Induction of thymic atrophy and loss of thymic output by type-I interferons during chronic viral infection

Abstract

Type-I interferon (IFN-I) signals exert a critical role in disease progression during viral infections. However, the immunoodulatory mechanisms by which IFN-I dictates disease outcomes remain to be fully defined. Here we report that IFN-I signals mediate thymic atrophy in viral infections, with more severe and prolonged loss of thymic output and unique kinetics and subtypes of IFN-α/β expression in chronic infection compared to acute infection. Loss of thymic output was linked to inhibition of early stages of thymopoiesis (DN1-DN2 transition, and DN3 proliferation) and pronounced apoptosis during the late DP stage. Notably, infection-associated thymic defects were largely abogated upon ablation of IFNαBR and partially mitigated in the absence of CD8 T cells, thus implicating direct as well as indirect effects of IFN-I on thymocytes. These findings provide mechanistic underpinnings for immunotherapeutic strategies targeting IFN-1 signals to manipulate disease outcomes during chronic infections and cancers.

INTRODUCTION

Persistent viral infections such as HIV (Katlama et al., 2013) and hepatitis C virus HCV) (Lauer, 2013) pose a major public health challenge. Although a strong antiviral immune response is initiated early after infection, induction of an immunosuppressive program subsequently leads to viral persistence (Barber et al., 2006; Brockman et al., 2009; Brooks et al., 2006; Day et al., 2006; Trautmann et al., 2006). Thus, a better understanding of the underlying mechanisms of immunosuppression is key to treatment of chronic viral infections.

Type-I interferon (IFN-I) signals are associated with disease progression in persistent infections (d’Ettorre et al., 2011), with evident timing-dependent effects of IFN-I administration or blockade (Elsaesser et al., 2020; Soper et al., 2017; Sullivan et al., 2015; Teijaro et al., 2013; Wang et al., 2012; Wilson et al., 2013). Chronic HIV infection is associated with high levels of IFN-I (Biron, 1998). Similarly, HCV-infected patients also exhibit an elevated IFN-I signature (Guidotti and Chisari, 2006; Su et al., 2002). It is notable that in experimental SIV infections of monkeys, elevated IFN-I levels and associated inflammatory signatures are found specifically in rhesus macaques that develop disease (Jacquelin et al., 2009; Manches and Bhardwaj, 2009), and not in sooty mangabeys. or African green monkeys (Bosinger et al., 2009), which manifest modest disease pathology despite similar viral loads. In mice also, IFN-I signaling during persistent infection with lymphocytic choriomeningitis virus (LCMV) is associated with disruption of lymphoid architecture and hyperimmune activation (Elsaesser et al., 2020; Teijaro et al., 2013; Wilson et al., 2013).

Mechanistically, IFN-I signals have been found to exert myriad effects on antiviral immunity such as: (i) direct antiviral activity and the induction of apoptosis in infected cells (Guidotti and Chisari, 2001; McNally et al., 2001; Tanaka et al., 1998); (ii) increased MHC-I expression on DCs to promote presentation of virally encoded epitopes to cytotoxic T cells (Cella et al., 1999); (iii) enhanced T cell clonal expansion during the acute phase of infections (Dondi et al., 2004; Kolumam et al., 2005); (iv) development of long-lived T cell memory (Grakoui et al., 2003; Kolumam et al., 2005; Lechner et al., 2000; McMichael and Rowland-Jones, 2001; Rickinson and Moss, 1997; Thimme et al., 2003). Contrary to these immunostimulatory antiviral effects of IFN-I, blockade of IFN-I signals before or after establishment of persistent lymphocytic choriomeningitis virus infection has been shown to promote viral clearance by lifting immune suppression through a CD4 T cell-dependent mechanism (Teijaro et al., 2013; Wilson et al., 2013). These studies identified that IFN-I signaling was essential for the expression of negative immune regulators IL-10 and PDL-1, in addition to exerting antiproliferative activities (Biron and Brossay, 2001; Samuel, 2001; Taniguchi and Takaoka, 2002).

While much mechanistic insight has been gained into the immunostimulatory role of IFN-I, relatively less is known about its immunosuppressive effects, as well as the respective contribution of the various IFN-α subtypes to a protective or deleterious immune response. In the present study, we sought to define the impact of IFN-I signaling on thymic function during chronic viral infection. T cell development and thymic output were defined in the LCMV model of acute and chronic viral infections. IFN-I or inducers of these molecules have been shown to play a detrimental role at numerous steps of T cell development, such as (i) accumulation of T cell precursors at the DN1-DN2 transition, (ii) inhibition of the DN3–4 proliferative stages, (iii) induction of DP subset apoptosis and (iv) impairment of T cell selection (Baron et al., 2008; Demoulins et al., 2008; Keir et al., 2002a; Keir et al., 2002b; Lin et al., 1998; Schmidlin et al., 2006; Vezys et al., 2006). Likewise, induction of IFN-I by TLR ligand polyinosine-polycytidilic acid (poly-IC) – possibly through thymus resident plasmacytoid dendritic cells (pDCs) known to produce high levels of IFN-α/β in T cell progenitor environment (Gurney et al., 2004) – has been shown to distinctively impact developing T cells when compared to mature peripheral T lymphocytes, which receive pro-survival signals. Combining these observations with impaired thymic function in chronic HIV infection (Dion et al., 2004; Douek et al., 2001; Hatzakis et al., 2000; Richardson et al., 2000), we hypothesized that IFN-I signaling during persistent viral infections inhibits the development and maintenance of a persistent immune response by inhibition of thymopoiesis. Using a head-to-head comparison of IFN-I responses during acute and chronic LCMV infections, our studies show that acutely infected mice display partial and transitional atrophy of the thymus, whereas chronically infected mice undergo severe and prolonged atrophy. Our studies further delineate a T cell developmental arrest in chronically infected mice linked to a divergent qualitative IFN-α response and a higher sensitivity to IFNs compared to acutely infected mice. A specific pattern of IFN-α subtypes was found to induce thymic defects through direct as well as CD8 T cell-dependent indirect mechanisms. These findings provide a mechanistic framework for targeting IFN-I signals during immunotherapy of chronic viral infections.

