MHC Class II Ubiquitination Regulates Dendritic Cell Function and Immunity

Major histocompatibility class II (MHC II) molecules present peptide Ags derived from the endosomal pathway to CD4⁺ T cells. This is essential for T cell activation during immune responses, but is also essential in thymic T cell selection and T cell maintenance in peripheral lymphoid organs. The stability and abundance of peptide-loaded MHC II (pMHC II) at the cell surface of APCs is regulated by ubiquitination. Ubiquitination occurs on lysine 225 (K225) in the β-chain of MHC II (1–3). The E3 ligase responsible for pMHC II ubiquitination in hematopoietic cells, including B cells and dendritic cells (DCs), is membrane-associated RING-CH-type finger (MARCH) 1 (4–6 and P. Schriek, H. Liu, A. C. Ching, P. Huang, N. Gupta, K. R. Wilson, M. Tsai, Y. Yan, C. F. Macri, L. F. Dagley, et al., manuscript posted on bioRxiv, DOI: 2021.04.15.439921), whereas MARCH8 is responsible for ubiquitination in thymic epithelial cells (7, 8, and P. Schriek et al., manuscript posted on bioRxiv, DOI: 2021.04.15.439921) and type II alveolar epithelial cells (P. Schriek et al., manuscript posted on bioRxiv, DOI: 2021.04.15.439921). Ubiquitination targets pMHC II for lysosomal degradation (9). As a result, DCs from mice lacking MARCH1 (Marchf1^−/−) (6, and P. Schriek et al., manuscript posted on bioRxiv, DOI: 2021.04.15.439921) or expressing MHC II molecules with K225 mutated to R (MHC IIKR^KI/KI), which cannot be ubiquitinated, have increased surface MHC II expression (10). Despite surface MHC II being critical for T cell development, mice that lack MHC II ubiquitination have only minor alterations in their T cell compartments. We and others reported reduced regulatory T cells (11–13), but there is little impact on steady-state conventional T cell populations (11–13). The functional consequences for immunity are unclear, although little analysis has been carried out to address this question. This is surprising, given the important role of APCs, particularly DCs, in initiating immunity.

An appealing hypothesis is that MHC II ubiquitination limits Ag presentation to prevent detrimental outcomes. In vitro Ag-presentation assays that investigate this scenario are inconclusive, showing increased (9), unaltered (14), or reduced (15–17) CD4⁺ T cell priming in response to MHC IIKR^KI/KI DC Ag presentation. This inconsistency may be due to differences in cell types, with some studies using bone marrow–derived DC (BMDC) (9, 14, 16) and others using bulk CD11c⁺ DC (15, 17). Moreover, the impact of MHC II ubiquitination on MHC class I (MHC I) presentation has largely been ignored. We have previously described that a lack of MHC II ubiquitination reduces MHC I surface expression and impairs cross-presentation in vitro (18). From these studies, it is clear that an in-depth in vivo investigation is required to fully elucidate the impact of MHC II ubiquitination on DC Ag presentation and immunity.

In this study, we have systematically investigated the role of MHC II ubiquitination on primary DC function and in vivo immune outcomes. We have identified that in mice lacking MHC II ubiquitination there is a decrease in the number of splenic DCs. The remaining DCs have an altered surface phenotype accompanied by defects in Ag proteolysis and cytokine secretion. Functionally, these changes reduce in vivo CD4⁺ and CD8⁺ T cell priming in response to soluble, cell-associated and DC-targeted Ag. Importantly, ex vivo T cell priming revealed these changes were not simply due to reduced DC numbers but an intrinsic defect in DC function. Additionally, impairment in CTL killing, T follicular helper (T_FH) cell generation, and Ab responses were observed in MHC IIKR^KI/KI mice following immunization. Our results demonstrate the importance of DC MHC II ubiquitination for effective immunity.

C57BL/6, MHC IIKR^KI/KI (12), IAα^−/− (19), OT-II × Ly5.1 and OT-I × Ly5.1 mice were used at 6–12 wk of age. All mice were bred and maintained in specific pathogen–free conditions at the Melbourne Bioresources Platform at Bio21 Molecular Science and Biotechnology Institute. Experimental procedures were approved by the Animal Ethics Committee of the University of Melbourne (1714375).

