MARCH1 Controls an Exhaustion-like Program of Effector CD4⁺ T Cells Promoting Allergic Airway Inflammation

Professional APCs contribute to allergic airway inflammation by sustaining Th cell activation and proliferation (1). Dendritic cells (DCs) capture inhaled allergens and present them to naïve CD4⁺ T cells in the mediastinal lymph node, inducing development of type 2 Th (Th2) cells (2). However, full development of Th2 cells is not completed in the lymph node (3, 4). The tissue of allergen exposure constitutes an important site for terminal differentiation and effectuation of Th2 cells (5). Tissue conditioning enables T cell production of IL-5 and IL-13 in high levels, which drives recruitment of eosinophils to the airway, establishing inflammation (6). Previous studies have shown that tissue DCs and macrophages are important contributors to effector inflammatory responses in allergic asthma. A human cohort study has shown that asthma characterized by high amounts of Th2 cytokines and eosinophils in the airway is accompanied by accumulation of DCs in the airway (7, 8). A mouse asthma model study has shown that depletion of airway DCs during allergen challenge abolished Th2 cell and eosinophilic inflammation, goblet cell hyperplasia, and bronchial hyperreactivity, identifying DCs as key immune cells necessary for Th2 cell stimulation during ongoing airway inflammation (9). Another mouse study demonstrated that depletion of macrophages impaired local Th2 cell homing and activation in the inflamed lung during IL-13–dependent airway inflammation (10). However, the molecular mechanisms by which DCs and macrophages mediate continuous Th2 cell activation in the airway remain poorly understood.

DCs and other APCs use MHC class II (MHCII) Ag-presenting molecule and the costimulatory molecule CD86 to stimulate Ag-specific CD4⁺ T cells (11). MHCII and CD86 are both posttranslationally modified by a ubiquitin ligase named membrane-associated RING-CH1 (MARCH1) (12–14). MARCH1 attaches ubiquitin chains to the cytosolic lysine residues of MHCII and CD86, inducing endocytosis and subsequent degradation (13, 15). MARCH1-mediated turnover of MHCII promotes thymic DC selection of regulatory T cells and promotes splenic DC activation of naive CD4⁺ T cells (16–18). Yet the role of MARCH1 in APC function in nonlymphoid tissues such as the airways is not known. Because MARCH1 mediates clearance of MHCII and CD86 from the APC surface, MARCH1 may control activation of tissue effector CD4⁺ T cells and negatively regulate CD4⁺ T cell–dependent tissue inflammation. Conversely, MARCH1 may sustain effector CD4⁺ T cell responses by preventing APCs from providing the cells with excessive and/or prolonged antigenic and costimulatory signals because such strong signals could exhaust T cells resulting in poor proliferation and cytokine production (19). In this study, we investigated how MARCH1 affects lung APC function of driving and maintaining Th2 cell–dependent inflammation in a mouse model of allergic asthma.

Wild-type (WT) C57BL/6J and B6.SJL-PtprcaPepcb/BoyJ (CD45.1⁺ “BoyJ”) mice were purchased from The Jackson Laboratory and maintained in our colony. MARCH1^−/−, MARCH1^fl/fl, CD11c^Cre, MHCII^K>R, CD86^K>R, and UBC^ERT2-Cre mice were previously described (14, 16, 17, 20–22). MARCH1^fl/flCD11c^Cre, MHCII^K>RCD86^K>R, or MARCH1^fl/flUBC^ERT2-Cre mice were generated by breeding MARCH1^fl/fl mice with CD11c^Cre mice, MHCII^K>R mice with CD86^K>R mice, or MARCH1^fl/fl mice with UBC^ERT2-Cre mice, respectively. Mice were housed in a specific pathogen-free facility in the Laboratory Animal Research Center at University of California, San Francisco (UCSF). Mice of both sexes were used at 5–20 wk of age when initiating an experiment and whenever possible, littermates or mice of distinct genotypes were cohoused. Sample sizes were guided by previous studies. Mouse genotypes were not blinded from the investigator. All experiments conformed to ethical principles and guidelines approved by the Institutional Animal Care and Use Committee at UCSF.