MATERIALS AND METHODS

Mice and infections

Four- to six-week-old female C57Bl/6 mice were purchased from Jackson Laboratory (Bar Harbor, Maine). Four- to six-week-old female Rag-2p-GFP mice were a generous gift from Michel Nussenzweig (The Rockefeller University, New York, USA). Four- to six-week-old female IFN-α/βR^−/− mice resulted from a 20-fold backcrossing of IFNaR^−/− mice (129Sv/Ev genetic background) on a C57BL/6 genetic background. CD8^−/− mice were procured from the Jackson Laboratory. Mice received 2 × 10⁶ PFU of LCMV Armstrong strain (LCMV_Arm) intravenously (i.v.) to initiate an acute infection, or 2 × 10⁶ PFU of LCMV Clone-13 strain (LCMV_Cl-13) intravenously to initiate chronic infection (Zajac et al., 1998).

Antibody depletion

CD8 T cells were depleted in 4–8 weeks old C57Bl/6 mice by intraperitoneal injection of 50μg/mouse anti-CD8 antibody (clone 53–6.7) 12–16 hours prior to LCMV infection. Depletion of peripheral CD8 T cells was confirmed by flow cytometry analysis of PBMC samples.

Isolation of organs and cell suspension

After sacrifice, thymuses and mesenteric lymph nodes (mLN) were isolated, mechanically disrupted, treated 10 minutes with DNase I (Sigma) and collagenase-D (Roche) (except for spleens), ACK solution, and prepared as single-cell suspensions.

Flow cytometry and tetramer staining

Single-cell suspensions were incubated with the appropriate Abs for 30 minutes on ice, followed by washing. Live events were collected based on forward- and side-scatter profiles on a FACSCalibur or LSR-II flow cytometer (Becton Dickinson, (BD)). Abs were purchased from BD and included CD3-APC, CD3-FITC, CD4-PE, CD4-PerCP, CD8-PerCP, CD8-FITC, CD44-FITC, CD25-APC, CD62L-APC, CD69-PE, CD24-PE, BrdU-Alexa 532, Annexin-V-FITC, MHC-I-PE and MHC-II-FITC. D^bGP276-APC, D^bGP33–41-APC, D^bNP396-APC tetramer and cell surface staining was performed as described previously (Kalia et al., 2010a; Kalia et al., 2010b; Sarkar et al., 2008). Thymic epithelial cell (TEC) enrichment was performed with collagenase D (Roche) and 0.1% (w/v) DNAse I (Sigma), as previously described (Gray et al., 2002).

For staining with anti-BrdU, animals were injected i.p. two times with 1 mg of BrdU (Sigma-Aldrich, St. Louis, MO) and the thymus was taken 1 hour after the last injection. Then, cells were stained for cell surface markers, fixed and permeabilized with PFA/Tween and stained using the FITC conjugated BrdU kit (BD Pharmingen) according to the manufacturer’s instructions. To specifically evaluate early thymocyte progenitors, we studied CD44 and CD25 expression in PerCP lineage-negative (lin^–) cells from mice (BD Pharmingen). Lin^– cells were defined as negative for lineage marker surface expression: CD11b (Mac-1), Ly-6G (Gr.1), CD45R/B220 and TER-119/Erythroïd cells (Ly-76) and CD3. MHC class I tetramers of H-2D^b complexed with LCMV NP_396–404, GP_33–41 or GP_276–286 were produced as previously described (Murali-Krishna et al., 1998). Biotinylated complexes were tetramerized using allophycocyanin-conjugated streptavidin (Molecular Probes, Eugene, Ore.).

Serologic Analysis

Serum levels of IFN-α and IFN-β were determined using an ELISA kit (PBL Biomedical Laboratories) (Demoulins et al., 2008).

IFN-α Heteroduplex-Tracking Assay (IFN-HTA) and type I IFN Real Time PCR analysis

IFN-HTA assay and real time PCR primers have been described elsewhere (Demoulins et al., 2009). Briefly, IFN-α PCR was carried out for 35–45 cycles with the use of consensual primers: sense 5′-ATGGCTAGRCTCTGTGCTTTCCT-3′, antisense 5′-AGGGCTCTCCAGAYTTCTGCTCTG-3′. PCR amplification conditions were established to avoid PCR recombination among IFN-α subtypes (35–45 × 94 °C 30 s, 60 °C 30s and 72 °C 10 mn). IFN-HTA probes (i.e., IFN-α2(s), -α8/6(as), α11(as) and -α14(as)) were generated by PCR in 100 μl reaction volume from 100 ng of purified subcloned IFN-α subtype using one of the two consensual primers coupled with fluorescein (Invitrogen, Montreal, Ca). The mixture of the HTA probe and target was denatured for 3 min at 95 °C using a thermocycler and directly cooled on ice. DNA heteroduplexes were separated on 16 cm high, 1.5 mm thick, 4% stacking and 8% polyacrylamide gels in 1× TBE at 150 V for 30 min (pre-run) and 200 V for 6 h. IFN-HTA gels were scanned for fluorescein with a Typhoon trio (Amersham Biosciences) and the results were visualized and quantified with ImageQuant analysis 5.1 software (Molecular Dynamics). ImageQuant allows for a densitometric quantitation of each heteroduplex band’s signals. As such, a subtype’s frequency can be calculated as the percentage of this particular transcript among all members of the IFN-α multigene family.