Primary splenic or thymic DCs were isolated as previously described (20). Splenic conventional DCs (cDCs) were stained with mAbs specific for CD11c (N418), CD8α (53-6.7), and CD11b (M1/70) (all BioLegend). Plasmacytoid DC (pDC) preparations were stained with BST-2 (927) and Siglec-H (551) (both BioLegend). Thymus DCs were identified with mAbs specific for CD11c (N418), CD8α (53-6.7), CD11b (M1/70), B220 (RA3-6B2), Sirpα (all BioLegend), and NK1.1 (PK136; BD Biosciences). In some experiments, cDC1 (CD11c⁺ CD8⁺ CD11b⁻) and cDC2 (CD11c⁺ CD8⁻ CD11b⁺) were sorted to purity by flow cytometry on a BD Influx (Murdoch Children’s Research Institute Flow Cytometry and Imaging Facility). To obtain absolute cell numbers, an internal microsphere counting standard (BD Biosciences) was used.

For determining the expression of cell surface markers, single-cell suspensions were generated. Splenocytes were either treated with RBC removal buffer, or light cells were separated by density gradient separation in 1.077 g/cm³ Nycodenz (Nycomed). Cells were stained with mAbs specific for CD3 (KT3-1.1), CD8α (53-6.7), CD11b (M1/70), CD11c (N418), CD19 (6D5), CD24 (M1/69), CD40 (FGK45.5), CD45R/B220 (RA3-6B2), CD80 (16-10A1), CD86 (GL1), CD135/Flt3 (A2F10), CD172 (Sirpα), CD205/DEC205 (NLDC-145), CD274/PD-L1 (10F.9G2), CD317/BST2 (927), CD370/Clec9A (7H11), H-2Kb (AF6-8815), IgD (11-26c.2a), IgM (RMM-1), Siglec-H (551), XCR1 (ZET) (all BioLegend), Ly-6C.2 (5075-3.6), or Ly-6G (1A8) (Walter and Eliza Hall Antibody Facility). Stained cells were acquired on the LSRFortessa (BD Biosciences) flow cytometer and analyzed on FlowJo software (Tree Star), with exclusion of cell doublets and dead cells in all cases identified based on forward and side scatter as well as staining with propidium iodide or DAPI. Statistical analysis was performed with Prism software.

Splenic cDC1 and cDC2 were isolated and sorted to purity. A total of 1 × 10⁵ DCs were cultured in completed medium (RPMI 1640 medium supplemented with 10% heat-inactivated FCS, 1% penicillin/streptomycin, 50 nM 2-ME, and 1% GlutaMAX) with or without 0.5 μM phosphorothioated CpG 1668 (Bioneer), 50 ng/ml IFN-γ (PeproTech), and 20 ng/ml GM-CSF (PeproTech) or 0.5 μM CpG alone. Supernatants were collected 18 h later and stored at −20°C. Cytokine secretion was determined using the BD Cytometric Bead Array Mouse Inflammation Kit according to the manufacturer’s instructions (BD Biosciences).

To assess Ag uptake, 5 × 10⁴ purified cDC1 or cDC2 were incubated with 50 μg/ml OVA-Cy5 in complete medium for 0–90 min at 37°C. DCs were washed twice, and the fluorescence was measured by flow cytometry. As a control, DCs were pulsed with OVA-Cy5 for 90 min at 4°C. To assess Ag proteolysis and endosomal pH, 2.5 × 10⁵ purified DCs were incubated with 20 μg/ml DQ-OVA (Life Technologies) or with 20 μg/ml dextran-pHrodo (Life Technologies) for 15 min at 37°C. Cells were washed twice, resuspended in complete medium and chased for the indicated time. At each time point, cells were kept on ice, and the fluorescence was measured by flow cytometry. As a control, DCs were pulsed with DQ-OVA or dextran-pHrodo for 15 min at 4°C, washed twice, and analyzed by flow cytometry. The signal was calculated for the 37°C chase time point by subtracting the geometric mean fluorescence intensity (gMFI) at 4°C. To normalize for dextran-pHrodo internalization, we simultaneously measured uptake of dextran-A647.

