M1 during infection, we adoptively transferred inducible Tregs (iTregs) cultured in the absence or presence (Th1-iTreg) of rIFN-γ for 7–9 d. To generate iTregs for transfer, we FACS sorted CD4+Foxp3− T cells from Foxp3eGFP reporter mice, converted them to iTregs in the presence of TGF-β, and expanded/conditioned them in vitro for 7–9 d. iTregs and Th1-iTregs were sorted on the basis of GFP+ (Foxp3+) cells after culture to obtain a purified population for transfer into infected mice (23 dpi) (>98%, data not shown). Day 23 of infection was chosen as the transfer date because M2 populations re-emerge and Tregs are actively proliferating at day 24 postinfection (Figs. 3C, 4A). Skeletal muscle was harvested 6 d posttransfer to monitor IM/M1 and M2. Transfer of iTregs did not significantly alter IM/M1 and M2 numbers and proportions, and proliferative capacity remained similar to transfer controls (Fig. 5). Surprisingly, Th1-iTregs adoptively transferred into mice at 23 dpi increase IM/M1 in the skeletal muscle (Fig. 5A–C). This increase was not due to the action of transferred Th1-iTregs on in situ IM/M1 proliferation (Fig. 5D). Transfer of 7.5 × 105 iTregs or Th1-iTregs did not appreciably alter the absolute number of skeletal muscle Treg (Supplemental Fig. 2A). Furthermore, these changes occurred in the absence of enhanced immunopathology, as assessed by elevated levels of liver enzymes (aspartate transaminase/alanine transaminase) (Supplemental Fig. 2B). To better understand how transfer of iTregs and Th1-iTregs results in differential effects on the macrophage population, we assessed their ability to produce a variety of cytokines following stimulation with PMA/ionomycin after culture. Notably, no differences were observed with regard to their capacity to produce IL-4, IL-13, IL-17, or IL-10 (Supplemental Fig. 1A, 1B). iTregs and Th1-iTregs produced Areg (Supplemental Fig. 1B). Although there was a small, but statistically significant, increase in the ability of Th1-iTregs to produce IFN-γ, IFN-γ–producing Th1-iTregs still only represent ∼2% of total Th1-iTregs (Supplemental Fig. 1C). Our data suggest that Th1-iTreg IFN-γ production is likely not the distinguishing factor promoting IM/M1 accumulation in the infected skeletal muscle. Together, our data suggest that long-term exposure of Tregs to IFN-γ may alter their canonical function and lead to the accumulation of IM/M1 instead of M2.

FIGURE 5.
FIGURE 5.

Th1-iTreg transfer increases M1 populations. HBSS, iTregs, or Th1-iTregs were adoptively transferred into infected mice at 23 dpi. Representative FACS plots (A), distribution (B), and absolute number (C) of IM/M1, M2, and resting macrophages 6 d after cell transfer. (D) Effect of Treg transfer on skeletal muscle M1 and M2 proliferation (Ki67+) 6 d following transfer. Results are representative of n ≥ 5 per group from two experiments, error bars represent SD. *p < 0.05, Student t test.

Treg ablation shifts macrophage populations in favor of M2 and rescues muscle regeneration