Statistical analysis

Data are expressed as means ± s.e.m. Statistical significance of differences was determined by the paired two-tailed Student t-test. Differences were considered statistically significant for p<0.05. Statistical analyses were performed using Excel software (Microsoft). P values of statistical significance are depicted by asterisk per the Michelin guide scale: * (P ≤ 0.05), ** (P ≤ 0.01), *** (P ≤ 0.001). (P > 0.05) was considered not significant (ns).

RESULTS

Chronic viral infection leads to severe and prolonged thymic atrophy and impaired peripheral T cell output

Previous studies from our Lab and others’ have shown an association of chronic HIV infection with impaired thymic function (Dion et al., 2004; Douek et al., 2001; Hatzakis et al., 2000; Richardson et al., 2000). Using a murine model of infection with LCMV strains that cause acute (LCMV_Arm) or persistent (LCMV_Cl-13) infections (Kalia et al., 2010a), we first investigated whether acute and chronic viral infections exert a differential impact on thymopoiesis. T cell development was monitored during early stages of the immune response following infection (from days 2 to 15 post-infection). Profound differences were observed between acute and chronic LCMV infections. Acute LCMV infection was associated with a 3-fold decrease in cellularity at day 8 (partial atrophy) which completely recovered at day 15 (day 8: 96.3 X 10⁶ ± 10.2 X 10⁶, day 15: 245.0 X 10⁶ ± 10.5 X 10⁶, p<0.001). Strikingly, chronic LCMV_Cl-13 infection was associated with a 26-fold decrease in cellularity at day 8 (severe atrophy), which was maintained at least up to day 15 (day 8: 13.6 X 10⁶ ± 3.8 X 10⁶, day 15: 17.0 X 10⁶ ± 2.0 X 10⁶, p=0.999) (Fig. 1A, upper panel). These differences were confirmed by visual inspection of the thymus (Un: 0.34 inches, Arm: 0.25 inches, LCMV_Cl-13: 0.19 inches) (Fig. 1B). The decrease in cellularity following chronic LCMV infection was specific to the thymus, and only a modest decrease was observed in the spleen for both infections (Fig. 1A, lower panel).

FIGURE 1. Chronic viral infection leads to severe and prolonged thymic atrophy that rapidly affects peripheral output.

(A) Loss of thymic cellularity. Absolute cell counts of thymus and spleen were obtained by cell numeration. (B) Shrinkage in overall thymus size. (C) Rapid reduction of peripheral naive T cell pool as a result of thymic atrophy during LCMV_Cl-13 infection. Splenocytes were stained for cell surface markers CD4, CD8, CD44 and CD62L and naive (CD44^-CD62L⁺) and memory (CD44⁺CD62L^-) phenotypes were defined. (D) Thymocytes numbers and numbers of Rag-2p-GFP⁺ cells in spleen of LCMV_Cl-13 infected mice are presented. These mice are transgenic for GFP driven by the RAG-2 promoter, allowing the detection of RTE in the periphery (Boursalian et al., 2004). (E) Peripheral CD4⁺ and CD8⁺ T cell counts were plotted as a function of thymic cellularity. (C-E) Data obtained at Day 15 post-infection. Data are representative of two independent experiments.

The strong thymic atrophy observed in chronic LCMV infection led to a substantial decrease in the absolute numbers of naive T cells in the spleen when compared to uninfected mice (CD4⁺: p=0.003; CD8⁺: p=0.004) or acutely infected mice (CD4⁺: p=0.051; CD8⁺: p=0.020). In contrast, the memory T cell pool remained largely unaltered in all groups (Fig. 1C). These data suggest that the severe thymic atrophy during chronic infections may exert a negative impact on the mature peripheral T cell output. Consistent with this, a strong correlation between thymic cellularity (Fig. 1D) and naïve CD4⁺ and CD8⁺ T cell subsets was evident, alongside a significant reduction of recent thymic emigrants (RTE) numbers at 15 days post-infection, as measured by frequencies and absolute numbers of Rag-2 GFP positive cells [Un.: 1.24 X 10⁶ ± 0.74 X 10⁶, Arm.: 0.90 X 10⁶ ± 0.64 X 10⁶, LCMV_Cl-13: 0.38 X 10⁶ ± 0.18 X 10⁶; p(Un./ LCMV_Arm)=ns, p(Un/ LCMV_Cl-13)=0.022] (Fig. 1E). These results establish a link between disease outcome and the extent of thymopoiesis alteration: acute infection was associated with partial and transient defects, whereas chronic infection exhibited severe and prolonged defects.

Multi-step T cell developmental arrest is associated with thymic atrophy during viral infection