On days 0 and 7–11, mice were anesthetized with isoflurane and given 10 μg house dust mite allergen (HDM) extract (Dermatophagoides pteronyssinus; Stallergenes Greer) by oropharyngeal aspiration. On day 14, mice were euthanized, and lungs were collected for flow cytometry. For experiments involving tamoxifen, mice were also challenged on the days of tamoxifen administration with 10 μg HDM extract and euthanized on day 49.

Lungs were digested with Collagenase D (Roche) and DNase I (Roche) for 30 min at 37°C and dissociated in C tubes with a gentleMACS Dissociator (Miltenyi Biotec). The digest was stopped with cold FACS buffer (0.5% BSA in PBS) and centrifuged at 500 × g for 5 min at 4°C. Cell pellets were resuspended in RBC lysis buffer for 5 min at room temperature and washed with FACS buffer. Cell suspensions were filtered with 70-μm nylon mesh strainers. Cells were centrifuged and resuspended in 1 ml FACS buffer and counted with a Z2 Particle Counter (Beckman Coulter). Cells then were stained for surface and/or intracellular markers with mixtures of fluorescent Abs (Supplemental Table I). In some experiments, cells were stimulated ex vivo with PMA (10 ng/ml; Sigma-Aldrich) plus ionomycin (1 μg/ml; Sigma-Aldrich) in the presence of brefeldin A (10 μg/ml; Sigma-Aldrich) for 3 h in a 37°C CO₂ incubator before surface and intracellular staining.

T cells were isolated using the EasySep CD4⁺ T cell Isolation Kit (STEMCELL Technologies) from single-cell suspensions of lungs derived from CD45.1⁺ donor mice at day 14 of HDM-induced allergic asthma protocol. For each experiment, CD4⁺ T cells were pooled from multiple donor mice. Purified T cells were labeled with CellTrace Violet (CTV) (Life Technologies) for 20 min at 37°C and quenched with RPMI supplemented with 10% FBS. Cells (∼0.5–1 × 10⁶) were transferred i.v. into recipient mice. Recipient mice were administered 10 μg HDMs on days 0–3 posttransfer. On day 4, recipient mice were euthanized, and lungs were collected for determining proliferation of donor T cells by staining cells for surface markers, and CTV dilution was detected by flow cytometry.

Tamoxifen was infused in corn oil and injected i.p. at a dose of 50 μg/g.

Single-cell RNA sequencing (scRNA-seq) libraries were prepared according to the 10× Chromium Next GEM 3′ kit according to the manufacturer’s instructions. In brief, FACS-sorted cells were washed once with PBS + 0.04% BSA and resuspended in PBS + 0.04% BSA. After reverse transcription and cell barcoding in droplets, emulsions were broken, and cDNA was purified using Dynabeads MyOne SILANE followed by PCR amplification. For gene expression library construction, amplified cDNA was used for fragmentation, followed by end-repair, double-sided size selection with SPRIselect beads, PCR amplification with sample indexing primers, and double-sided size selection with SPRIselect beads. Sequencing was performed on a MiSeq (Illumina).

