Myeloid-derived suppressor cells increase and inhibit donor-reactive T cell responses to graft intestinal epithelium in intestinal transplant patients

2.1 Patients and patient characteristics

Thirty-seven patients who underwent ITx at the Cleveland Clinic Transplant Center from 2009 to 2016 (n = 36) or at the Intestinal Rehabilitation and Transplant Center, University of Pittsburg Medical Center in 2008 and visited the Intensive Care Unit at Cleveland Clinic (n = 1) were enrolled in this study. The clinical study was approved by the regional ethic committee (IRB no. 11-966 and no. 17-471). All participating patients provided informed consent. Additional information is available in Supplementary Materials.

3.1 MDSCs increase in peripheral blood mononuclear cells (PBMCs) of patients after ITx

We first investigated whether myeloid cells expressing the MDSC phenotype were present and increased in PBMCs of patients after ITx without histopathologic evidence of ACR. The flow cytometry gating strategy excluded doublets and dead cells, and MDSCs were identified as lineage⁻HLADR^−/lowCD33⁺CD11b⁺–expressing cells (Figure 1A), consistent with other reports.7, 10, 12-14 Within the MDSC population, 3 subsets were identified based on CD14 and CD15 expression: CD14⁺CD15⁻ monocytic MDSCs (M-MDSCs), CD14⁻CD15⁺ polymorphonuclear MDSCs (PMN-MDSCs), and CD14⁻CD15⁻ early-stage MDSCs (e-MDSCs) (Figure 1A).13 As reported in previous studies,12, 13, 15-17 M-MDSCs had kidney-shaped nuclei and PMN-MDSCs had lobulated nuclei; these MDSCs were relatively smaller than conventional monocytes and granulocytes, respectively (Figure 1B). In contrast, e-MDSCs contained a mixture of cells with small round, lobulated, and kidney-shaped nuclei, and the cell size was smaller than that of the other MDSC subsets (Figure 1B).

Although small numbers of cells expressing the MDSC phenotype were detected in PBMCs before transplant, the total MDSC numbers in PBMCs were significantly increased by 2 months after ITx and maintained past 1 year posttransplant (Figure 1C, D). Although levels of each of the MDSC subset were increased after ITx, they showed different patterns. Peripheral blood PMN-MDSC and e-MDSC numbers were significantly increased from pretransplant levels within 2 months after ITx (Figure 1E, F), and e-MDSC numbers were maintained throughout the study period (Figure 1F). However, M-MDSC numbers were not significantly increased until 2 months after ITx and then increased after 2 months following ITx (Figure 1G). As part of the induction therapy at the time of transplant, 32 (84%) of the 37 patients were treated with alemtuzumab (Campath-1H), a humanized anti-CD52 mAb, known to be a depleting antibody, which has rapid and long-lasting effects.18 Therefore, to investigate whether alemtuzumab affect the numbers of MDSC subsets, we assessed CD52 expression on MDSCs. We consistently observed high CD52 expression on M-MDSCs, low-absent expression on PMN-MDSCs, and heterogeneous expression on e-MDSCs across the patient samples (Figure S1). Consistent with the CD52 expression, M-MDSCs were detected 4 weeks after transplant in peripheral blood of ITx patients not treated with alemtsuzumab; however, M-MDSCs were not detected in those patients treated with alemtuzumab (Figure S2A, B). In serial samples from ITx patients treated with alemtsuzumab, M-MDSCs were detected > 4 months after ITx when other CD52⁺lineage(CD3, CD19, and CD56)⁺ cells were detected but not within a month after ITx (Figure S2C). Overall, these findings suggest that the 3 MDSC subsets increase in the peripheral blood of ITx patients after transplant and that alemtuzumab induction therapy affects the number of M-MDSCs.

Next, the MDSC-phenotype cells isolated from the peripheral blood of ITx patients were tested for the ability to suppress T cell activation. CD33-depleted cells from pretransplant PBMC samples were used as a source of responder T cells and cultured in anti-CD3/CD28 mAb–coated wells, with autologous MDSCs enriched from posttransplant PBMC samples of the same recipient. The MDSCs inhibited both CD4 and CD8 T cell proliferation in response to anti-CD3/CD28 stimulation (Figure 2A) and interferon (IFN)γ production (Table S2). To examine the suppressive activity of each MDSC subset, each subset was sorted on a flow cytometer. Each MDSC subset suppressed T cell proliferation in a dose-dependent manner (Figure 2B).

3.2 Exogenous steroid hormone enhances expansion of MDSCs from human bone marrow cells (BMCs)

Because GM-CSF and proinflammatory cytokines, such as tumor necrosis factor (TNF)-α and IL-6, induce MDSC expansion in tumor rodent models,7 we investigated whether these factors contribute to MDSC accumulation in ITx patients and determined the plasma levels of IL-6, TNF-α, and GM-CSF and their relationship to peripheral blood MDSC numbers. Plasma IL-6 levels, but not those of TNF-α and GM-CSF, were correlated significantly with MDSC numbers in PBMCs (Figure 3A; r = 0.6888, P = .0278 vs 0.3638 and 0.1942, respectively). In addition, we investigated the correlations of MDSC numbers in PBMCs with the days after ITx, tacrolimus trough levels and administered doses of exogenous steroids per body weight, which was normalized to a value based on the potency of each steroid drug (Table S3). MDSC numbers were positively correlated with the exogenous steroid dose (P = .0013) and negatively with tacrolimus trough levels (P < .0001). In contrast to the significant increases in MDSC numbers in peripheral blood before versus after transplantation (Figure 1D), no significant correlation was observed between MDSC numbers and the days after ITx (Figure 3B).