Single-cell suspensions were generated from lymph nodes of OT-I × Ly5.1 or OT-II × Ly5.1 mice. Cells were stained with rat anti-mouse mAbs specific for CD11b (M1/70), F4/80 (F4/80), RBCs (TER119), Ly-6G/Ly-6C (RB68C5), MHC II (M5/114), CD45R (RA36B2), and CD4 (GK1.5) for OT-I T cells or CD8α (53-6.7) for OT-II cells. Cells were washed and incubated with BioMag anti-rat IgG-coupled magnetic beads (QIAGEN). After magnetic depletion, the CD4⁺ or CD8⁺ T cell–enriched supernatant was recovered. Cells were washed with PBS supplemented with 0.1% BSA and labeled with CellTrace Violet (CTV; Thermo Fisher Scientific). To enable accurate quantification of the number of OT-II or OT-I cells, mice also received transfer of a control population of CFSE (Thermo Fisher Scientific)–labeled splenocytes. Single-cell suspensions were treated with RBC removal buffer. Cells were washed with PBS, 0.1% BSA, and labeled with CFSE. Equal numbers of CTV-labeled OT-II or OT-I cells and CFSE-labeled splenocytes were pooled and resuspended in RPMI 1640, 2% FCS at 1 × 10⁷ cells/ml, respectively, and injected i.v. Twenty-four hours later, mice received 50 µg (for OT-II–transferred mice) or 25 µg (for OT-I–transferred mice) soluble OVA (Sigma-Aldrich), 20 × 10⁶ OVA-coated splenocytes, or 0.2 µg anti-Clec9A-OVA mAb (21) via i.v. injection. For OVA-coated splenocytes, single-cell suspensions were generated from spleens of C57BL/6 mice treated with RBC removal buffer, washed, and incubated with 10 mg/ml OVA protein (Sigma-Aldrich) in FCS-free RPMI 1640 at 37°C. Sixty-four hours after immunization, spleens were harvested, single-cell suspensions were generated, and RBCs were lysed. Cells were stained with mAbs specific for CD8α (53-6.7, BioLegend) or CD4 (GK1.5; Walter and Eliza Hall Antibody Facility) and Ly-5.1 (A20; BioLegend). The number of divided OT-II and OT-I was determined as the number of CD4⁺ or CD8⁺ Ly-5.1⁺ cells that had undergone CTV dilution. The number of cells was normalized to the number of CFSE-labeled splenocytes recovered.

Mice were immunized i.v. with 1 μg of anti-Clec9A-OVA mAb (22). One day later, spleens were harvested, and DC were isolated, as previously described, before sorting to purity. Sorted cDC1 (CD11c⁺ CD8⁺ CD11b⁻) or cDC2 (CD11c⁺ CD8⁻ CD11b⁺) (numbers indicated in figures) were cultured in vitro with 2.5 × 10⁴ CTV-labeled OT-II or OT-I cells in complete medium. Four or five days later, the number of divided OT cells was determined.

DCs were isolated as previously described, and cDC1 (CD11c⁺ CD8⁺ CD11b⁻) and cDC2 (CD11c⁺ CD8⁻ CD11b⁺) were sorted to purity. A total of 2.5 × 10⁴ sorted cDC1 or cDC2 were cultured in vitro with 2.5 × 10⁴ CTV-labeled OT-II cells and OVA_(323–339) peptide. Sixty-eight hours later, the number of divided OT-II cells was determined by flow cytometry.

Mice were injected with 2 × 10⁷ OVA-coated splenocytes (prepared as previously described) or 1 µg of anti-Clec9A-OVA mAb, both with 1 µg of LPS as adjuvant. Six days later, single-cell suspensions of target cells (splenocytes) were prepared from spleens of wild-type (WT) mice. Half of the suspension (OVA⁺ splenocytes pulsed with OVA_257–264 labeled with a high concentration of CTV [CTV^hi]) was pulsed with 1 µg of OVA_257–264 (Worthington Biochemical) and labeled with 5 mM CTV (Thermo Fisher Scientific). The other half of the suspension (OVA⁻ splenocytes labeled with a low concentration of CTV [CTV^lo]) was labeled with 0.5 mM CTV only. An equal number of OVA⁺ CTV^hi and OVA⁻ CTV^lo target cells were pooled, and 10 × 10⁶ target cells were injected i.v.; 36–42 h later, spleens were harvested and the percentage of OVA-specific lysis was determined as follows:

• R = (% CTV ^lo/ % CTV ^hi)

• % OVA-specific lysis = [1 − (r_unprimed/r_primed)] × 100.

A total of 1 × 10⁴ OT-II × Ly-5.1 or OT-I × Ly-5.1 cells were adoptively transferred into WT and MHC IIKR^KI/KI mice. One day later, mice were immunized with 1 µg of anti-Clec9A-OVA mAb and 1 µg of LPS. Seven days later, spleens were harvested, single-cell suspensions were generated, and RBCs were lysed. Splenocytes were stained with mAbs specific for TCR Vα2 (B20.1; Walter and Eliza Hall Antibody Facility), Ly-5.1 (A20), PD-1 (29F.1A12), CD44 (IM7), and CD62L (MEL-14) (all BioLegend). For IFN-γ analysis, splenocytes were incubated with 1 μg/ml OVA_257–264 in the presence of BD GolgiPlug for 5 h at 37°C. For intracellular staining of IFN-γ and T-bet, splenocytes were permeabilized and fixed with a BD Cytofix/Cytoperm Kit or an eBioscience Foxp3/Transcription Factor Staining Buffer Set. Cells were stained for IFN-γ (XMG1.2) or T-bet (REA102; Miltenyi Biotec).

