T Cell–Intrinsic CX3CR1 Marks the Most Differentiated Effector CD4⁺ T Cells, but Is Largely Dispensable for CD4⁺ T Cell Responses during Chronic Viral Infection

During chronic viral infections, such as with HIV in humans or lymphocytic choriomeningitis virus clone 13 (LCMV13) infection in mice, the long-term persistence of Ag results in the exhaustion of the virus-specific CD8⁺ T cells characterized by the progressive loss of T cell effector functions (1, 2). The depletion of CD4⁺ T cells from mice prior to infection with LCMV13 (3–5) or the loss of CD4⁺ T cells during progressive HIV infection (6) causes increased viral burden, immune dysregulation, and functional exhaustion of the CD8⁺ T cells. In contrast, adoptive transfer of virus-specific CD4⁺ T cells into mice chronically infected with LCMV13 enhances the effector function of exhausted CD8⁺ T cells and increases the control of viral replication (7). Thus, CD4⁺ T cells are critical for the control and clearance of virus during chronic viral infections.

Many factors contribute to the CD4⁺ T cell response during viral infection, including cytokines, T cell inhibitory receptors, and costimulatory receptors. The glucocorticoid-induced, TNFR-related protein (GITR), a costimulatory receptor, plays a critical T cell–intrinsic role in accumulation of T cells during acute and chronic viral infection (8–10). During chronic LCMV13 infection, CD4⁺ T cell–intrinsic GITR signaling is critical for the accumulation of IL-2⁺ Th1 cells, IL-2–dependent help for the CD8⁺ T cell response, and viral control (9, 10). Using adoptive transfer of GITR^+/+ and GITR^−/− TCR-transgenic SMARTA cells to distinguish intrinsic versus cell extrinsic effects of GITR on the CD4⁺ T cell response, we previously showed that GITR-dependent signals on CD4⁺ T cells occurred postpriming and resulted in increased surface expression of several prosurvival receptors, including CD25, CD127, and OX40 (11). RNA sequencing and flow cytometry experiments also showed that the fractalkine receptor, CX3CR1, is upregulated on Ag-specific CD4⁺ T cells, with higher levels on GITR^+/+ compared with GITR^−/− cells during LCMV13 infection. However, the role of CX3CR1 on T cells during lymphocytic choriomeningitis virus (LCMV) infection was not investigated (11).

Until now, there have been limited data on the role of CX3CR1, specifically on T cells. During allergic and atopic diseases, CX3CR1 regulates pathological outcomes by controlling effector CD4⁺ T cell survival in inflamed lungs or CD4⁺ T cell retention in inflamed skin, respectively (12, 13). In viral infection models, CX3CR1 has mainly been studied as a marker of distinct CD8⁺ T cell subsets (14–16). For example, during acute LCMV infection, CX3CR1 expression distinguishes three distinct populations of CD8⁺ T cells, with the highest expression of CX3CR1 defining the most differentiated effector population (15). Similarly, during influenza infection, CX3CR1 is most highly expressed on the most differentiated CD8⁺ T cells, whereas CD4⁺ T cells expressed minimal CX3CR1 levels in this context (16). CX3CR1 expression also defines the most highly polarized cytotoxic effector memory CD4⁺ T cells after dengue virus infection (17), but whether CX3CR1 contributes to T cell responses during viral infection remains poorly understood.

In this study, we explored the intrinsic role of CX3CR1 on CD4⁺ T cells during chronic infection with LCMV13. We show that CX3CR1 is induced on the CD4 T cells upon Ag stimulation and expressed on CD4⁺ effector T cells at the peak of the response to LCMV13. We also show that high levels of CX3CR1 mark the CD4⁺ T cells with the most highly differentiated Th1 T-bet^hi phenotype. Using mixed adoptive transfer with CX3CR1^+/+ and CX3CR1^−/− CD4⁺ SMARTA T cells, we found that CX3CR1 expression did not impact the number or tissue distribution of CD4⁺ T cells at the effector stage of the response but had a marginal, albeit significant, effect on levels of T-bet and IFN-γ per cell. At the chronic phase, CX3CR1^−/− CD4⁺ T cells were slightly enriched over CX3CR1^+/+ tissue-resident memory T cells (Trm) potentially linked to lower T-bet in the Trm precursors. Taken together, this study shows that CX3CR1 marks the most differentiated CD4 effector subset but is largely dispensable for CD4⁺ T cell responses during chronic infection with LCMV13.