Given the remarkable finding that Th1-iTregs enhance IM/M1 numbers in infected muscle, we asked whether skeletal muscle Tregs were directly contributing to the pathology observed during chronic infection. To this end, we used transgenic mice expressing the simian DT receptor driven by Foxp3 (DTR) to systemically deplete Tregs at 23 dpi. DTR− and DTR+ mice were infected and treated with DT starting on day 23, and depletion of skeletal muscle Tregs was confirmed (Supplemental Fig. 2C). Given that adoptive transfer of Th1-iTregs increased IM/M1 (Fig. 5C), we asked whether Treg depletion restores M2 proportions, leading to improved repair. Indeed, Treg ablation (DTR+) increases muscle M2 proportions, whereas DT treatment in DTR− mice had no effect (Fig. 6A, 6B). The shift toward a statistically significant increase in the M2 to IM/M1 ratio was due to a combination of a decrease in the absolute numbers of IM/M1 and an increase in the absolute numbers of M2 (Fig. 6C). Furthermore, the effects of Treg depletion on IM/M1 and M2 numbers occurred in the absence of increased immunopathology, as assayed by serum aspartate transaminase/alanine transaminase levels and total T cell/IFN-γ–producing T cells in the skeletal muscle (Supplemental Fig. 2D, 2E). The frequency and number of total iNOS-producing macrophages remained unaltered following Treg depletion (Supplemental Fig. 2F). However, reactive nitrogen species may not be the key agent inciting damage, and the re-emergence of macrophages expressing M2 phenotypic markers, such as the scavenger receptor CD206, indicates that an increased proportion of macrophages may have enhanced regenerative functions, despite also producing iNOS.

FIGURE 6.
FIGURE 6.

Treg depletion is accompanied by a shift toward increased M2. Systemic depletion of Tregs following DT treatment of DTR-transgenic mice. Representative FACS plots (A), distribution (B), and absolute number and fold change (C) of IM/M1 and M2 1 d following the final DT treatment. (D) Effect of Treg depletion on skeletal muscle M1 and M2 proliferation (Ki67+) 1 d following the final DT treatment. (A–D) Results are representative of n = 10 per group from four experiments. (E) Gating scheme for Ly6chi and Ly6clow monocytes. Frequencies and absolute numbers of bone marrow (F) and blood (G) Ly6chi and Ly6clow monocytes (CD45+TCRβ−Ly6G−SiglecF−NK1.1−TER119−CD11b+CD115+) 1 d following the final DT treatment. Results are representative of n = 13 per group from three experiments; error bars represent SD. *p < 0.05, **p < 0.01, Kruskal–Wallis test (C, E, and F), Student t test (D–F).

To understand how Treg ablation influences the macrophage compartment, we first investigated the in situ proliferation of IM/M1 and M2 following depletion. Ki67 expression in IM/M1 and M2 remained unaltered (Fig. 6D). We next asked whether Treg ablation acted on a systemic level, influencing M1 and M2 precursor monocytes from the bone marrow. Specifically, it was reported that Ly6chi and Ly6clow monocytes preferentially give rise to M1 and M2, respectively (4, 33–36) (gating scheme in Fig. 6E). Following Treg depletion during infection, the frequency of Ly6chi and Ly6clow monocytes is not notably altered (Fig. 6F). Interestingly, the absolute number of Ly6clow monocytes is significantly increased, whereas the number of Ly6chi monocytes trends upward (Fig. 6F). We next asked whether this change was also reflected in the blood; however, no detectable changes were observed in the frequency or number of monocyte subsets (Fig. 6G).

Strikingly, the depletion of Tregs is associated with an increased incidence of centrally nucleated muscle fibers demonstrating enhanced muscle regeneration (Fig. 7A, 7B), suggesting that Tregs actively inhibit muscle regeneration during chronic infection. We did not observe alterations in the weights of mice during DT treatment (Supplemental Fig. 2G). Previously, it was demonstrated that M2 promote the early, proliferative stage of myogenesis following sterile injury (37). To assess whether increased proportions of M2 influence the myogenic processes, we measured levels of the transcription factors MyoD and myogenin, which regulate distinct phases of the myogenic-differentiation program in skeletal muscle stem cell populations. MyoD is highly expressed in the mid-G1 phase and between the S and M phases of the cell cycle (38). Myogenin is expressed upon differentiation of myoblasts into multinucleated myotubes (38). Although tissue level expression of MyoD is not significantly altered in infected DTR+ mice compared with DTR− mice, myogenin expression decreases following Treg depletion (Fig. 7C).

FIGURE 7.
FIGURE 7.