For T_FH induction, 1 × 10⁶ OT-II × Ly-5.1 cells were adoptively transferred into mice. One day later, mice were immunized with 0.5 µg anti-Clec9A-OVA mAb or PBS (no Ag control). Six days later, spleens were harvested, single-cell suspensions were generated, and RBCs were lysed. Cells were stained with mAbs specific for CD4 (GK1.5; Walter and Eliza Hall Antibody Facility), Ly-5.1 (A20), PD-1 (29F.1A12), CD44 (IM7) (all BioLegend), and CXCR5 (2G8; BD Biosciences). The number of OT-II and T_FH cells was determined as the number of CD4⁺ Ly-5.1⁺ cells or CD4⁺ Ly-5.1⁺CXCR5⁺ PD-1⁺. For anti-rat Ig response, mice were injected i.v. with 2 μg of rat anti-Clec9A mAb (10B4), and serum samples were obtained at 2 wk postinjection. Anti-rat Ig reactivity was determined by ELISA. Nunc MaxiSorp (Thermo Fisher Scientific) plates were coated overnight at 4°C with 1.5 μg/ml rat GL117 mAb (Walter and Eliza Hall Antibody facility). Unbound Ab was removed by washing with PBS plus 0.05% Tween 20, PBS, and Milli-Q water. Serum samples were serially diluted in 5% skim milk powder in PBS and incubated at 4°C overnight. Captured mouse anti-rat Ig Abs were detected using anti-mouse IgG-HRP and visualized using 1-Step Ultra 3,3′,5,5′-Tetramethylbenzidine ELISA Substrate (Thermo Fisher Scientific). Absorbance at 440 nm was detected with a CLARIOstar plate reader (BMG LABTECH). Titers were considered positive when the absorbance was above 0.1 compared with blank.

To generate a large number of DCs, WT and MHC IIKR^KI/KI mice were injected s.c. with 5 × 10⁶ Flt3L-secreting B16 melanoma cells (22). Nine days later, spleens were harvested, and DCs were isolated as previously described (20). cDC1 and cDC2 were further separated by positive selection using MACS magnetic separation (Miltenyi Biotec). Purity >90% was determined by flow cytometry. Cells were lysed in PBS 0.1% Triton X-100 on ice for 20 min. Postnuclear supernatants equivalent to 80 μg of protein were acidified by adding 10× citrate buffer (final concentrations were 50 mM citrate [pH 5.5], 0.5% CHAPS, 0.1% Triton X-100, and 4 mM DTT). Where indicated, lysates were preincubated with the pan-cysteine cathepsin inhibitor E-64 (25 μM; Sigma-Aldrich) or DMSO vehicle at 37°C. A total of 1 μM BMV109, a fluorescently quenched pan-cysteine cathepsin activity-based probe, was added, and lysates were incubated at 37°C. Labeling reactions were stopped by addition of 5× sample buffer (50% glycerol, 200 mM Tris-Cl [pH 6.8], 10% SDS, 0.05% bromophenol blue, 6.25% 2-ME) followed by heating at 95°C for 5 min. Proteins were resolved on a 15% SDS-PAGE gel, and the gel was scanned for Cy5 fluorescence using a Typhoon flatbed laser scanner (GE Healthcare). For cystatin C immunoblots, proteins were transferred to nitrocellulose membranes and probed with goat anti-cystatin C (R&D Systems) and rabbit anti-actin (Sigma-Aldrich) Abs. After washing with PBS plus 0.05% Tween 20, blots were incubated with donkey anti-goat HRP (Novex; Life Technologies) and goat anti-rabbit IRDye 800CW (LI-COR Biosciences). IRDye 800CW immunoblots were scanned using Typhoon, and HRP labeling was visualized on a ChemiDoc MP Imaging System (Bio-Rad Laboratories) with the use of Amersham ECL Western blotting reagents (GE Healthcare). Protease activity and cystatin C (band intensity) were measured by densitometry using ImageJ. Values were normalized to actin (band intensity). Statistical analyses were performed using GraphPad Prism 8.

To investigate the impact of MHC II ubiquitination on DC biology, we used mice harboring a mutant MHC II unable to undergo ubiquitination (MHC IIKR^KI/KI) (12). First, we analyzed the number of splenic DCs. cDC1, cDC2, and pDCs were enriched by DNAse/collagenase digestion, followed by density centrifugation. A reduction in the number of cDC1 (61% decrease), cDC2 (50% decrease), and pDC (30% decrease) was detected in MHC IIKR^KI/KI mice compared with WT mice (Fig. 1A). The decreased cell number was specific for splenic cDC and pDC, with no changes observed for splenic B cells (Fig. 1A) or thymic cDC1 and cDC2 (Fig. 1B).