LCMV13, obtained from M. Oldstone, Scripps Research Institute, was propagated on L929 cells. Age-matched female mice (6–8 wk old) were infected i.v. with 2 × 10⁶ focus forming units of LCMV13. C57BL/6 mice were obtained from Charles River Laboratories (St. Constant, QC, Canada). GITR^−/− mice were kindly provided by Dr. C. Riccardi (University of Perugia) and Dr. P. Pandolfi (Harvard Medical School) and underwent additional backcrossing. After, GITR^−/− mice were analyzed by single-nucleotide polymorphism analysis across 1500 single-nucleotide polymorphisms differing in the 129 and B6 genome and found to be a minimum of 92% C57BL/6 (The Centre for Applied Genomics, University of Toronto). The mice were then further crossed with TCR-transgenic SMARTA mice (18), which express a transgenic TCR specific for the I-A^b–restricted LCMV gp_61–80 (kindly provide by Dr. P. Ohashi, Princess Margaret Cancer Center, Toronto, ON). F2 wild-type (WT) and GITR^−/− littermate SMARTA mice were then further crossed with CD45.1 mice (Charles River Laboratories) for use in adoptive transfer studies. CX3CR1^−/− mice were obtained from The Jackson Laboratory (Bar Harbor, ME) and were crossed with CD45.1 SMARTA mice to generate CD45.1 SMARTA CX3CR1^−/− and CD45.1/2 SMARTA CX3CR1^+/+, and the F2 littermates were then bred separately for adoptive transfer experiments. For bone barrow chimera experiments, Thy1.1 (B6.PL-Thy1^a/CyJ) and CD45.1 (B6.SJL-Ptprc^a Pepc^b/BoyJ) mice were obtained from The Jackson Laboratory. All mice were housed under specific pathogen-free conditions in the Division of Comparative Medicine at the Terrence Donnelly Centre for Cellular and Biomolecular Research (University of Toronto). All animal procedures were approved by the animal care committee at the University of Toronto in accordance with the Canadian Council on Animal Care.

Anti-CD11a (clone: M17/4) was purchased from BD Biosciences (San Jose, CA). Anti-CD4 (clone: RM4-5), anti-CX3CR1 (clone: SA011F11), and anti-TCR Vα2 (clone: B20.1) were purchased from BioLegend (San Diego, CA). Anti-CD16/32 (Fc Block, clone: 93), anti-CD3ε (clone: 145-2C11), anti-CD45.1 (clone: A20), anti-CD45.2 (clone: 104), anti-CD69 (clone: [¹H].2F3), anti-GITR (clone: DTA-1), anti–IFN-γ (clone: XMG1.2), anti–T-bet (clone: eBio4B10), anti-Thy1.1 (clone: HIS51), anti-CD44 (clone: IM7), and fixable viability dye eF506 were purchased from Thermo Fisher Scientific (Waltham, MA).

Organs (lung, liver, and spleen) were harvested at the indicated times postinfection (p.i.). Lung and liver were perfused with 10 ml of PBS and then minced and digested with 2 mg ml⁻¹ of collagenase IV (Invitrogen, Carlsbad, CA) for 45 min at 37°C in a shaker. Lung, liver, and spleen tissue were mechanically disrupted through a 70-μm cell strainer to generate single-cell suspensions, which were then subjected to RBC lysis with ammonium–chloride potassium buffer. For Trm detection, mice were injected i.v. with 3 μg of anti-mouse anti-CD3ε (clone: 145-2C11) Ab, and mice were sacrificed 10 min later, and tissues were processed as described above.

