Control (USP28^+/+) and USP28 knockout (USP28^-/-) mice were obtained as described previously (24). All animals were maintained at the Laboratory Animal Center of the University of Oulu. All experimental procedures were performed in accordance with the license number ESAVI/7374/2019, approved by the National Project Authorization Board of Finland.

DSS was added in the drinking water of 10-12 weeks old USP28^+/+ and USP28^-/- male mice to induce colitis. For the acute DSS colitis model, mice were treated with 2% DSS-water for seven days. DSS-water was replaced by autoclaved water for the following 3 days. Mice were sacrificed on Day 10. For the chronic DSS colitis model, mice received 3 cycles alternating 7 days of 1.5% DSS-water and 14 days of autoclaved water. Blood, spleen, mesenteric lymph nodes (mLN) and colon were collected on the day of the sacrifice for cytokines detection, flow cytometry, and histological staining, respectively.

CD4+ and CD8+ cells were isolated from mouse spleen and lymph nodes respectively using L3T4 microbeads and CD8a (Ly-2) microbeads (Miltenyi). Naïve CD4+ T cells were obtained by CD4+ enrichment (CD4+ T cell isolation kit, Miltenyi) followed by positive isolation of naïve cells (CD62L microbeads, Miltenyi). All T cell isolations were performed according to the manufacturer´s instructions.

Complete RPMI1640 medium containing 10% fetal bovine serum (FBS), 2 mM glutamine, penicillin (100 IU/ml), streptomycin (0.1 mg/ml; Sigma-Aldrich), and 2.5 μM β-mercaptoethanol was used in in vitro cultures expect when mentioned otherwise.

Naïve CD4+ T cells were used for activation, proliferation, and polarization assays.

For surface staining, cells were stained in PBS + 0.5% BSA with the following antibodies: CD3 (560590), CD4 (550954), CD8 (553035), CD44 (561860), CD62L (561919), CD11b (553311), Gr1 (553129), B220 (553089), NK1.1 (553164, all from BD Biosciences, Franklin Lakes, NJ), CD69, CD25, and CD11c (11-0114-82, eBioscience, San Diego, CA) for 40 min at 4°C.

For intracellular staining of cytokines, cells were stimulated in the presence of phorbol 12-myristate 13-acetate (PMA) and ionomycin plus Golgi inhibitor for 4 hours before permeabilization and fixation steps. Foxp3 staining kit (eBioscience) and the following antibodies were used: anti-IL17A (17-7177-81), anti-IFNγ (53-7311-82), anti-Foxp3 (12-4774-42, all from eBioscience) and anti-T-bet (sc-21749, Santa Cruz Biotechnology). Stained cells were analyzed on an LSR Fortessa flow cytometer (BD Biosciences). Data were analyzed using FlowJo software (version 10, Tree Star, Ashland, OR).

RNA was extracted using the RNeasy kit (Qiagen) according to the manufacturer´s instructions. Total RNA was reverse transcribed using the SuperScript VILO cDNA Synthesis Kit (Invitrogen). Quantitative PCR was performed using either Luna universal qPCR Master Mix (Biolabs) for the SYBER Master Mix or PROBE FAST ABI Prism 2X qPCR Master Mix (Kapa Biosystems) for the TaqMan Master Mix. The corresponding primer sequences are listed below in Table 1. Data were analyzed using the Hprt gene (Applied Biosystems) as an endogenous control.

Primary antibodies against mTOR (2983s), Akt (9272), MAPKAPK5 (7419s), Stat3 (9132), p-Stat3 (9145), Stat5 (9363s), p-Stat5 (9359s), Jak1 (3332), p-Jak1 (3331s), Jak2 (3229), and p-Jak2 (3771l) were purchased from Cell Signaling Technology; USP28 (HPA006778), and β-actin (A5441) were from Sigma Aldrich. Results were normalized according to β-actin expression that served as loading control.

Luminex technology [ProcartaPlex Mouse 11-Plex Mix and Match Panel (PPX-11-MXH6CNK); Thermofisher Scientific] was used to measure IFNγ, IL1β, IL10, IL12/IL23p40, IL17A, IL2, IL22, IL4, IL6, MIP-1α and TNFα in mice plasma samples. All quantifications were done according to the protocols provided by the manufacturer.