Thymic atrophy induced by TLR ligands has been shown to significantly impair DN2/DN3 as well as DP thymocyte maturation (Baron et al., 2008; Demoulins et al., 2008). Hence, we next evaluated T cell development in the context of virus infection-induced thymic atrophy. Kinetics of the decrease of DN3 and DP numbers in acute LCMV infection were identical to those observed for total thymocyte population: partial decrease observed at day 8 post-infection that progressively returned to basal levels at day 15. In contrast, chronic LCMV infection was associated with significant decrease of DN3 and DP numbers at day 8 post-infection, which persisted up to at least 15 days; (DP at day 8: p(Un/ LCMV_Arm)=0.004, p(Un/ LCMV_Cl-13)=0.001, p(LCMV_Arm/ LCMV_Cl-13)<0.001) (DP at day 15: p(Un/ LCMV_Arm)=0.0945, p(Un/LCMV_Cl-13)<0.0001, p(LCMV_Arm/LCMV_Cl-13)=0.0002) (Fig. 2A). At day 8, the distribution of the DN1–4 subsets was only slightly altered in acute infection, whereas the DN2 and DN3 phenotype cells completely disappeared in chronic infection (p=0.001 and p<0.001, respectively) (Fig. 2B, upper panel). Additionally, DP T cell frequency was more profoundly affected in chronic infection, [Day 8: p(Un/LCMV_Arm)=0.0125, p(Un/LCMV_Cl-13)<0.0001 and p(LCMV_Arm/LCMV_Cl-13)=0.0001] (Fig. 2B, lower panel). Finally, SP CD4⁺ and SP CD8⁺ thymocytes displayed a continuous decline in chronic infection, whereas these subsets were restored by day 15 in acute infection after a modest transient decline at day 8 (Fig. 2A). Supp. Fig. 1 displays the frequencies of the various thymocyte subsets over the course of LCMV infection. The data demonstrate that DN thymocyte blockade occurs as early as Day 2 post-infection (frequency of DN2 thymocytes: Un.=5.8 %, LCMV_Cl-13=3.4%), thus justifying the use of early time points (i.e., Days 1–2 post-infection) for subsequent investigations of DN subsets. On the other hand, the DP thymocyte depletion was clearly obvious at day 8 post-infection (frequency of DP thymocytes: Un.=85 %, LCMV_Cl-13=15.3%), thus identifying later time points (i.e., Days 6–8 post-infection) as optimal for evaluating DP thymocytes.

FIGURE 2. Characterization of the thymic atrophy during chronic viral infection.

(A) Absolute cell count of thymocyte progenitor’s subsets: DN3 (CD4^-CD8^-CD44^-CD25⁺), DP (CD4⁺CD8⁺), SP CD4⁺ and SP CD8⁺ were determined by flow cytometry analysis and absolute counts were obtained by cell numeration at indicated time-points. (B) Upper panel. Blockade in DN1-DN2 transition. Representative dot plots are shown from gated CD4^-CD8^- population. DN1 (CD44⁺CD25^-), DN2 (CD44⁺CD25⁺), DN3 (CD44^-CD25⁺) and DN4 (CD44^-CD25^-) were determined by flow cytometry analysis. Lower panel. Severe and prolonged decrease of the frequency of DP thymocytes. Shown are representative dot plots with the percentages of thymic subsets: DN, DP, SP CD4⁺ and SP CD8⁺. Data were obtained at Day 8 post-infection and represent the mean of 3 mice. (C) Increased apoptosis of thymocyte in chronic infection. Cells were stained for cell surface markers CD3, CD4, CD8 and Annexin-V. Apoptosis levels were obtained on each subset by flow cytometry. Representative FACS plots and bar graphs depicting means of 3 mice per group are presented from Day 8 post-infection. (D) Slow down of early thymocyte progenitor proliferation. Mice were injected i.p. twice with 1 mg of BrdU and the thymus was taken 1 hour after the last injection. Cells were stained with cell surface markers CD3, CD4, CD8, CD25, CD44 and Lin-SAV. Level of proliferation was determined by BrdU uptake using flow cytometry. Representative FACS plots from 3 mice per group are presented along with bar graphs showing average values from all mice. Data were obtained at Day 2 post-infection. (E) Increased retention of SP CD8⁺ T cells within the thymus of LCMV infected mice. Shown are representative dot plots of CD69 in DN3, DP, SP CD4⁺ and SP CD8⁺ thymocyte subsets from two independent experiments. Bar graphs depict mean values from all mice. Data were obtained at Day 2 post-infection. (F) Prolonged loss of thymic stromal cellularity following chronic virus infection. Absolute cell counts are presented at indicated time-points. (G) MHC-I and -II expression levels on TECs. Thymii were collected and prepared as single-cell suspension and stained for cell surface markers MHC-I, -II and CD45. TECs were defined as MHC-II⁺CD45^-. Level of marker expression from infected mice relative to naïve mice are presented as bar graphs with means ± SD of 3 mice per group per time point. Stars indicate significance levels. *, p < 0.05; **, p < 0.01.

The majority of DP cells undergo apoptosis during normal thymopoiesis, whereas the DN3 thymic developmental stage is characterized by high levels of proliferation. Hence, we next investigated whether the heightened loss of DP and DN thymocytes in chronic viral infection was related to increased apoptosis and decreased proliferation, respectively. Results illustrated in Fig. 2C clearly show the involvement of apoptosis in the thymic atrophy of chronic infection where mice displayed exacerbated levels of Annexin-V staining for all thymocyte subsets [ex: SP CD8⁺: p(Un/ LCMV_Arm)=0.059, p(Un/ LCMV_Cl-13)=0.002, p(LCMV_Arm/LCMV_Cl-13)= 0.132]. With respect to thymocyte proliferation, we observed a significant decrease in BrdU⁺ cells in most thymocyte subsets during both acute and chronic infections. This inhibition of proliferation was more pronounced during chronic infection for DN3, DP and SP CD8⁺ thymocyte subsets (e.g.: p(Un/ LCMV_Arm)=0.055, p(LCMV_Arm/ LCMV_Cl-13)=0.001) (Fig. 2D). Of note, the decrease in cellular proliferation was specific to the thymus and T cell proliferation in the periphery was similarly increased during both acute and chronic viral infections (Supp. Fig. 2).