Freshly isolated single-cell suspensions from the lung, liver, and spleen were treated with Fc Block for 15 min at 4°C, followed by surface staining for 30 min at 4°C. Samples were fixed with 4% paraformaldehyde following surface staining. Intracellular staining, when applicable, was performed for 30 min at 4°C following surface staining described above and permeabilization with FoxP3 Transcription Factor Staining Buffer Set (eBioscience). For intracellular cytokine staining, spleen samples were restimulated ex vivo with 4 μg/ml I-A^b–restricted LCMV gp_61–80 in the presence of 20 U/ml murine rIL-2 (eBioscience) and GolgiStop (BD Pharmingen, San Jose, CA) for 5 h at 37°C. After restimulation, cells were surface stained, fixed and permeabilized with Fixation/Permeabilization solution kit (BD Biosciences), and stained for intracellular cytokine production. Unstimulated samples (no peptide) were used as negative controls. Samples were acquired with an LSRFortessa X20 (BD Biosciences) with FACSDiva software and analyzed with FlowJo VX (Tree Star, Ashland, OR).

Naive CD4⁺ T cells were purified from spleens of GITR^+/+, GITR^−/−, CX3CR1^+/+, and CX3CR1^−/− CD45.1 SMARTA TCR-transgenic mice with EasySep Mouse Naive CD4⁺ T Cell Isolation Kit (STEMCELL Technologies, Vancouver, BC, Canada). Samples of either genotype were counted by trypan blue exclusion three times and mixed in a 1:1 ratio where indicated. SMARTA TCR-transgenic CD4⁺ T cell purity and GITR^+/+/GITR^−/− or CX3CR1^+/+/CX3CR1^−/− ratios were confirmed by flow cytometry. A total of 1 × 10⁴ cells from the mixed sample were adoptively transferred i.v. in 200 μl of PBS into recipient mice 1 d prior to infection.

Thy1.1 host mice were lethally irradiated with two doses of 550 rad and reconstituted with a 1:1 mixture of CX3CR1^−/− Thy1.2 CD45.2/CX3CR1^+/+ Thy1.2 CD45.1 bone marrow cells delivered i.v. for a total of 5 × 10⁶ cells. Reconstituted chimeric mice were given water supplemented with 2 mg ml⁻¹ neomycin sulfate (Bio-Shop, Burlington, ON, Canada) for 2 wk consecutively following irradiation and were rested for a total of 90 d before chimerism in the blood was assessed prior to infection. Chimeric mice were infected according to the indicated schedules.

All statistical analyses were performed using GraphPad Prism 6 (San Diego, CA), with the specific test performed indicated in the figure legends. The p values applied were *p < 0.05, **p < 0.01, and ***p < 0.001.

Previous work identified Cx3cr1 as a GITR-regulated gene by RNA-sequencing analysis of GITR^+/+ and GITR^−/− CD4⁺ T cells sorted from mouse spleens at day 3 after LCMV13 infection (11). Consistently, CX3CR1 protein was minimally expressed on resting T cells but expressed at a higher level on GITR^+/+ CD4⁺ T cells compared with GITR^−/− CD4⁺ T cells at day 3 p.i. (11). To determine if GITR-dependent regulation of CX3CR1 persisted through to the peak of the effector response, we adoptively transferred CD45.1 GITR^+/+ and CD45.1 GITR^−/− TCR-transgenic SMARTA CD4⁺ T cells in a 1:1 ratio into naive CD45.2 GITR^−/− mice, followed by infection with LCMV13 virus, and the expression of CX3CR1 on CD4⁺ T cells was measured at day 8 p.i. As GITR is expressed throughout the response to LCMV13 (19), we used expression of GITR to distinguish GITR^+/+ from GITR^−/− T cells (Fig. 1A). Before infection, both GITR^−/− and GITR^+/+ SMARTA T cells showed minimal levels of CX3CR1, and no difference was observed between GITR-deficient and WT cells (Fig. 1B), consistent with previous results (11). At day 8 p.i., CX3CR1 expression was higher in Ag-specific SMARTA T cells when compared with those cells before infection and compared with the host cells p.i. (Fig. 1B). Ag-specific GITR^−/− SMARTA T cells showed a lower frequency of CX3CR1⁺ cells and reduced CX3CR1 expression per cell when compared with GITR^+/+ SMARTA cells in the spleen (Fig. 1B–D). These results show that LCMV13 infection induces CX3CR1 expression on Ag-specific CD4 T cells and confirms that GITR costimulation increases the frequency of CD4⁺ T cells expressing CX3CR1 as well as the level of expression per cell.