The remaining cDC in MHC IIKR^KI/KI spleen were profiled for their surface expression of characteristic DC markers (Fig. 1C, Supplemental Fig. 1). As expected, increased MHC II expression was observed on MHC IIKR^KI/KI cDC1 (3.4-fold increase) and cDC2 (2.4-fold increase), whereas no changes were observed in CD86, a well-known MARCH1 substrate (15). Increased MHC II expression was accompanied by decreases in MHC I, CD24, and CD8, which we and others have previously reported as being reduced in the absence of MHC II ubiquitination (11, 15, 18). In addition, we detected previously undescribed, to our knowledge, reductions in CD11b, Sirpα, CD40, CD80, Flt3, XCR1, and DEC205. The only molecule besides MHC II to show increased expression was CD11c, most notably for MHC IIKR^KI/KI cDC1 (Fig. 1C, Supplemental Fig. 1). Similar patterns were observed in splenic B cells, with increased MHC II expression correlating with small but significant decreases in the surface expression of IgM, PD-L1, CD19, and CD40 (Supplemental Fig. 1). These results indicate that impaired MHC II ubiquitination was sufficient to alter a variety of surface molecules, including integrins, chemokine receptors, and C-type lectin receptors.

Given that we observed reduced costimulatory molecule expression (CD40 and CD80) on MHC IIKR^KI/KI DCs, we next aimed to assess other parameters of Ag presentation. For efficient presentation by DCs, Ag must undergo endocytic uptake and proteolytic degradation. First, we measured OVA protein uptake. WT and MHC IIKR^KI/KI cDC were purified and incubated with fluorescent soluble OVA-Cy5. In agreement with previous studies (23), we found cDC1 and cDC2 captured similar amounts of OVA. Similar uptake was observed by MHC IIKR^KI/KI and WT cDC, demonstrating that a lack of MHC II ubiquitination does not alter uptake of OVA (Fig. 2A).

Once internalized, Ag is degraded and loaded onto MHC molecules. To investigate whether Ag proteolysis was impacted by a loss of MHC II ubiquitination, we used DQ-OVA, a self-quenched form of OVA that fluoresces once degraded. MHC IIKR^KI/KI cDC1 and cDC2 were sorted to purity, and equal numbers of DCs were incubated with DQ-OVA. After Ag pulsing, excess DQ-OVA was removed by washing, and assessment of DQ-OVA fluorescence over time, as a read out of OVA proteolysis, was assessed by flow cytometry. In comparison with WT cDC1, MHC IIKR^KI/KI cDC1, and to some extent cDC2 displayed reduced DQ-OVA proteolysis (Fig. 2B). The defect was specific to OVA, given that DQ-BSA, a self-quenched form of BSA underwent similar proteolysis in MHC IIKR^KI/KI DC as in WT DC (Fig. 2C). Impaired proteolysis of DQ-OVA could result from alterations in the endosomal environment in MHC IIKR^KI/KI cDC. To investigate this, we assessed endosomal pH using dextran-pHrodo. This is a pH-sensitive probe for which fluorescence is only detected once the probe enters an acidic environment. Purified WT and MHC IIKR^KI/KI cDC1 and cDC2 were pulsed with dextran-pHrodo and chased for 120 min (Fig. 2D). As expected, fluorescence increased with time, indicative of dextran-pHrodo accessing an acidic environment. No difference was observed between MHC IIKR^KI/KI and WT cDC, indicating no changes to endosomal pH in the absence of MHC II ubiquitination.

To examine protease activity, a fluorescently quenched activity-based probe, BMV109, was used to measure the activity of four cysteine cathepsins, which are key Ag-degrading enzymes (24–26). BMV109 irreversibly binds to cathepsins in an activity-dependent manner, giving rise to Cy5 fluorescence upon release of a QSY21-quenching group. As the binding is covalent, cathepsin binding can be detected by in-gel fluorescence as a surrogate measure of protease activity. A pan-cysteine cathepsin inhibitor, E-64, was first used to confirm the specificity of BMV109 for active papain-fold cysteine proteases. As expected, BMV109 did not bind cathepsins in the presence of E-64, confirming its specificity (Supplemental Fig. 2A). In agreement with previous studies (27), cysteine protease inhibitor cystatin C (CTS3) could only be detected in cDC1 (Supplemental Fig. 2B). Next, cysteine cathepsin protease activity was compared between MHC IIKR^KI/KI DC and WT DC, and no differences were observed (Fig. 2E).