We have previously shown that the thymic egress of mature thymocytes is inhibited following TLR signaling (Demoulins et al., 2008). This inhibition of egress is caused by an upregulation of CD69, which complexes with the chemokine receptor S1P₁ thus resulting in sequestration (Feng et al., 2002; Nakayama et al., 2002; Shiow et al., 2006). Hence, we further queried whether the decrease in naïve T cells was not only a consequence of decreased proliferation and increased death, but also resulted from inhibition of egress of SP cells to peripheral compartments. A moderate, increase in SP CD8⁺ CD69⁺ thymocytes was observed during both acute and chronic LCMV infections. However, the DN3, DP and SP CD4⁺ T cells were largely unchanged after acute or chronic infections, suggesting that inhibition of egress from the thymus to peripheral lymphoid compartment was not particularly important in the decrease of naïve T cells observed in chronic infection (Fig. 2E). Strikingly, this observation was specific to the thymus as most T cells (≥ 96.8% in all T cell compartment) in the spleen upregulated CD69 (Supp. Fig 3). Altogether, these data demonstrate evident perturbation of early and late stages of T cell development during acute and chronic viral infections. Moreover, these data suggest that prolonged decrease in peripheral naïve T cells observed in chronic viral infection may be related to more profound and prolonged defects in T cell survival and proliferation and not reduced thymic egress.

Negative selection eliminates by apoptosis DP thymocytes whose αβTCRs bind with high affinity/avidity to the MHC/self-peptide complex. Based on increased apoptosis in DP thymocytes, we next sought to determine whether an aberrant negative selection was induced during viral infection. Profound differences were observed in thymic stroma cellularity (Fig. 2F): acute LCMV infection was associated with a decrease that entirely reversed by day 8 post-infection, whereas chronic LCMV infection was associated with a prolonged decrease in cellularity (Fig. 2F). In addition to evident differences in numbers of thymic stromal cells during acute and chronic LCMV infection, we also observed a transient increase in MHC-I expression on the thymic stromal compartment following LCMV_Cl-13 infection (Fig. 2G, top panel). The MHC-I up-regulation was also detected on CD45^- MHC-II⁺ TECs, thus providing evidence that CD8⁺ thymocyte selection milieu is likely modified. In contrast, chronic LCMV infection led to an overall decrease of MHC-II expression levels (Fig. 2G, bottom panel). These results of quantitative and qualitative aberrancies in thymic stroma are suggestive of altered thymic selection during chronic LCMV infection possibly causing a restriction of the peripheral T cell repertoire in exiting thymocytes, as we have previously reported (Demoulins et al., 2008).

Role of peripheral antigen-specific CD8⁺ T-cells in mediating thymocyte loss during viral infections.

Decreased thymic output could be attributed to direct cytotoxicity of thymocytes by functional mature T cells that traffic to the thymii in infected mice. To test this hypothesis, we assessed mature peripheral T cells in the thymii of naïve and virus-infected mice. We used CD24 expression to distinguish developing thymocytes from peripheral T cells, which characteristically lose CD24 expression following maturation (Demoulins et al., 2008; Marodon and Rocha, 1994). We observed increased frequencies of SP CD4⁺ CD24^- and SP CD8⁺ CD24^- mature T cells in atrophied thymii of both acutely and chronically infected mice, and this was exacerbated upon chronic infection (ex for SP CD8⁺: Un: 24%, LCMV_Arm: 43%, LCMV_Cl-13: 77%; p(Un/ LCMV_Arm)=0.015, p(Un/ LCMV_Cl-13)<0.001, p(LCMV_Arm/ LCMV_Cl-13)=0.024) (Fig. 3A). We then assessed if SP cells infiltrating the thymus were specific for LCMV epitopes. We tested multiple dominant [H-2D^b-restricted NP396–404 (D^bNP396), GP33–41 (D^bGP33) and GP276–286 (D^bGP276)] epitope-specific CD8 T cells in thymii of acutely or chronically infected mice (percentages in Fig. 3B, 3C; absolute counts in Supp. Fig. 4A). Increased LCMV-specific SP CD8⁺ T cells were found similarly in both acute and chronic LCMV infection (ex for D^bGP33: Un: 05 %, LCMV_Arm: 1.4 %, LCMV_Cl-13: 1.7 %; p(Un/ LCMV_Arm)=0.010, p(Un/ LCMV_Cl-13)=0.033, p(LCMV_Arm/LCMV_Cl-13)= 0.551). These data demonstrating similarly increased proportions of LCMV specific peripheral CD8 T cells in acutely and chronically infected mice suggested that differential thymic atrophy noted in acute and chronic viral infections is likely independent of deleterious activity by cytolytic antigen-specific cells within the thymus. Nonetheless, we directly investigated the role of CD8 T cells in mediating cytolysis of thymocytes by engaging two distinct strategies: germline ablation of CD8 T cells using CD8^−/− mice, and antibody-mediated (clone 53–6.7) depletion of CD8 T cells prior to acute or chronic LCMV infection. The second strategy was employed to bypass any thymic developmental aberrancies associated with germline loss of the CD8 coreceptor. Evaluation of thymic cellularity in the presence or absence of CD8 T cells revealed an evident rescue of thymocyte numbers in the absence of CD8 T cells in both acute and chronic infections (Fig. 3C-D). Notably, the DP+ thymocyte subset, which is impacted in a profound and prolonged manner during chronic LCMV infection, was significantly rescued in the absence of CD8 T cells (Fig 3F). Heightened thymic loss in chronic infection compared to acute infection was associated with higher levels of granzyme B+ effector CD8 T cells in the thymus. Notably, restoration of thymic cellularity in the absence of CD8 T cells was associated with a reduction in granzyme B+ effector CD8 T cells in the thymus in both acute and chronic LCMV infection models (Supp. Fig. 4B, 4C ). Together, these data showing partial recovery of thymic cellularity by loss of CD8 T cells in LCMV infection demonstrate that CD8 T cells partially contribute to infection-induced thymic atrophy, and further support a role for granzyme B+ effector CD8 T cells in the process.