To examine the phenotype of CX3CR1 high and low effector cells, we transferred CD45.1 SMARTA CD4⁺ T cells into WT mice, infected them with LCMV13, and analyzed CX3CR1 and T-bet expression. Cells recovered from spleen were also restimulated with LCMV gp_61–80 peptide to analyze IFN-γ production (Fig. 2A). In contrast to published reports on CD8⁺ T cells that show three populations of CD8⁺ T cells based on CX3CR1 surface expression (15), we observed more of a continuum of CX3CR1 expression on activated Ag-specific CD4 T cells, ranging from low to high. Therefore, we divided the transferred SMARTA T cells into two populations, high versus low/intermediate for CX3CR1 (Fig. 2B). Based on this gating strategy, the CX3CR1^hi SMARTA population showed about a 2-fold increase in the proportion of T-bet–positive cells and a modest increase in the mean fluorescence intensity (MFI) for T-bet. There was a small decrease in the proportion of IFN-γ⁺ CX3CR1^int/low CD4⁺ T cells compared with the CX3CR1^hi cells, and this was reflected in about a 2-fold decrease in IFN-γ MFI in this population (Fig. 2C, 2D). To analyze the cytotoxic profile of these cells, we also measured perforin and granzyme B in the two populations. We observed an increased proportion of granzyme B–positive as well as perforin-positive cells among the CX3CR1^hi SMARTA population compared with the CX3CR1^int/low CD4⁺ T cells (Fig. 2E). However, we did not see a change in the level of perforin or granzyme B per cell (data not shown). These findings show that high levels of CX3CR1 mark the most activated splenic CD4⁺ Th1 effectors, consistent with previously reported findings for CD8⁺ T cells (15, 16).

To investigate the role of CX3CR1 on the CD4⁺ T cell response, we crossed TCR-transgenic SMARTA mice with CX3CR1^−/− mice. To confirm the deletion of CX3CR1 at the protein level, we stimulated the splenocytes from SMARTA CX3CR1^+/+ or SMARTA CX3CR1^−/− mice with gp_61–80 and checked the CX3CR1 expression after 48 h. As expected, we observed a shift in CX3CR1 expression after stimulation only in the CX3CR^+/+ SMARTA cells, but not in the CX3CR1^−/− cells (Fig. 3A). Next, we adoptively transferred CD45.1/2 CX3CR1^+/+– and CD45.1 CX3CR1^−/−–transgenic SMARTA T cells in a 1:1 ratio into CD45.2 WT hosts. One day later, mice were infected with LCMV13, the organs were harvested at day 8 p.i. (Fig. 3B), and the transferred cells were analyzed by flow cytometry with gating as shown in Fig. 3C. Allowing the WT and CX3CR1-deficient T cells to compete in the same mouse eliminates the potential effects of different viral load and inflammation levels that could be observed if CX3CR1^−/− or ^+/+ SMARTA T cells were analyzed in separate hosts. The mixed adoptive transfer approach also makes it easier to observe subtle effects that might be missed because of additional biological noise if T cells were transferred into separate mice. Moreover, this mixed adoptive transfer approach allows us to analyze CD4⁺ T cell–intrinsic effects of CX3CR1 signaling.