For colitis experiments, colons were excised, washed with PBS, sectioned and divided into four equal parts: proximal (prox), middle 1 (mid1), middle 2 (mid2) and distal segments. Colon segments were then fixed in 10% neutral buffer formalin for 24h at room temperature before paraffin embedding. Tissues cross sections of 5µm were stained with hematoxylin and eosin and histologic evaluation of colitis severity was performed.

Each colon section was analyzed and scored separately. The degree of inflammation was scored according to the Wirtz protocol. Sections were scored according to the following criteria: 0 = no evidence of inflammation, 1 = low level of inflammation, 2 = moderate level of inflammation, 3 = high level of inflammation, and 4 = maximum inflammation. The overall inflammation score was determined as the sum of the scores for the proximal, middle1, middle2, and distal segments.

Statistical analyses were performed using the Prism 9.0 software (GraphPad Software, La Jolla, CA, USA). P values between groups were calculated using the student t-test. Differences were considered statistically significant at p <0.05.

Very little is known about the function of the deubiquitinating enzyme USP28 in T cell biology so far. First, we determined whether USP28 is expressed in different Th cell subsets and CD8+ cells. Analysis of RNA-seq data from our group (25) showed that USP28 was expressed in CD4+ cells and was preferentially expressed in in vitro differentiated Th17 compared to Th0 (control) and Treg cells ( Figure 1A ). In Treg cells, USP28 expression appeared to be increased at both the mRNA and protein levels compared to Th0 ( Figures 1A, B ). In CD8+ T cells activated under control conditions (Tc0) or differentiated under Tc1 conditions to induce IFNγ production, no difference in USP28 mRNA level was observed when comparing the two conditions ( Supplementary Figure S1A ). However, a trend towards increased USP28 protein expression was detected in CD3/CD28 activated helper T cells at 72h compared to resting naïve T cells ( Figure 1C ), suggesting that USP28 protein levels are induced by T cell activation.

To better understand the role of USP28 in T cells, USP28 knockout (USP28^-/-) and their littermate control mice (USP28^+/+) were generated as previously described (24). First, the knockout of USP28 in T cells was confirmed by Western blot analysis ( Figure 2A ). Characterization of immune cell populations was then performed by flow cytometry analysis in the thymus, spleen, and blood of these mice. Overall, in the thymus, although USP28^-/- mice exhibited a decreased proportion of DN1 cells (CD44+CD25-), no further defect of thymic T-cell development could be observed, as similar proportions of single positive CD4+ and CD8+ cells were quantified in both genotypes ( Supplementary Figure S1B ). In the spleen, the proportions of T cells (CD4+, CD8+ and, CD4+ naïve cells), NK cells and, myeloid cells are similar between USP28^-/- and USP28^+/+ mice ( Supplementary Figure S1C ). Next, the lymphoid cell proportions of B and T cells appear to be altered in the blood of USP28^-/- mice, with a decrease in B cells and an increase in T cells compared to USP28^+/+ mice ( Figure 2B ).

Looking at the T cells population (CD3+ cells), CD8+ cells were significantly increased while CD4+ cells (naïve and helper cell subsets such as Th1, Th17 and Treg cells) remained unchanged in the blood of USP28^-/- mice ( Figure 2B and Supplementary Figure S1C ). Similarly, the proportions of other cell types such as NK cells (NK1.1+), dendritic cells (CD11b+CD11c+), granulocytes (Gr1+), myeloid cells (CD11b+) and myeloid-derived suppressor cells (CD11b+Gr1+) were similar between USP28^-/- and USP28^+/+ mice ( Supplementary Figure S1C ).

Overall, we observed a preferential expression of USP28 in Th17 and Treg cells, and an altered T cell distribution in USP28 knockout mice. These results led us to hypothesize that USP28 may play a role in T cell homeostasis or function.

Since T cells play an essential role in the development of inflammation, to investigate the role of USP28 on T cell effector function, a DSS-induced colitis model was applied to USP28^-/- and littermate control mice. Chronic DSS-induced colitis was induced by three cycles of DSS treatment. Compared to control mice, USP28^-/- mice tended to lose more weight after the first two cycles of DSS feeding. However, at the end of the experiment, no differences in weight loss, spleen weight and colon weight/length ratio were observed between USP28^-/- and USP28^+/+ mice ( Figures 3A, B and Supplementary Figure S2A ). Histological analysis was performed on the colon samples. The overall score was slightly increased in USP28^-/- mice compared to control mice ( Figure 3C ), and an increase in immune cell infiltration was particularly observed in the mid2 segment ( Figure 3D ). Furthermore, a disruption of mucosal structure in the mid2 segment can be observed in USP28^-/- DSS-challenged mice compared to USP28^+/+ DSS-challenged mice ( Figure 3E ). Characterization of immune cells in spleen and mLN did not show any difference between the two groups ( Supplementary Figure S2B ).