Treg ablation rescues skeletal muscle fiber regeneration. (A) Representative H&E stains of hindlimb muscles 1 d following final DT treatment in DTR− and DTR+ mice 28 dpi. Arrowheads indicate centrally nucleated myofibers. (B) Quantification of regenerating fibers (centrally nucleated myofibers) in muscle sections per square millimeter. (C) Fold expression of myogenic targets MyoD and myogenin in infected DTR+ skeletal muscle normalized to DTR− skeletal muscle following DT treatment. Statistics were calculated based on log-transformed fold change values. qRT-PCR quantification of total parasite burden by B1 (D) and parasite stage-specific transcripts Eno2 tachyzoite (left panel) and Bag1 bradyzoite (right panel) (E) following DT treatment. Results are representative of n = 10 per group from four experiments; error bars represent SD. *p < 0.05, Student t test.

Further, it was demonstrated that M2 have anticyst activity (39). We assessed whether a shift in proportions to favor M2 improved regeneration, in part through decreasing parasite burden. Total parasite burden was assessed by the T. gondii tandem-arrayed 35-fold repetitive gene, B1. Treg depletion resulted in a nonsignificant downward trend in global parasite burden (Fig. 7D). Transcripts Eno2 (tachyzoite) and Bag1 (bradyzoite) were also measured to determine whether changes in life stages of the parasite occur following Treg depletion. Although detection of Eno2 does not change significantly, expression of Bag1 decreases (Fig. 7E). Collectively, our data show that chronic T. gondii infection causes ongoing inflammation and muscle damage. Furthermore, skeletal muscle Tregs in T. gondii–infected mice directly contribute to muscle pathology by hindering muscle regeneration through promoting IM/M1 accumulation and, consequently, influencing the myogenic program.

Discussion

In recent years, great progress has been made in examining the role of inflammation in sterile muscle injury and genetic diseases. These studies showed that muscle repair is highly dependent on the sequential and discrete actions of M1 and M2 (1–3). Importantly, dysregulation of macrophage populations and the inability to repair tissue damage are central to the pathogenesis of many inflammatory myopathies. In many myopathies, including polymyositis and dermatomyositis, inappropriate macrophage accumulation and activation lead to invasion and destruction of nonnecrotic muscle fibers (7, 8, 16) Notably, a prolonged M1 presence during muscle injury may lead to delayed or aberrant repair (7–13), whereas exuberant M2 activity may cause muscle fibrosis (6).

Previous reports demonstrated sensorimotor deficits during T. gondii infection; however; these findings were attributed solely to neurologic abnormalities (40–42). Our data compellingly show muscle damage, ongoing myositis, and loss of muscle function within infected muscle. This previously underappreciated pathology is associated with prolonged accumulation of IM/M1 in skeletal muscle during infection. Furthermore, it appears that M2 polarization is aberrant, because we observe iNOS expression by these cells. A previous report demonstrated that macrophages exposed to T. gondii express features of M1 and M2 (43). Recent studies show a tissue-specific role for resident Tregs in the critical transition of proinflammatory IM/M1 to M2 during the tissue repair process (12, 13). Surprisingly, we find that Tregs acquire a pathogenic capacity to promote skeletal muscle damage during chronic toxoplasma infection.

The ability to adapt to local inflammatory environments is important for Treg homeostasis and function (30, 44). It is known that exposure to Th1 inflammation endows Tregs with an enhanced capacity to traffic to and survive within local Th1-inflamed environments through acquiring Th1 effector traits, such as Tbet and CXCR3 expression (30, 45). Furthermore, transcriptomic analysis of Tregs cultured in Th1 environments identifies Tregs with hallmark features like Tbet and CXCR3 as phenotypically distinct from Tregs cultured in neutral conditions (30). Consistent with this, we show that, in the Th1 environment of chronically infected skeletal muscle, Tregs highly express Tbet. However, little is known about the long-term consequences of extended exposure to a proinflammatory milieu on Treg physiology, such as that experienced during chronic inflammatory disease states. We show in this article that there can be detrimental effects on the ability of Tregs to continue to perform homeostatic functions in the face of long-term inflammation. However, it remains to be determined whether Tbet directly plays a role in the pathogenic capacity of these Th1-Tregs. Taken together with our current data, the paradigm of repair must be expanded to account for how pre-existing inflammation may modulate Treg physiology and ultimately alter progression of the repair process.