Using this approach, we found that at day 8 p.i., the ratio of CX3CR1^+/+ to CX3CR1^−/− SMARTA T cells was close to 1 in the spleen, lung, and liver (Fig. 3D). We next examined whether the CD4⁺ T cell–intrinsic effects of CX3CR1 signaling affected the Th1 profile of CD4⁺ T cells by analyzing the frequency of T cells expressing T-bet and IFN-γ within the SMARTA population as well as the MFI of T-bet and IFN-γ per cell. We found that CX3CR1^+/+ SMARTA cells show a slightly higher frequency and level of T-bet and IFN-γ per cell (Fig. 3E, 3F) compared with CX3CR1^−/− SMARTA T cells in all the three organs analyzed (spleen, lung, and liver). These data show that CX3CR1 has little or no impact on the overall number of T cells accumulating in the spleen and tissues but that T cell–intrinsic CX3CR1 has a small, but significant, effect on the Th1 phenotype, as measured using T-bet and IFN-γ expression.

We next asked if the effects of CX3CR1 on the Th1 cells would be the same if the adoptively transferred CX3CR1^+/+- or CX3CR1^−/−-transgenic SMARTA T cells were not in competition with each other. To this end, we transferred CX3CR1^+/+- or CX3CR1^−/−-transgenic SMARTA T cells into separate WT hosts (Fig. 4A). One day after the cell transfer, the mice were infected with LCMV13, and the organs were harvested at 8 d p.i. Similar to the results in the competitive adoptive transfers, CX3CR1^+/+ SMARTA cells and CX3CR1^−/− cells were recovered at similar frequency (Fig. 4B). Also consistent with the mixed adoptive transfer experiment, a slightly higher proportion of CX3CR1^+/+ SMARTA became Th1 (CD44^hi T-bet⁺) than CX3CR1^−/− cells (Fig. 4C). Moreover, CX3CR1^+/+ T cells had a higher MFI for IFN-γ in spleen and lung, whereas this was NS in the liver (Fig. 4D). Thus, when WT or CX3CR1-deficient SMARTA cells are analyzed in separate mice, CX3CR1 does not affect the accumulation of T cells in spleen or tissues but intrinsically affects acquisition of a Th1 phenotype in some tissues with less robust results than under conditions of competition.

Trm are defined as noncirculating memory T cells that reside in the peripheral tissue and are critical for protective immunity (20, 21). Although LCMV13 is a chronic infection, cells with a Trm phenotype have been noted in chronic LCMV (22). During acute viral infections, CX3CR1^−/lo CD8⁺ T cells give rise to the Trm (15), but whether CX3CR1 impacts the accumulation of CD4⁺ Trm is unknown. To address this question in the LCMV13 chronic infection model, we used competitive mixed–bone marrow chimeras in which CX3CR1^+/+ and CX3CR1^−/− cells compete within the same mouse (Fig. 5A). Chimeric mice were rested for 90 d and blood chimerism checked before infection with LCMV13. LCMV13 is a systemic infection that can affect many different organs, and in this study, we analyzed the Trm accumulation in a representative lymphoid and nonlymphoid organ, spleen, and lung, respectively. Prior to harvest, we used intravascular infusion of fluorescently labeled Ab to distinguish lung vasculature (LV)– or splenic vasculature (SV)–exposed cells from cells located within the lung or splenic parenchymal tissue (23). CD4⁺ lung Trm were defined based on CD69 and CD11a coexpression, and CD4⁺ splenic Trm were defined based only on CD69 (Fig. 5B). At day 30 p.i., the CX3CR1^−/− CD4⁺ Trm had a small, but significant, advantage over their CX3CR1^+/+ counterparts in both the spleen and lung tissue (Fig. 5C–E). In contrast, CX3CR1^+/+ and CX3CR1^−/− Ag-specific CD4⁺ T cells in the LV and SV were found in 1:1 ratio after correcting for preinfection ratio (Fig. 5C–E). Fig. 5E shows a representative flow cytometry plot showing proportions of CX3CR1^+/+/CX3CR1^−/− of CD4⁺ T cells before infection and p.i. A similar proportion of CX3CR1^+/+/CX3CR1^−/− p.i. was also observed in the total Ag-specific CD4⁺ T cells residing in the lung and spleen, suggesting that CX3CR1 has a negative impact only on cells expressing Trm markers, including CD69 and CD11a (Fig. 5C–E), and not on the other CD4⁺ T cell populations in the lung. Moreover, we also saw a similar negative effect of CX3CR1 on the accumulation of effector memory T cells, whereas there was no impact on central memory T cells in the spleen (Fig. 5F). When we looked at IFN-γ production by CD4⁺ T cells, no difference was observed within the different compartments (Fig. 5G). These results show that the CX3CR1^−/− CD4⁺ Trm population is found in a slightly higher proportion than CX3CR1^+/+ CD4⁺ Trm in both spleen and lung.