FIGURE 3. Role of peripheral antigen-specific CD8+ T-cells in thymic atrophy.

(A) Both acute and chronic infection displayed an increase of the numbers of CD3⁺ peripheral T cells in the thymus at day 6 post-infection. CD24^- cells within the thymus have been reported to be mature and to recirculate from the periphery (Marodon and Rocha, 1994). (Left) Numbers are the percentage of CD3⁺CD24^- thymocytes measured by flow cytometry. (Right) as in left, but with histograms integrating all mice. (B) CD8⁺ tetramer positive cells in thymus during acute and chronic viral infection are similar in numbers at day 8 post-infection. Responses to three LCMV T-cell epitopes were determined using MHC tetramer staining. The percentage of SP CD8⁺ T cells staining positive for each of the three MHC class I tetramers (D^bNP396, D^bGP33 or D^bGP276 epitope) is shown for representative LCMV_Arm and LCMV_Cl-13 infected mice, and for an uninfected mice. (C) Composite data for proportions of CD8⁺ tetramer+ cells (D^bNP396, D^bGP33 or D^bGP276 epitopes) in the thymus and spleen of control and LCMV-infected mice at day 8 after infection are presented as means +/− SEM. (D) Wild-type and CD8^−/− C57Bl/6 mice were either left uninfected or infected with LCMV_Arm or LCMV_Cl-13 strains. At day 8 post-infection thymic cellularities were assessed. (E, F) C57Bl/6 mice were intraperitoneally administered PBS or anti-CD8 depleting antibody. PBS control and CD8-depleted mice were either left uninfected, or infected with LCMV_Arm or LCMV_Cl-13 and thymocytes were assessed. Raw data and summary bar graphs for proportions of double positive (DP) thymocytes are presented at day 8 after infection. Data are the average for 3–4 mice per group and are representative of two independent experiments.

Specific action of type I IFNs in the loss of thymic output during chronic viral infection

IFN-I signals are critical for driving robust clonal expansion of virus-specific CD8 T cells (Kolumam et al., 2005). Moreover, we have previously shown that induction of IFN-I by TLR ligand polyinosine-polycytidilic acid (poly-IC) distinctively impacts developing T cells compared to mature peripheral T lymphocytes. Hence, we next investigated the role of IFN-I in driving the differential levels of thymic atrophy in acute and chronic chronic LCMV infections. We hypothesized that the strong and prolonged thymic atrophy observed in chronic LCMV infection could be attributed to a divergent IFN response (quantitatively and qualitatively) compared to acutely infected mice. IFN-α and IFN-β were quantified in the sera of LCMV_Arm or LCMV_Cl-13 infected mice by ELISA at various time points over a 144 hour period (Fig. 4A). Plasma levels of IFN-α and IFN-β were significantly elevated in infected mice compared to uninfected controls. Levels of type I IFNs reached a peak at 18–24 hours and remained easily detectable up to 60 hours post infection. However, induction of IFN-I was faster in acute infection such that higher levels of both IFN-α and IFN-β were observed in Arm infected mice at 12 hours post-infection. Production of IFN-β appeared slightly higher in acute infection at early time-points, and returned to low levels earlier than IFN-α (36 hours post-infection versus 72 hours) (Fig. 4A). Overall, these data demonstrate that LCMV infection induces a tightly regulated IFN-α/β production: strong, early and transient.

FIGURE 4. Specific action of type 1 IFNs in the loss of thymic output during chronic viral infection.

(A) Serum IFN-α and -β protein production quantification by ELISA. Data presented are the means 3 mice that were bled at various time points on a 144 hours length period. (B) IFN-α mRNA subtype quantification by IFN-HTA within the spleen (12h, 24h post infection). This figure plots real-time PCR IFN-α absolute level by the IFN-HTA calculated frequencies to obtain a quantitative measurement of IFN-α subtypes: (S (Mean RT-PCR IFN-α X HTA Mice A) + (Mean RT-PCR IFN-α X HTA Mice B)…)/3 (Demoulins et al., 2009). Results are the mean and standard deviation of 3 individual mice per group per time point (C-E) Specific role of type I IFNs on thymic cellularity. (C-E) WT and IFN-α/βR^−/− mice were uninfected, infected with LCMV_Arm or LCMV_Cl-13 and were sacrificed 6 days post infection. (C) Data represent absolute thymic cellularity of WT versus IFN-α/βR^−/− mice. (D) DN3 and DP cellularity of WT versus IFN-α/βR^−/− mice day 6 post-LCMV infection. (E) IFN-α/βR^−/− mice chronically infected no more exhibited an increased apoptosis of thymocyte, as observed in WT mice. Cells were stained for cell surface markers CD3, CD4, CD8 and Annexin-V. Apoptosis levels were obtained on each subset by flow cytometry. Data are presented as means and standard deviation of 4 mice per group, and are representative of two independent experiments.