Given that cathepsin activity was normal in MHC IIKR^KI/KI DCs, the reduction in Ag proteolysis together with altered cell surface phenotype suggest that a lack of MHC II ubiquitination could impact endocytic events. Because the production of cytokines is dependent on the endosomal trafficking of TLRs and their recognition of TLR ligands (28), we next investigated the capacity of MHC IIKR^KI/KI DC to produce inflammatory cytokines. Splenic cDC1 and cDC2 were sorted to purity, and equal numbers were incubated in the presence of TLR9 ligand, CpG, and inflammatory cytokines IFN-γ and GM-CSF, conditions that elicit maximal IL-12 production (29). Supernatants were harvested after 18 h, and the presence of IL-6, IL-10, IL-12p70, TNF-α, and MCP-1 were measured using a BD Cytometric Bead Array assay. IL-10 and MCP-1 were not produced by either cDC1 or cDC2. MHC IIKR^KI/KI cDC1 produced increased amounts of IL-6, IL-12, and TNF-α compared with WT cDC1, whereas the cytokine production of MHC IIKR^KI/KI cDC2 was unaltered (Fig. 2F). Similar patterns, albeit at lower concentrations, were observed when DCs were treated with CpG alone (Supplemental Fig. 2C).

In summary, an inability to ubiquitinate MHC II reduces DC numbers, alters their surface phenotype, reduces proteolysis of some forms of Ag, and enhances cytokine production, in particular for the cDC1 subset, but does not impact DC endosomal pH or cysteine cathepsin protease activity.

MHC II expression levels, per se, rather than the extent of its ubiquitination, may regulate endosomal functions. If this were the case, such functions might also be affected in cells deficient in MHC II expression. To address this, we analyzed DCs from IAα^−/− mice. First, the cell surface phenotype was investigated. In contrast to the MHC IIKR^KI/KI DCs, no alterations were observed, except for a slight increase in CD80 (Supplemental Fig. 3A). Next, we investigated the cytokine secretion of IAα^−/− cDC1 and cDC2 (Supplemental Fig. 3B). In agreement with previous studies (30), DCs lacking MHC II produced slightly less IL-6, IL-12, and TNF-α than WT DCs. Finally, we investigated Ag proteolysis using DQ-OVA, which proceeded similarly in WT and IAα^−/− DCs (Supplemental Fig. 3C). Overall, the impact of MHC II deficiency was negligible compared with that caused by lack of MHC II ubiquitination, suggesting that the altered phenotype of MHC IIKR^KI/KI DCs is due to changes in MHC II trafficking rather than to altered MHC II surface expression.

Given the important role of MHC II in B cell and T cell immunity (31) and the dysregulation in Ag processing observed in MHC IIKR^KI/KI DCs, we next investigated the impact of MHC II ubiquitination on adaptive immunity. To investigate Ag presentation in a physiologically relevant setting, WT or MHC IIKR^KI/KI mice were injected i.v. with CTV-labeled OT-II or OT-I. The following day, mice were immunized i.v. with one of three different forms of the model Ag OVA: 1) OVA protein, 2) cell-associated (splenocytes preincubated in vitro with OVA protein), and 3) OVA protein genetically fused to mAb against Clec9A (anti-Clec9A-OVA) (21). Clec9A was chosen as a receptor to measure receptor-mediated Ag uptake, as it is highly expressed by cDC1 (32) and is not altered in DCs with impaired MHC II ubiquitination (Fig. 1C). T cell priming with OCS (33) and anti-Clec9A-OVA (34) is cDC1 dependent. The number of dividing OT-II and OT-I cells in spleen was determined by flow cytometry 3 d following i.v. immunization. For all Ags tested, MHC IIKR^KI/KI mice displayed a significant reduction in the number of dividing OT-II and OT-I cells (Fig. 3A). Therefore, a lack of MHC II ubiquitination impacts both MHC II and MHC I Ag presentation of soluble, cell-associated, and DC-targeted Ag in vivo.