During chronic viral infection, CD4⁺ T cells are critical for viral clearance because they play a key role in maintaining the effector function and persistence of Ag-specific CD8⁺ T cells (24). Many factors can contribute to the CD4⁺ T cell response during viral infection, including signals through GITR, a costimulatory receptor that plays an important CD4⁺ T cell–intrinsic role in T cell accumulation during viral infections (8, 9, 16). The identification of CX3CR1 as a GITR-induced molecule on CD4⁺ T cells during chronic LCMV infection (11) prompted us to analyze the intrinsic role of this chemokine receptor in this context. In this study, we confirmed that during LCMV13 infection, CX3CR1 is induced on CD4⁺ effectors. We also confirm that CX3CR1 is minimally expressed on resting CD4 T cells and increases in expression upon Ag stimulation. GITR intrinsically increases the level of CX3CR1 on the Ag-specific effector CD4 T cells after LCMV infection. Whether GITR impacts levels CX3CR1 through direct signaling or via allowing selective survival of CX3CR1^high cells cannot be ascertained from this study. We also show that high levels of CX3CR1 mark the CD4⁺ T cells with the most differentiated Th1 phenotype. However, genetic deficiency in CX3CR1 did not affect the accumulation of CD4⁺ T cells in the spleen, lung, or liver at the effector stage of the response, arguing that CX3CR1 is not required for migration of the CD4⁺ T cells to the organs during chronic LCMV infection. In contrast, at late time points, CX3CR1^−/− CD4⁺ T cells were slightly enriched over CX3CR1^+/+ Trm, suggesting a small role for CX3CR1 in limiting Trm formation or persistence. These effects on Trm are, however, very small, making it doubtful that they would have much biological significance.

The chemokine receptor CX3CR1 has been studied as a marker of distinct CD8⁺ T cell subsets during viral infections. CX3CR1 expression correlates with the degree of effector CD8⁺ T cell differentiation (15, 25), and CX3CR1⁺ CD8⁺ T cells define the terminally differentiated cytotoxic effector cells (15, 26). It has also been shown that CX3CR1⁺ CD8⁺ T cells are required to control chronic viral infection (27); however, no changes in the functional phenotype of CX3CR1-deficient memory CD8⁺ T cells were observed in another study (28), suggesting that CX3CR1-mediated signals are not the cause of direct cytotoxic effector functions in memory T cells. We also show in this study that CX3CR1 marks the most activated splenic CD4⁺ Th1 effectors, consistent with previously reported findings for CD8⁺ T cells (15). The CX3CR1^hi CD4⁺ T cells had about a 2-fold higher frequency of T-bet⁺ cells, with a slightly increased MFI for T-bet. The CX3CR1^hi CD4⁺ T cells also had a modest increase in the proportion of IFN-γ⁺ among the Th1 cells, with about a 2-fold effect on IFN-γ MFI compared with the CX3CR1^lo/int CD4⁺ T cells. We also observed an increased proportion of granzyme-positive and of perforin-positive CD4 T cells in the CX3CR1^hi SMARTA effector population. Of note, after acute dengue virus infection, CX3CR1 also marks a cytotoxic subset of effector memory CD4⁺ T cells (17).