To confirm the role of IFN-I in induction of thymic atrophy, we employed IFN-α/βR^−/− mice that lack the IFN-α/β receptor. Arm or Cl-13 infected WT and IFN-α/βR^−//− mice were sacrificed at day 6 post-infection to dissociate the effect of type I IFNs from viral clearance. Remarkably, the severe (LCMV_Cl-13) and partial (LCMV_Arm) thymic atrophies were completely abogated in IFN-α/βR^−//− mice [p(Un/ LCMV_Arm)= 0.2180, p(Un/LCMV_Cl-13)=0.3736, p(LCMV_Arm/LCMV_Cl-13)= 0.5011] (Fig. 4B). These differences between IFN-α/βR^−//− and WT mice could not be attributed to differences in the average thymus cellularity for both strains (WT Un: 202.7 X 10⁶ ± 115.2 X 10⁶, IFN-α/βR^−//− Un: 173.4 X 10⁶ ± 58.4 X 10⁶,p=0.4610) and clearly demonstrate that IFN-α/β signaling pathway is a major determinant in the induction of thymic atrophy in LCMV infection. Contrary to WT mice, numbers of DN3 and DP subsets in IFN-α/βR^–/– infected mice remained similar to uninfected controls (ex. For DN3: p(Un/ LCMV_Cl-13 WT)=0.012; p(Un/ LCMV_Cl-13 IFN/βR^−/−)=0.592) (Fig. 4C), demonstrating that the contribution of type I IFNs on thymus reduction in size occurs as early as the DN stages. Moreover, analysis of apoptotic cells in thymii of IFN-α/βR^−/− infected mice (Fig. 4D) established the direct impact of type I IFNs on apoptosis – unlike WT mice, which showed increased Annexin-V⁺ cells in the thymus compared to uninfected controls, thymii of IFN-α/βR^−//− mice exhibited similar levels of apoptosis as uninfected mice [WT DP: p(Un/ LCMV_Cl-13)=0.0048, WT CD4⁺: p(Un/ LCMV_Cl-13)=0.0053, WT CD8⁺ : p(Un/LCMV_Cl-13)=0.0550) (IFN-α/βR^−/− DP: p(Un/LCMV_Cl-13)=0.1733, IFN-α/βR^−/− CD4⁺: p(Un/LCMV_Cl-13)=0.1281, IFN-α/βR^−/− CD8⁺ : p(Un/LCMV_Cl-13)=0.1414]. Finally, unlike WT mice, infiltration of SP CD4⁺ CD24^- and SP CD8⁺ CD24^- was completely abogated in the thymus of LCMV-infected IFN-α/βR^−/− mice (Supp. Fig. 5). Reduced thymic infiltration by mature peripheral CD8 T cells occurred despite increased viral loads and largely similar numbers of virus-specific CD8 T cells in the spleens of IFN-α/βR^−/− mice (Supp. Fig 5). Combined with our observations of CD8 T cell-dependent loss of thymocytes (Fig 4), these findings support the notion that IFN-I signals mediate thymic loss in chronic infection in part through direct effects on virus-specific CD8 T cell thymic infiltration. This is consistent with recent study showing mitigated thymic loss in chronic infection when virus-specific CD8 T cells specifically lacked IFNR1 (Elsaesser et al., 2020).

IFN-β exerts a dominant role in regulating effector CD8 T cell responses and viral control (Ng et al., 2015), and exerts minimal effects on thymic loss during chronic LCMV infection (Elsaesser et al., 2020). We next characterized the IFN-α response diversity in terms of individual subtypes between acute and chronic infection. To assess this, we used an IFN heteroduplex-tracking assay (IFN-HTA) (Demoulins et al., 2009) (Fig. 4E). At 12 hours post-infection, LCMV_Cl-13 and LCMV_Arm infected mice showed distinct distributions of the different IFN-α subtypes: IFN-α1 (p=0.04447), IFN-α11 (p=0.0113), IFN-αB (p=0.0338) were lower, whereas IFN-α6T (p=0.0254), IFN-α7/10 (p=0.0010), IFN-αψ2 (p=0.0047) were higher in LCMV_Cl-13 infected mice compared to LCMV_Arm infected mice. Together with serum IFN-I levels, these data indicate an altered type I IFN program in acute versus chronic viral infections. These data are consistent with previous reports showing altered expression patterns of IFN-I between acute and chronic LCMV infections (Clingan et al., 2012; van Pesch et al., 2004; Wang et al., 2012). Altogether, these results demonstrating an evident upregulation of IFN-I during early stages of infection, with a distinct expression pattern of IFN-α in acute and chronic infections, and mitigation of chronic virus infection-induced thymic loss in the absence of IFNα/βR demonstrate a role for IFN-I in promoting thymic atrophy, possibly through induction of apoptosis and/or inhibition of proliferation.

DISCUSSION

This study highlights a critical role of IFN-I signals in establishing severe and prolonged thymic atrophy in the context of chronic infections. Further, providing finer resolution of the IFN-I response in acute and chronic infections, this study establishes an association of altered kinetics and pattern of IFN-α subtype expression with the survival and proliferation of specific subsets of developing T cell lineages in the thymus. The unique pattern of IFN-I expression in acute and chronic infections suggests that both an altered pattern of type-I IFN subsets as well as altered sensitivity of thymocytes to IFN-α in the context of chronic viral infections might be responsible for increased apoptosis, decreased proliferation and loss of thymic cellularity. These quantitative and qualitative differences in the type I IFN response between acute and chronic LCMV infections can provide new insights for the understanding of human chronic infection, given that similar observations of aberrant thymopoiesis related to a modified IFN-α subtype production have also been noted in SIV-infected rhesus macaques (Dutrieux et al., 2014).

Our observations of modestly enhanced IFN-I at 12 hours of acute (compared to chronic) LCMV infection and largely similar levels at 48 hours post-infection are consistent with previous reports (Sullivan et al., 2015; Teijaro et al., 2013). It is possible that more rapid induction of specific IFN-I subsets with more potent antiviral functions during acute infection contributes to more rapid viral clearance, thus creating an environment that is more conducive for thymopoiesis. Consistent with this proposal, previous report showing enhanced chronic viral control following initial priming with acute strain of LCMV, further provide a functional context for the distinct IFN-I milieu induced during early stages of acute and chronic LCMV infections (Sullivan et al., 2015). Consistent with largely immunosuppressive role of IFN-β and anti-viral role of IFN-α in chronic LCMV infection (Ng et al., 2015), our IFN-α subtyping studies show significantly reduced expression of subtypes with higher antiviral activities (IFN-α11 and IFN-ß), and significantly increased expression of IFN-α7/10 with lower antiviral activity due to the lack of the Cys1 residue (van Pesch et al., 2004). Specific role of IFN-α in mediating thymic loss is supported by collective findings from our group and others showing differential expression of unique antiviral IFN-α subsets, and reduced chronic infection associated thymic atrophy in the absence of IFNα/βR, but not IFN-β, which is largely associated with immunosuppressive effects (Elsaesser et al., 2020; Ng et al., 2015).