The role of Tregs during chronic infection with T. gondii was suggested to be limited during late-stage infection (46, 47). Studies showed that depletion of CD25-expressing cells with the anti-CD25 Ab administered after 2 mo of infection did not affect the course of disease. Interestingly, previous studies and our data highlight that Tregs found in chronic infection express low levels of CD25 compared with Tregs during acute infection (23, 46, 48) (Fig. 4). Our study provides compelling evidence that Tregs do indeed participate in the immune response during chronic infection, albeit in a proinflammatory role instead of a suppressive role. Our results show that these Tregs modulate the IM/M1 and M2 balance by promoting the accumulation of IM/M1 in the skeletal muscle. This role is independent of IFN-γ production by Tregs because we do not find that Tregs produce IFN-γ ex vivo or with mitogenic stimulation. This is in contrast to previous studies that showed Foxp3-expressing cells producing the proinflammatory cytokine IFN-γ during acute lethal infection with T. gondii (23).

We show that ablation of pathogenic Tregs induced during infection coincides with increases in the proportion of M2 accompanied by increases in the number of regenerating fibers. We did not observe an appreciable decrease in the production of iNOS by M2, suggesting that reactive nitrogen species may not be the key agent inciting damage because we observe a dramatic increase in muscle fiber regeneration. Treg depletion does not alter in situ proliferation of IM/M1 or M2. However, we do observe increased numbers of Ly6clow monocytes in the bone marrow. Although this increase is not reflected in the blood, increased extravasation into the tissue may counter the increased input of Ly6clow monocytes into the blood. Alternatively, the increase in Ly6clow monocytes may represent an accumulation in the bone marrow resulting from hindered egress and therefore, the effect of Treg depletion is at the level of the muscle IM/M1 and M2. In either circumstance, our results indicate a correlative relationship among the development of pathogenic Tregs during chronic infection, altered tissue macrophage homeostasis, and impaired muscle regeneration.

The results presented are consistent with previous in vitro coculture studies in which M2 stimulated myoblast proliferation (37). Earlier reports note that this effect of M2 is not reliant on myoblast MyoD or myogenin expression (37). We show that enhanced proportions of M2 following Treg depletion correlate with increases in regenerating fibers concurrent with a decrease in myogenin expression. Although myogenin is important for myotube terminal differentiation and fusion, its expression was shown to inhibit myoblast responsiveness to epidermal growth factors (49). Additionally, decreased myogenin expression may represent a myogenic checkpoint to ensure genetic stability in differentiated cells and proper regeneration (50). Taken together, these data suggest that pathogenic Tregs may cause a defective myogenic differentiation program through modulating macrophage subsets.

Alternatively, it is possible that Treg interactions with satellite cells are involved (12, 51). During sterile injury it was shown that Treg recruitment only occurred during the time when satellite cells expand. Furthermore, in vitro studies showed that iTregs enhanced proliferation of satellite cells while preventing their further differentiation (12, 51). It is unclear whether this prevention of differentiation occurs in vivo. Furthermore, Tregs in injured muscle show marked increases in the production of epidermal growth factor Areg that are concurrent with M2 accumulation and proliferation of satellite cells (12). We find that Tregs from infected skeletal muscle express higher levels of Areg than do naive skeletal muscle Tregs, although they do not express as much as Tregs in a sterile injured muscle where more than half of the population produced Areg (12). Given that Treg ablation leads to dramatic increases in regenerating fibers, our data suggest that the regenerative program during T. gondii infection is not reliant on Areg production by Tregs.