Changes in in vivo T cell priming could be due to intrinsic alterations in DC function and/or the observed reduction in DC numbers (Fig. 1A). To control for the latter, ex vivo Ag-presentation assays were performed using equivalent numbers of isolated WT and MHC IIKR^KI/KI DC. In this case, WT or MHC IIKR^KI/KI mice were immunized with anti-Clec9A-OVA (21); 22 h later, spleens were harvested, and cDC1 were sorted to purity before an equal number of cells were incubated with CTV-labeled OT-II or OT-I cells. Proliferation was determined 4 or 3 d later for OT-II or OT-I, respectively. In agreement with in vivo analysis, reduced OT-II and OT-I priming was observed for MHC IIKR^KI/KI cDC1 (Fig. 3B). Second, we tested the capacity of WT and MHC IIKR^KI/KI cDC1 to present OVA peptide in vitro. WT and MHC IIKR^KI/KI DCs were incubated with OVA_323–339 peptide and IA^b-OVA_323–339–specific OT-II T cells for 3 d. No change in OT-II proliferation was observed in vitro (Supplemental Fig. 4). Previously, we have demonstrated reduced OT-I proliferation when MHC IIKR^KI/KI DC are pulsed with OVA_254–267 peptide (18). Together, these data demonstrate that a lack of MHC II ubiquitination causes DC-intrinsic impairments in Ag presentation.

We investigated whether a lack of MHC II ubiquitination alters generation of effector CTL. WT and MHC IIKR^KI/KI mice were immunized with cell-associated OVA or anti-Clec9A-OVA, with LPS used as an adjuvant. Six days later, mice were injected with target cells that comprised an equal mix of syngeneic splenocytes pulsed with CTV^hi or without CTV^lo.” Robust CTL responses were observed in WT mice immunized with cell-associated OVA (35) or anti-Clec9A OVA mAb (21). In contrast, reduced CTL killing was observed in MHC IIKR^KI/KI mice in response to both immunogens (Fig. 4).

To further investigate T cell responses in MHC IIKR^KI/KI mice, OT-II or OT-I cells were transferred into WT and MHC IIKR^KI/KI mice before immunization with anti-Clec9A-OVA together with LPS as an adjuvant. Six days later, their number and phenotype were examined (Fig. 5). In response to immunization, MHC IIKR^KI/KI mice had reduced numbers of OT-II cells accompanied by reduced PD-1 expression, a marker expressed by activated T cells (36), and a reduction in the percentage of CD62L⁻CD44⁺ OT-II cells. Expression of T-bet, indicative of Th1 differentiation, was unaltered (Fig. 5A). Reduced OT-I numbers were also observed in MHC IIKR^KI/KI mice following immunization. These cells expressed lower PD-1, but did not differ in their expression of CD62L and CD44 or IFN-γ production when restimulated with Ag (Fig. 5B).

Finally, a role for MHC II ubiquitination in humoral immunity was assessed. WT and MHC IIKR^KI/KI mice were immunized with rat anti-mouse Clec9A mAb, which elicits anti-rat Ig reactive Abs (21, 32). Generating germinal centers and Ab responses requires CD4⁺ T_FH cells. To investigate T_FH cell induction, OT-II cells were transferred into WT and MHC IIKR^KI/KI mice, and 1 d later, mice were immunized with anti-Clec9A-OVA mAb. Six days later, we examined production of T_FH, which upregulates PD-1 and CXCR5. WT mice injected with anti-Clec9A-OVA mAb had robust expansion of OT-II cells, with increased PD-1 and CXCR5 expression (21, 37). In contrast, reduced numbers of T_FH cells were detected in MHC IIKR^KI/KI mice (Fig. 6A). The reduction in T_FH cells was associated with a barely detectable Ab response. Although a robust anti-rat Ig response was detected in WT mice, MHC IIKR^KI/KI mice failed to generate serum anti-rat Ig in response to anti-Clec9A (Fig. 6B). Therefore, MHC IIKR^KI/KI mice display significant impairments in humoral immunity.

Ubiquitination of MHC II plays an important role in regulating cell surface MHC II; however, the biological consequences of this posttranslational modification are unclear. In this study we conduct, to our knowledge, the first comprehensive analysis in vivo and show that an absence of MHC II ubiquitination reduces splenic DC numbers, alters DC cytokine production, impairs DC Ag proteolysis, and ultimately reduces in vivo Ag presentation, CTL, and humoral immunity.