Because greater weight loss was observed in USP28^-/- mice after the first cycle of DSS treatment, we decided to evaluate the effect of USP28 in an acute DSS-induced colitis model. Throughout the acute DSS treatment, USP28^-/- mice lost significantly more weight compared to USP28^+/+ control mice ( Figure 3F ). Reduced colon length and a trend towards increased inflammatory scoring of whole colon samples were consistently observed between the two groups ( Figures 3G, H ). Flow cytometric analyses of immune cells in the mLN showed changes in the CD8+ cell subset in both spleen and mLN. USP28 deficiency resulted in a reduced proportion of CD8+ cells compared to control mice, along with a shift in the proportion of central memory, effector and naïve CD8+ cells. This shift was characterized by a decrease in memory and naive CD8+ cells in favor of effector CD8+ cells ( Figures 3I, J and Supplementary Figures S3A, B ). Notably, a trend toward increased frequency of IFNγ+ cells was observed in CD4- cells, possibly from CD8+ cells. No differences were observed between the two groups for B cells, NK cells, myeloid cells, T cells or CD4+ T cell subsets ( Supplementary Figures S3C, D ).

Together, USP28 protects mice against early DSS-induced colitis development and long-term intestinal structural integrity.

We then characterized the cytokine profile in the peripheral blood of USP28 deficient and littermate control mice from the colitis experiments. In non-challenged mice, IL22 levels stood out. They were significantly higher in USP28 deficient mice compared to control mice, whereas no significant differences in other cytokine levels were observed between the two groups. In both acute and chronic DSS-induced colitis settings, cytokine levels were comparable between USP28^-/- and USP28^+/+ mice and the level of IL22 was still increased in USP28^-/- mice compared to control mice ( Table 2 and Figure 4A ). To further elucidate the alterations in IL22, we conducted mRNA quantification of IL22 along with IFNγ and IL2, known to be necessary for optimal IL22 production (26) in mesenteric lymph node (mLN) samples from the acute DSS-challenged mice. The results revealed a significant upregulation in the mRNA expressions of both IL22 and IFNγ in USP28^-/- mice ( Figure 4B ). Together, this suggests a potential regulatory role of USP28 in modulating IL22 and IFNγ expression during acute DSS-induced colitis, with implications for the involvement of the IL2 pathway in this context.

IL22 is known to activate the JAK/STAT and MAPK pathways (27). IL2 and its downstream signaling play an important role in T cell activation and proliferation. Therefore, we decided to investigate the role of USP28 on T cell activation and proliferation in vitro by using USP28^-/- mice compared to control mice. For the T cell activation assay, naïve CD4+ cells were activated with elevated concentrations of anti-CD3 and anti-CD28 for 24h and 48h. T cell activation markers, CD69 for early activation and CD25 for mid-late activation, were then analyzed by flow cytometry ( Figure 5A ). At 24h post activation, cells only activated with anti-CD3 showed no differences in the proportion of CD69+ cells between USP28^-/- and USP28^+/+ T cells. However, when the cells were activated with different concentrations of both anti-CD3 and anti-CD28, a significantly reduced frequency of CD69 expression was observed in USP28^-/- T cells compared to control T cells ( Figure 5A ). This effect was lost after 48h of activation ( Supplementary Figure S4A ). In response to activation, the cytokine IL2 is produced by T cells before binding to its own receptor (IL2R) on the surface of the T cells, inducing a positive feedback loop that promotes the IL2 signaling pathway (28). Therefore, the expression of CD25, the alpha subunit of the IL2R, was analyzed by flow cytometry. CD25 expression was observed in T cells at 24h, 48h and 72h after activation. Our data show that CD25 expression is significantly lower in USP28^-/- T cells at 24h post activation compared to USP28^+/+ cells. The reduced CD25 and CD69 expression in USP28^-/- T cells were observed at an even earlier time point ( Supplementary Figure S4B ). However, with prolonged activation, CD25 expression became similar in both USP28^-/- and USP28^+/+ cells ( Figure 5B ).