The chemokine receptor CX3CR1 and its ligand, fractalkine (CX3CL1), are known for directing leukocyte migration from the bloodstream into tissues during infections; however, their role in T cells has been more enigmatic (29–31). In this study, we showed that CX3CR1 is dispensable for CD4⁺ T cell trafficking to the spleen, lung, or liver by the peak of the response 8 d after LCMV13 infection. Consistently, previous work has demonstrated using adoptive transfer of CX3CR1-proficient and CX3CR1-deficient Th2 cells into WT mice that CX3CR1 is dispensable for T cell migration into the airways in a mouse asthma model (12). Moreover, during Mycobacterium tuberculosis infection, CX3CR1 deficiency in effector CD4⁺ T cells enhanced (by ∼2-fold), rather than decreased, the rate of T cell lung entry (32), whereas other chemokine receptors such as CXCR3, CXCR6, CCR2, and CCR5 were implicated in T cell trafficking to tissues during M. tuberculosis infection (32). Thus, the evidence that CX3CR1 is important in T cell trafficking is limited, a finding that is further supported by the current study.

Although CX3CR1 did not affect T cell accumulation in the tissues, we found that CX3CR1 deficiency marginally affected the proportion of those cells that acquire a Th1 phenotype. Whether this is due to an effect of CX3CR1 on the differentiation of the Th1 cells or an effect on the survival of the more differentiated effectors was not investigated in this study. However, in the literature CX3CR1, which is required for airway disease in asthma models, was shown to provide a survival signal to Th2 cells through Bcl-2 (12). Interestingly, Sallin et al. (33) found that intravascular CX3CR1⁺ CD4⁺ T cells were nearly completely absent from p40^−/− mice during M. tuberculosis infection and that IL- 12/23p40 and T-bet were involved in the production of a subset of terminal effector–like CX3CR1⁺ KLRG1⁺ Th1 cells. Whether CX3CR1⁺ cells respond better to IL-12 or whether CX3CR1 signaling directly leads to cell differentiation with increased expression of T-bet and IFN-γ remains to be investigated.

At the memory phase of chronic LCMV infection, we observed that there was a slightly higher proportion of CX3CR1^−/− CD4⁺ Trm in both lung and spleen compared with their WT counterparts after correcting for preinfection ratios, whereas total CD4⁺ T cells did not show this difference. This finding is in line with previous reports showing that within CD8⁺ T cells, the CX3CR1^-/low subset is the least differentiated and gives rise to Trm (15, 34). T-bet induces IFN-γ production and Th1/Tc1 differentiation and is more highly expressed in terminally differentiated effector T cells than in memory precursors (35, 36). Moreover, in the absence of T-bet, there is a striking increase in parenchymal CD69⁺ CD103⁺ CD4⁺ T cells during M. tuberculosis infection, consistent with the view that T-bet suppresses the development of CD69⁺ CD103⁺ tissue-resident phenotype effectors in lung (33). Similarly, T-bet overexpression prevents the development of skin tissue-resident memory CD8⁺ T cells during acute HSV infection (37). In the current study, we observed that CX3CR1-deficient CD4⁺ T cells expressed slightly lower levels of T-bet compared with their WT counterparts during the effector phase, which might explain why the CX3CR1^−/− CD4⁺ population is found in slightly greater proportion than the CX3CR1^+/+ CD4⁺ Trm population. Whether CX3CR1 contributes to formation, survival, and/or maintenance of Trm was not addressed because of the marginal nature of this phenotype.

In sum, in the current study, we have shown that CX3CR1 marks the most differentiated Th1 effector population during LCMV13 infection of mice. CX3CR1 is dispensable for T cell accumulation in the spleen, lung, and liver, arguing against a major role in trafficking or survival of CD4⁺ T cells during LCMV13 infection. CX3CR1 deficiency marginally decreases the proportion of cells with a T-bet^hi Th1 phenotype at the effector stage and marginally increases the proportion of cells that acquire a Trm phenotype at day 30 p.i. Taken together, these studies show CX3CR1, although useful as a marker of effector cell differentiation, has a minimal role in T cell responses to a chronic viral infection.

We thank Birinder Ghumman for technical support and Dionne White and Joanna Warzyszynska for assistance with flow cytometry.

The authors have no financial conflicts of interest.