Previously, different in vitro Ag-presentation studies have yielded disparate outcomes and have focused on CD4⁺ T cell priming only (9, 14–18). To our knowledge, this is the first study to demonstrate a reduction in MHC II and MHC I Ag presentation in response to soluble, cell-associated and anti-Clec9A mAb-targeted Ag following immunization of MHC IIKR^KI/KI mice. These data are in agreement with a recent study identifying that MHC IIKR^KI/KI mice have reduced CD4⁺ T cell priming following immunization with OVA in the presence of CFA (17). Our study extends this work to different forms of Ag, including soluble, cell-associated, and Clec9A-targeted Ag, demonstrating that MHC II ubiquitination impacts MHC II presentation of Ag acquired via different intracellular routes. In addition, we demonstrated that MHC II ubiquitination impacts MHC I presentation in vivo. Reduced splenic DC numbers may contribute to this impairment, and in addition, ex vivo Ag-presentation assays demonstrate a cell-intrinsic defect in DC function. In agreement with this, MHC IIKR^KI/KI DC isolated from mixed bone marrow chimeric mice generated from MHC IIKR^KI/KI bone marrow have reduced in vitro CD4⁺ T cell priming in comparison with the WT DC from the same mouse (17). Several parameters of DC biology are likely involved, including reduced costimulatory molecule and MHC I cell surface expression, altered cytokine secretion, and altered Ag proteolysis. An altered cell surface phenotype has previously been observed in cells lacking MHC II ubiquitination. Indeed, altered expression of CD80, CD40, CD71, PD-L1, ICOS-L (P. Schriek et al., manuscript posted on bioRxiv, DOI: 2021.04.15.439921), CD4, and CD8 (15) is detected on APCs lacking Marchf1, and MHC IIKR^KI/KI BMDCs have reduced tetraspanin expression (11).

Elevated IL-6, IL-12, and TNF-α secretion described in this study is in contrast to other studies demonstrating that MHC IIKR^KI/KI and Marchf1^−/− DC have reduced IL-12 and TNF-α production (15, 17). This difference is likely due to differences in stimuli tested, with CpG (TLR9 agonist), IFN-γ, and GM-CSF used in this study, in contrast to LPS (TLR4 agonist). This study also shows that MHC IIKR^KI/KI cDC1 have reduced OVA degradation, but overall, MHC IIKR^KI/KI cDC1 or cDC2 do not exhibit a global Ag proteolysis defect. BSA degradation was similar to WT and, likewise, no defects in endosomal acidification or cathepsin activity were detected.

Impairments in DC Ag presentation in the absence of MHC II ubiquitination are likely due to excessive MHC II accumulation on the plasma membrane, which disrupts plasma membrane and lipid raft structure (11). Lipid rafts are important for Ag presentation and if disrupted can lead to reduced APC and T cell conjugate formation, and reduced T cell activation (38, 39). In agreement with this, Marchf1-deficient and MHC IIKR^KI/KI BMDC engage thymocytes at a lower frequency (11), and when pulsed with OVA, Marchf1-deficient and MHC IIKR^KI/KI spleen CD11c⁺ DCs are unable to form stable conjugates with CD4⁺ T cells to the same extent as WT DCs (17). Altered MHC II trafficking in the absence of its ubiquitination has been previously described, with reduced pMHC II degradation (9), MHC II internalization (13, 18), and increased recycling (9). The downstream consequences for T cell priming were examined, and reduced numbers and impaired phenotype of CD4⁺ (OT-II) and CD8⁺ (OT-I) T cells following Clec9A-OVA immunization was identified. This likely accounts for impaired CTL immunity following immunization with cell-associated or Clec9A-targeted Ag and, similarly, poor Ab responses. Indeed, we also observe reduced induction of CD4⁺ T_FH cells in MHC IIKR^KI/KI mice, in accordance with longer TCR:pMHC II interactions being required to promote their differentiation (40, 41).

MHC IIKR^KI/KI mice failed to generate Ab responses to DC-targeted Ag. This is in conflict to previous studies that show normal Ab responses for MHC IIKR^KI/KI mice (14). The discrepancy could stem from the different mouse models, with MHC IIKR^KI/KI mice used in this study as opposed to MHC IIKR-GFP mice, or differences in immunization timing and Ag. Our study immunized with rat anti-Clec9A mAb, which specifically relies on cDC1 presentation, whereas the previous analysis used protein (OVA and hen egg lysozyme) and influenza A virus infection, all of which will engage other DC subsets, including cDC2, that we show in this study are less impacted by a lack of MHC II ubiquitination. Importantly, changes in Ab production could also be due to alterations in B cell biology. Indeed, MHC II is regulated by MARCH1-mediated ubiquitination in B cells. In germinal center B cells, in particular, MHC II expression is dynamically regulated by ubiquitination as the cells cycle between the light and dark zones (42). A lack of MHC II ubiquitination in germinal center B cells leads to prolonged peptide presentation that may impact the presented peptide repertoire and, in turn, delay affinity maturation (42). Therefore, it is possible that this too contributes to the decreased Ab production observed in MHC IIKR^KI/KI mice.

In this study we identify the importance of MHC II ubiquitination in DC, T cell, and B cell responses. Evolutionary conservation of MARCH ligases and the target residue of ubiquitination in MHC II (K225), may therefore have been driven not by a simple requirement to regulate MHC II surface expression but to prevent secondary effects induced by the excessive deposition of MHC II on the cell surface.

The authors have no financial conflicts of interest.