Next, we examined in vitro T cell proliferation in USP28^+/+ and USP28^-/- helper T cells after 3 and 4 days of activation. The distribution of proliferative T cells for each cell division is altered in USP28^-/- T cells ( Figure 5C ). Our results show a decrease in the percentage of proliferative cells in the undivided and early stage of division whereas there is an increase in the percentage of proliferative cells in the late stage of division in USP28^-/- T cells ( Figure 5C ).

Collectively, our results suggest that USP28^-/- T cells have an early defect in activation that may be mediated by IL2R/CD28 signaling. However, at a later stage an increase in their proliferation rate was observed upon TCR activation compared to USP28^+/+ control T cells. Overall, USP28 appears to alter the early phase of T cell activation and proliferation.

We then investigated the effect of USP28 on in vitro T cell differentiation and effector functions. First, CD8+ cells were isolated and cultured in vitro under anti-CD3/CD28 activation conditions (Tc0) or Th1 like inflammatory conditions (Tc1). Flow cytometric analysis of IFNγ+ and T-bet+ cells as well as mRNA levels of these genes in USP28^+/+ and USP28^-/- cells under either Tc0 or Tc1 condition did not show any differences between these two groups ( Supplementary Figures S5A, B ). However, the mRNA levels of granzyme B and perforin in Tc1 condition seemed to be increased in USP28^-/- CD8 cells compared to their littermate controls ( Supplementary Figure S5B ).

Next, we investigated the role of USP28 in the differentiation of CD4+ cells into Th17 or Treg cells and analyzed the Treg suppressive function. USP28^-/- and USP28^+/+ naïve CD4+ T cells were isolated and cultured in different differentiation media for 3 days before flow cytometric analysis of Foxp3 and IL17 expression in these cells. No significant differences were observed in the proportion of Foxp3+ cells in Treg differentiation between USP28^-/- and USP28^+/+ cells ( Figure 6A ). Since other members of the USP family, USP7 and USP44 have been shown to be involved in Treg suppressive function (11, 12), we then performed an in vitro Treg suppression assay using Treg from USP28^+/+ and USP28^-/- mice. The ability of Treg cells to inhibit the proliferation of responder T cells was assessed by Celltrace labelling and flow cytometry analysis. A decrease in the proliferation of responder T cells was observed when the cells were co-cultured with USP28^-/- Treg cells compared to control Treg cells ( Figure 6B ). On the Th17 side, we observed a consistently and significantly reduced proportion of IL17+ cells in USP28^-/- vs USP28^+/+ cells under Th17 polarizing conditions at both protein and mRNA level ( Figures 6C, D ). Surprisingly, we also detected a significantly increased expression of IL22 at the mRNA level ( Figure 6D ).

Taken together, our results indicate that USP28 is not required for CD4+ Treg differentiation and Tc1 differentiation. However, USP28 is involved in Th17 cell differentiation, and more importantly, it contributes to Treg cell suppressive function.

Given our observations of decreased CD25 expression at early stages of T cell activation, but increased T cell proliferative capacity at later stages of activation, and increased IL22 expression in polarized Th17 cells, our aim was to investigate the underlying mechanisms. Given the established links between the CD28 costimulatory, IL2-activated JAK/STAT, and PI3K/AKT/mTOR pathways (28–33), we analyzed the expression of proteins related to the these pathways in activated naive CD4+ T cells ( Figure 7A and Supplementary Figure S6A ). The relative protein expressions of mTOR, AKT, MAPKAPK5, STAT3, p-STAT3, STAT5, JAK1, JAK2 and p-JAK2 were similar between USP28^-/- and control T cells, whereas we observed an increase of p-JAK1 and p-STAT5 expression in USP28^-/- T cells compared to control T cells ( Figure 7B ). Our results indicate that USP28 deficiency in T cells leads to an increase in the activation of JAK1/STAT5 signaling pathways. Next, we analyzed the kinetic activity of the STAT5 pathway in response to IL2 stimulation using Western blot analysis ( Figure 7C ). In response to IL2 stimulation, USP28^-/- T cells showed an increase in STAT5 phosphorylation after 30min and 6h, but not after 24h of stimulation compared to control T cells ( Figure 7D ). Furthermore, the increased STAT5 phosphorylation was also observed in USP28^-/- T cell-depleted splenocytes in response to IL7 ( Supplementary Figure S6B ). Together, these results suggest that USP28 regulates the activity of the STAT5 pathway by affecting STAT5 phosphorylation.