Reads from 10× scRNA expression libraries were aligned to the mouse genome assembly GRCm38 (mm10) and quantified using cellranger count (version 3.1.0; 10× Genomics). The filtered feature-barcode matrices containing only cellular barcodes were used for further analysis. Single-cell gene expression matrices were imported into R and analyzed using Seurat (version 4.0). Cells were filtered for high mitochondrial RNA content (keep cells < 5% mitochondrial gene expression) and number of genes captured (keep cells > 800 and < 2500). Each cell was assigned a score for S and G2/M cell-cycle phases based on previously defined gene sets using the CellCycleScoring function. Scaled Z scores for each gene were calculated using the ScaleData function and regressed against the number of unique molecular identifiers per cell, S phase score, and G2/M phase score. Scale data were used as input into a principal-component analysis on the basis of variable genes. Dimensionality reduction was performed using Uniform Manifold Approximation and Projection (UMAP) with the first eight principal components. To compare cells with previously described transcriptional states, we computed gene signature scores for each cell with Seurat’s AddModuleScore() function using gene signatures from Crawford et al. (23) for exhaustion score and from Tibbitt et al. (4) for Th2 module score. For the exhaustion signature score, inhibitory surface molecules overexpressed in exhausted CD4⁺ T cells during lymphocytic choriomeningitis virus clone 13 infection compared with naive CD4⁺ T cells (Ctla4, Pdcd1, Lag3, Pdcd1lg2, Cd274, Tigit, Cd200, Havcr2) were used. For the Th2 module score, the top 15 genes expressed by bronchoalveolar lavage Th2 cells after HDM challenge (Igfbp7, Il13, Il1rl1, Plac8, Bhlhe40, Gata3, Nfkb1, Rbpj, Fgl2, Ltb4r1, Epas1, Il5, Zcchc10, Hlf, Mns1) were used. scRNA-seq data have been deposited in the Gene Expression Omnibus (https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE211511) under accession number GSE211511.

Flow cytometry data were analyzed by FlowJo (TreeStar). Graphs were drawn by Prism (GraphPad). Unpaired two-tailed Student t test was performed to assess the significance for most of the data. For the scRNA-seq exhaustion score and Th2 module score data, statistical significance between groups was calculated by Wilcoxon rank-sum test.

WT and MARCH1-deficient mice were administered with HDMs as shown in (Fig. 1A. On day 14, at the peak of allergic inflammation, lung APCs in MARCH1-deficient mice displayed higher levels of MHCII and CD86 on the surface than those in WT mice, indicating that MARCH1 regulates the surface abundance of MHCII and CD86 in lung APCs (Fig. 1B). Despite higher levels of MHCII and CD86 on lung APCs, the number of effector CD4⁺ T cells (CD44^hiCD62L^loCD4⁺ cells) was significantly lower in MARCH1-deficient mice compared with WT mice (Fig. 1C, Supplemental Fig. 1A). We hypothesized that the increased MHCII and CD86 on APCs in MARCH1-deficient mice might have induced exhaustion of effector CD4⁺ T cells, leading to impaired proliferation. To test this possibility, we examined the expression of PD-1, CTLA-4, TIGIT, TIM-3, or LAG-3 immune-inhibitory receptors upregulated during T cell exhaustion (19). The percentage of effector CD4⁺ T cells upregulating each of these inhibitory receptors was higher in MARCH1-deficient mice than in the WT mice (Fig. 1D, Supplemental Fig. 1B). Expression of multiple inhibitory receptors is a cardinal feature of T cell exhaustion (19). The percentage of effector CD4⁺ T cells coexpressing multiple inhibitory receptors was also higher in MARCH1-deficient mice than in WT mice (Fig. 1E), implicating greater T cell exhaustion in MARCH1-deficient mice. To determine whether upregulation of these inhibitory receptors is associated with functional exhaustion, we examined proliferative status of the effector CD4⁺ T cells by determining the expression of Ki-67, a marker indicative of cell-cycle progression. The percentage of Ki-67⁺ effector CD4⁺ T cells was significantly lower in MARCH1-deficient mice than in WT mice (Fig. 1F, Supplemental Fig. 1C). These results strongly suggest that MARCH1 restrains exhaustion of effector CD4⁺ T cells during allergic airway inflammation.