Our observations of partial restoration of thymic cellularity in the absence of CD8 T cells and full restoration of thymic cellularity in the absence of IFN-α/βR support the notion that IFN-I signals induce thymic atrophy through both CD8 T cell-dependent and independent mechanisms, which is in line with a recent study (Elsaesser et al., 2020) showing mitigated thymic loss when IFNR1 was specifically ablated on virus-specific CD8 T cells. In this study, IFN-g+ LCMV-specific (but not LCMV non-specific OT-1) TCR-transgenic CD8⁺ T cells were shown to rapidly migrate into the thymus to induce thymocyte loss. Consistent with these findings, our data show that mature granzyme B+ peripheral CD8 T cells traffic to the thymii, albeit to different extents in acute and chronical infections. Together, these findings confirm that activated virus-specific (and not bystander T cells) CD8 T cells contribute to thymic ablation in an IFN-I-dependent manner. It is plausible that thymocyte loss during early stages of LCMV infection might be dominantly mediated by CD8 T cell-dependent IFN-I signals, whereas CD8 T cell-independent effects of IFN-I might be primarily responsible for the severe and prolonged thymic atrophy that occurs during chronic infections, in the face of CD8 T cell exhaustion. Mechanistically, it is possible that IFN-I signals exert direct effects on T cell progenitors, which might respond with increased sensitivity or mount an aberrant response to the altered expression pattern of type-I interferons observed during later stages of chronic infection. Additionally, IFN-I could also cause thymic atrophy via indirectly inhibiting the migration or numbers of T cell progenitors from bone marrow (Binder et al., 1997; Binder et al., 1998; Isringhausen et al., 2021; Pascutti et al., 2016). These possibilities are the target for future investigations.

Our observations of largely similar numbers of splenic antigen-specific CD8 T cells in IFN-I sufficient and deficient (through germline deletion or antibody blockade) settings of chronic LCMV infection are consistent with previous reports (Ng et al., 2015; Teijaro et al., 2013; Wilson et al., 2013) Nonetheless, possibly owing to model system-specific differences in the highly dynamic nature of IFN-I expression, and the range of mechanisms by which IFN-I signals may modulate CD8 T cell responses (such as through direct effects on CD8 T cells, or indirectly through CD4 T cells, Treg cells, DCs, macrophages, NK cells and systemic cytokine alterations) increased or even decreased splenic numbers of antigen-specific CD8 T cells have been reported in the absence of IFN-1 signaling (Gangaplara et al., 2018; Mbanwi et al., 2017). Findings of reduced thymic infiltration in the absence of IFN-1 despite apparent changes in splenic numbers are suggestive of thymus-specific defects in survival or trafficking of virus-specific CD8 T cells.

Thymic atrophy induced during chronic infection is paradoxical with the necessity of a diverse and robust peripheral T cell pool, and raises into question the relative importance of available peripheral T cell repertoire versus RTEs in early phases of infection. Using Rag2-GFP transgenic mice it has been shown that 20–25% of peripheral T cells are comprised of GFP⁺ RTEs, and about one week is the estimated time for entry into the naive T cell pool (Boursalian et al., 2004). Therefore, it is conceivable that in the first days of infection, delay of RTEs to enter the naive pool will not impact the early phase of viral clearance. Consistent with the assumption that thymic output is not required in early phases of immune response, a study showed that thymectomy had minimal impact on the expansion phase of the CD8 T-cell response in acute LCMV infection. However, this study was restricted to the observation of only 3 viral epitopes (Miller et al., 2005). Whether the lack of generation of RTE is compensated by an increased half-life and proliferation rate remains to be assessed. Also, despite strong thymic atrophy mediated by IFN-I, several reports have shown thymic capacity to regenerate and produce new T cells after about a month of infection (Demoulins et al., 2008; Elsaesser et al., 2020), thus suggesting that thymic functionality is likely restored in the context of a subsequent new infection. However, it is important to consider that multiple infections may accrue alterations in the naive T cell repertoire over time. Our observations of altered thymic stromal and epithelial cells and related selection process, support this possibility of alterations in the T cell repertoire due to aberrant negative selection, thus compromising the immune ability to effectively tackle future new infections. Moreover, although numerous infections are successfully cleared by the host’s immune response, certain viral infections are only slowly or never resolved, as shown in a previous study that HIV infection disrupts the development of T cells early in the course of disease progression and thymic defect was maintained through time (Dion et al., 2004).

Collectively, our findings demonstrate a critical role of type-I interferon signals in promoting immunosuppression during persistent infections by inducing thymic atrophy through a unique program of IFN-I subset induction, and present this pathway as a target for regulating chronic viral infections and cancers.

RESEARCH HIGHLIGHTS.

This study highlights a critical role of type-I interferon signals in promoting immunosuppression during persistent infections by inducing thymic atrophy through a unique program of IFN-I kinetics and subset induction. This study further presents novel insight into the effects of dysregulated IFN-I on the specific subsets of developing T cell lineages in the thymus, and presents this pathway as an immunomodulatory target for potentially manipulating the outcome of chronic viral infections and cancers.