Our studies highlight the complexity of the potential interactions among the parasite, skeletal muscle fiber, and immune response. Although the depletion of Tregs may affect total parasitic activity, we do not observe significant overall alterations in the parasite burden following depletion, as measured by the B1 gene. Previous studies showed that M2 have anticyst properties during T. gondii infection in the brain (39). Interestingly, we show that, following Treg depletion, increases in M2 proportions are associated with decreased expression of the bradyzoite gene bag1. Conversely, we do not detect alterations in tachyzoite-specific gene expression, as evidenced by nonsignificant changes in the eno2 transcript levels between Treg- replete and -depleted mice. Host cell differentiation status can also directly influence parasite growth. Notably, terminally differentiated syncytial myotubes promote stage conversion to bradyzoites (22). Taken together, these findings raise intriguing questions about whether the parasite may be influencing the local environment to induce pathogenic Tregs as a means to remain encysted, evade immune detection, and increase the likelihood of transmission via the infectious cyst stage because ingestion of undercooked contaminated meats is the primary route of transmission. In addition, it is tempting to speculate that lasting loss of muscle function may have an impact on an organismal level, such that the host cannot escape predators and, thus, the parasite is more likely to be transmitted.

Although the importance of innate cells to muscle fiber regeneration has been well characterized, the role of the adaptive immune response is still being explored. Moreover, the regulatory mechanisms that allow for the progression of the wound repair response have just recently begun to be elucidated. The critical role that Tregs play in promoting tissue repair mechanisms in response to damage of the skeletal muscle was recently recognized (12, 13, 51). The interplay between these cell populations during chronic infection of T. gondii has not been examined until now. Previous work showed that, in the gut and other tissues, Tregs have a protective role during T. gondii infection (23, 30). Our data show the unexpected finding that Tregs promote muscle damage during chronic infection. As Treg-directed therapies gain traction, a deeper understanding of the consequences of chronic inflammation on Treg physiology and tissue-specific reparative programing will be a powerful asset in producing safe and efficacious therapies. Our results show that Tregs residing in infected skeletal muscle play a decidedly different role than what is normally the suppressive function associated with Foxp3-expressing cells. Identifying the determinants of immunoregulation during skeletal muscle infection will provide targets for novel immune-modulatory therapeutics for a range of inflammatory myopathy etiologies, not limited to infection.

Disclosures

The authors have no financial conflicts of interest.

Acknowledgments

We thank the University at Buffalo Histology Core and the Confocal Microscopy and Flow Cytometry Core Facility at the University at Buffalo for technical assistance, the National Institutes of Health Tetramer Core Facility for the T. gondii ME49 hypothetical protein tetramers, and Dr. Yasmine Belkaid, Dr. Joanne Konkel, and Dr. John Grainger for discussion and critical reading of the manuscript.

Footnotes

  • This work was supported by the Jacobs School of Medicine and Biomedical Sciences, University at Buffalo, State University of New York (to E.A.W.) and by National Institutes of Health Grants AI124677 (to I.J.B.) and AI007614 (to R.M.J.).

  • The online version of this article contains supplemental material.

  • Abbreviations used in this article:

    Areg
    amphiregulin
    dpi
    d postinfection
    DT
    diphtheria toxin
    DTR
    DT receptor driven by Foxp3
    IM
    inflammatory monocyte
    iNOS
    inducible NO synthase
    iTreg
    inducible Treg
    M1
    proinflammatory macrophage
    M2
    proregenerative macrophage
    qRT-PCR
    quantitative RT-PCR
    Tconv
    conventional T cell
    Th1-iTreg
    Th1-polarized inducible Treg
    Th1-Treg
    Th1-polarized Treg
    Treg
    regulatory T cell.

  • Received May 26, 2016.
  • Accepted November 1, 2016.

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