To determine whether the role of MARCH1 in restraining exhaustion of effector CD4⁺ T cells is dependent on the expression of this protein in DCs and/or macrophages, but not B cells or other unconventional APCs, we performed similar experiments using MARCH1^fl/flCD11c^Cre mice in which Cre recombinase is expressed under the control of the promoter of CD11c (9, 20, 24), a marker of murine DCs and lung macrophages (Supplemental Fig. 2A). As expected, the lung myeloid APCs of MARCH1^fl/flCD11c^Cre mice displayed higher levels of MHCII and CD86 on the surface than those of MARCH1^fl/fl mice during allergic airway inflammation (Supplemental Fig. 2B). The number of effector CD4⁺ T cells was lower in MARCH1^fl/flCD11c^Cre mice than in MARCH1^fl/fl mice (Supplemental Fig. 2C). The percentage of effector CD4⁺ T cells that express inhibitory receptors such as CTLA-4, TIGIT, and TIM-3 was significantly higher in MARCH1^fl/flCD11c^Cre mice than in MARCH1^fl/fl mice (Supplemental Fig. 2D). The percentage of effector CD4⁺ T cells that concurrently express multiple inhibitory receptors was also higher in MARCH1^fl/flCD11c^Cre mice than in MARCH1^fl/fl mice (Supplemental Fig. 2E), implicating greater T cell exhaustion in MARCH1^fl/flCD11c^Cre mice. Lastly, the frequency of Ki-67–expressing effector CD4⁺ T cells was lower in MARCH1^fl/flCD11c^Cre mice than in MARCH1^fl/fl mice (Supplemental Fig. 2F). These findings indicate that the role of MARCH1 in restraining exhaustion of effector CD4⁺ T cells during allergic airway inflammation is largely dependent on the expression of this protein in DCs and/or macrophages, but not B cells or other APCs.

MARCH1 is not expressed in T cells (14); however, MARCH1 deficiency might have caused a defect in the development of effector CD4⁺ T cells in a T cell–extrinsic manner, and this defect could have resulted in functional abnormality of the cells in MARCH1-deficient mice. To bypass this potential problem, we isolated effector CD4⁺ T cells from CD45.1⁺ WT “BoyJ” mice treated with HDMs, labeled the cells with CTV dye, and transferred the cells to either WT or MARCH1-deficient mice that express CD45.2. These recipient mice were subsequently challenged with HDMs, and proliferative responses of transferred T cells were examined by determining CTV dilution (Fig. 2A). We found the extent of CTV dilution was significantly lower in MARCH1-deficient mice than in WT mice (Fig. 2B, 2C). When the same experiment was performed using MARCH1^fl/flCD11c^Cre mice and MARCH1^fl/fl mice, the transferred T cells proliferated less in MARCH1^fl/flCD11c^Cre mice than in MARCH1^fl/fl mice (Fig. 2D). These findings demonstrate that MARCH1 in myeloid APCs significantly contributes to promoting tissue proliferation of effector CD4⁺ T cells during allergic airway inflammation.

Then, we considered whether the adoptively transferred cells in the MARCH1-deficient mice exhibited an exhaustion signature as we had observed with endogenous effector CD4⁺ T cells during allergic airway inflammation (Fig. 1D, 1E). To address this, we investigated the gene expression profile of the proliferating transferred cells in WT or MARCH1-deficient mice, which were sorted and subjected to scRNA-seq. HDM-reactive transferred cells in the WT or in the MARCH1-deficient recipients were visualized by dimensionality reduction in a UMAP plot (Fig. 3A). Expression of genes such as Ctla4, Pdcd1, and Lag3 encoding for the inhibitory receptors CTLA-4, PD-1, and LAG-3, respectively, was higher in the transferred cells in MARCH1-deficient mice than in WT mice (Fig. 3B, 3C). We computed an exhaustion signature score based on inhibitory surface molecules overexpressed in exhausted CD4⁺ T cells during lymphocytic choriomeningitis virus clone 13 infection (23) and extrapolating this to the HDM-reactive cells showed that the transferred cells in the MARCH1-deficient mice exhibited a higher exhaustion score than in the WT recipients (Fig. 3D). The transferred cells in the MARCH1-deficient recipients were also expressing less Il13, Il4, and Il5 encoding for effector cytokines IL-13, IL-4, and IL-5, respectively (Fig. 3E). We calculated a Th2 module score based on top genes expressed in Th2 cells during HDM challenge (4), and the Th2 module score was lower for the transferred cells in the MARCH1-deficient mice (Fig. 3F), suggesting loss of effector function. These findings corroborate a role for MARCH1 in suppressing exhaustion of effector CD4⁺ T cells, which coincides with a vigorous proliferative and effector response of these cells during allergic inflammation.

Next, we sought to identify the molecular mechanisms by which MARCH1 suppresses exhaustion of effector CD4⁺ T cells and promotes proliferation of the cells. Our hypothesis was that MARCH1-dependent ubiquitination of MHCII and CD86 allows APCs to express MHCII and CD86 in a regulated fashion, which in turn allows the effector CD4⁺ T cells to express inhibitory receptors in a controlled manner eliciting robust proliferative responses. To test this hypothesis, we generated MHCII^K>RCD86^K>R mice where all the ubiquitin-accepting lysines (K) of MHCII and CD86 were mutated to arginines (R) (16, 21). MHCII^K>RCD86^K>R mice and WT mice were treated with HDMs (Fig. 4A). We found that lung APCs in MHCII^K>RCD86^K>R mice displayed increased abundance of MHCII and CD86 on the surface compared with those in WT mice, which confirms that surface abundance of these two molecules is indeed regulated by ubiquitination (Fig. 4B). Remarkably, the number of effector CD4⁺ T cells was significantly low in MHCII^K>RCD86^K>R mice compared with WT mice (Fig. 4C). The frequency of effector CD4⁺ T cells that express PD-1, CTLA-4, TIGIT, or TIM-3 was higher in MHCII^K>RCD86^K>R mice than in WT mice (Fig. 4D). The percentage of the effector CD4⁺ T cells that concurrently express multiple inhibitory receptors was also higher in MHCII^K>RCD86^K>R mice than in WT mice (Fig. 4E). The frequency of Ki-67⁺ effector CD4⁺ T cells was low in MHCII^K>RCD86^K>R mice compared with WT mice (Fig. 4F). MHCII^K>RCD86^K>R mice were also inferior in supporting proliferation of HDM-experienced CD4⁺ T cells compared with WT mice (Fig. 4G, 4H). This poor proliferation was accompanied by increased expression of the inhibitory molecules PD-1 and TIM-3 by the T cells in the MHCII^K>RCD86^K>R mice (Fig. 4I). Taken together, ubiquitination of MHCII and CD86 plays a significant role in restraining exhaustion of effector CD4⁺ T cells during allergic airway inflammation.

The findings suggesting that the ubiquitin ligase activity of MARCH1 promotes the proliferation of effector CD4⁺ T cells during allergic airway inflammation led us to hypothesize that inhibiting MARCH1 may exert a therapeutic benefit in allergic asthma by ameliorating inflammation driven by Th2 cells, the pathologic effector CD4⁺ T cells in this disease (6). We conceived a proof-of-principle experiment that uses a mouse strain (MARCH1^fl/flUBC^ERT2-Cre) where the expression of MARCH1 can be abolished in an inducible manner with tamoxifen. MARCH1^fl/flUBC^ERT2-Cre mice and the control UBC^ERT2-Cre mice were administered for 2 wk with HDMs alone, which would induce allergic asthma to both groups of mice. From the third week, the mice were administered with HDMs and tamoxifen, which would disable the expression of MARCH1 selectively in MARCH1^fl/flUBC^ERT2-Cre mice. After 7 wk of HDM treatment in total (Fig. 5A), we found that the surface level of MHCII and CD86 was markedly higher in APCs of MARCH1^fl/flUBC^ERT2-Cre mice than that of UBC^ERT2-Cre mice, which confirms the ablation of MARCH1 in MARCH1^fl/flUBC^ERT2-Cre mice (Supplemental Fig. 3A). We also found that the effector CD4⁺ T cells in MARCH1^fl/flUBC^ERT2-Cre mice expressed PD-1, TIM-3, and TIGIT to a higher frequency than those in UBC^ERT2-Cre mice (Supplemental Fig. 3B). The frequency of effector CD4⁺ T cells that concurrently express multiple inhibitory receptors was also higher in MARCH1^fl/flUBC^ERT2-Cre mice (Supplemental Fig. 3C). This finding is consistent with the role of MARCH1 in restraining the exhaustion of effector CD4⁺ T cells. Next, we examined the frequency of IL-4–, IL-13–, and IL-5–producing Th2 cells, which drive inflammation in allergic asthma. We also determined the frequency of eosinophils, which also contribute majorly to inflammation in allergic asthma. We found the numbers of Th2 cells and eosinophils were both significantly lower in MARCH1^fl/flUBC^ERT2-Cre mice compared with UBC^ERT2-Cre mice (Fig. 5B, 5C). These findings support our hypothesis that inhibiting MARCH1 could be an effective strategy for ameliorating airway inflammation associated with allergic asthma.

Collectively, we conclude that MARCH1-dependent control of MHCII and CD86 grants tissue APCs an ability to sustain robust effector CD4⁺ T cell responses by limiting a T cell exhaustion-like program, and that this function plays a significant role in sustaining airway inflammation associated with allergic asthma. Mice deficient in either MARCH1 or the ubiquitin acceptor sites of MHCII and CD86 elicited poor effector CD4⁺ T cell responses compared with WT mice. The effector CD4⁺ T cells in these mice upregulated the expression of a broad range of inhibitory receptors and proliferated poorly, which are characteristic of exhausted T cells (19). We propose that MARCH1 limits MHCII and CD86 ligand availability on the APC surface and thus prevents persistent antigenic signaling to responding effector T cells, which restrains the effector cells from becoming exhausted.

Kim et al. (18) have shown that MARCH1-mediated ubiquitination of MHCII promotes DC function of inducing naive CD4⁺ T cell proliferation in the spleen, although a role for MARCH1 during an effector CD4⁺ T cell response was not addressed. We show in this study that MARCH1 ubiquitination of MHCII and CD86 functions in APCs to support effector Th2 cell responses in the lung independent of a primary response. In addition, a previous study has shown that MARCH1-deficient mice exhibited defective Th2 cell responses but competent eosinophilia in the lung (25). The discrepancy of this study from ours may represent inherent differences in the mouse models of asthma. We did not address the distinct or redundant contribution of MARCH1 among myeloid APCs (i.e., DC or macrophage subsets), although our previous work showed that MARCH1 in the lymph node resident DCs plays a critical role for Th2 cell priming and sequential Th2 cell responses (26). This previous work did not exclude a role for MARCH1 among lung APCs (i.e., DC or macrophage subsets), which we have shown in this study could also contribute to effectuation of a Th2 cell response. Although the question of the dependency of MARCH1 among individual lung APC subsets remains to be addressed, we identify MARCH1 as a key player for lung APCs to mediate optimal effector CD4⁺ T cell responses and propose inhibiting MARCH1 could bring a therapeutic benefit for the control of allergic asthma.

We thank H. K. Li (UCSF) for mouse colony management, S. L. Gonzalez (UCSF) for performing preliminary experiments, and K. M. Ansel (UCSF), R. M. Locksley (UCSF), and R. A. Rahimi (Massachusetts General Hospital, Harvard) for helpful discussion.

A.T.S. is a scientific founder of Immunai and founder of Cartography Biosciences and receives research funding from Allogene Therapeutics and Merck Research Laboratories.