These results led us to hypothesize that serum IL-6 and exogenous steroid induce MDSC expansion. Whereas PBMCs from healthy volunteers cultured with IL-6 and methylprednisolone (MP) did not result in MDSC expansion (Figure S3), IL-6 and MP enhanced the numbers of MDSC-phenotype cells when BMCs from healthy volunteers were cultured in basic medium supplemented with G-CSF and GM-CSF (Figure 3C, Figure S4). MP was more potent than IL-6 in expanding the MDSCs (Figure 3C), and the effect of MP was dose dependent (Figure 3D). Mifepristone, a glucocorticoid receptor antagonist, abrogated the effect of MP (Figure 3E). MDSCs generated from BMCs grown with IL-6 and MP suppressed both autologous CD4 and CD8 T cell proliferation in cultures with allogeneic B cells (Figure 3F, G) and IFNγ production (Figure 3H).

3.3 Positive correlation between MDSCs and T regulatory cells in ITx recipients

In kidney transplant recipients, MDSCs induce FoxP3⁺CD4⁺ T cell proliferation.19 Therefore, we investigated the relation between peripheral blood MDSCs and total regulatory CD3⁺CD4⁺CD25⁺CD127⁻Foxp3⁺ T cell (Treg) numbers (Figure 4A). In this study, we also dissected subsets of Tregs based on Helios expression, which was preferably expressed in naturally occurring regulatory CD4 T cells,20-23 and examined the relations between MDSCs and the numbers of cells in subsets of Treg (Figure 4A). The numbers of total, Helios⁺, and Helios⁻ Treg cells in PBMCs of ITx patients were correlated with MDSC numbers (Figure 4B: r = 0.5584, P = .0056; 4C: r = 0.4622, P = .0263; and 4D: r = 0.6029, P = . 0023, respectively).

3.4 MDSCs increase in the allograft intestinal mucosa

To investigate whether MDSCs infiltrate into transplanted grafts, mononuclear cells from lamina propria components (LPCs) were analyzed by using flow cytometry. MDSC levels were low or undetectable in the allograft mucosa before implantation, but they significantly increased in percentage among CD45⁺ cells after ITx (Figure S5, Figure 5A). For each MDSC subset, the percentages of each of the 3 MDSC subsets increased in the graft biopsies in the first 12-month period after ITx; however, similar to the PBMCs of the recipients, a minimal increase in M-MDSCs within the first 2 months after ITx was evident compared with the graft infiltration of the other 2 subsets (Figure 5B-D).

To investigate potential mechanisms directing MDSCs into transplanted intestinal grafts, comprehensive chemokine RNA levels were assessed on the NanoStrings^® platform. Because distinct MDSC populations were detectable in the intestinal mucosa of grafts after ITx, we focused on chemokines upregulated after ITx. We detected 2 clusters of chemokine RNA expression levels, including CCL25, CCL15, CXCL6, CCL28, CXCL16, and CCL11, upregulated after ITx (Figure S6). Therefore, MDSC subsets in PBMCs were labeled with antibodies to the corresponding chemokine receptors, including CCR1, CCR2, CCR3, CCR7, CCR9, CXCR2, CXCR6, and β7 integrin, and analyzed by using flow cytometry (Figure 5E). M-MDSCs showed strong expression of CCR1 and CCR2 and weak expression of CCR3, e-MDSCs showed weak expression of CCR1 but strong expression of CCR2 and CCR3, and PMN-MDSCs did not express these chemokine receptors. In contrast, PMN-MDSCs and some e-MDSCs showed strong expression of CXCR2, whereas M-MDSCs did not. None of the MDSC subsets expressed CXCR6, CCR7, CCR9, or β7 integrin. Graft expression of the CCR1 and CCR3 ligand CCL15 increased approximately 3-fold at 3 months posttransplant, and this expression level was maintained through 6 months posttransplant (Figure 5F). Expression of the CCR3 ligand CCL11 was low at the time of transplant and at 3 months posttransplant; however, it was markedly increased at 6 months posttransplant (Figure 5F). In contrast, expression levels of other CCR1, CCR2, and CCR3 ligands were as follows: CCL7, CCL16, and CCL26 levels were low in allografts before and after transplant; CCL2, CCL3, and CCL23 levels were low in allografts after transplant; and there was no significant increase in the expression levels of CCL5, CCL8, CCL13, CCL14, CCL24, and CCL28 after transplant (Figure 5F, Figure S6). Graft expression of CXCR2 ligands CXCL1, CXCL2, CXCL3, and IL-8 decreased to near absent levels after transplant when tested at 3 and 6 months posttransplant. However, graft expression of the CXCR2 ligand CXCL6 was low at the time of transplant and increased by > 3-fold by 3 months posttransplant (Figure 5F, Figure S6). Expression of the IFNγ inducible chemokines CXCL9, CXCL10, and CXCL11 that direct recruitment of alloantigen-primed T cells did not increase after the transplant (Figure S6). Overall, these results support a role for CCL11 and CCL15 in directing the recruitment of CCR1- and CCR3-expressing M-MDSCs and e-MDSCs and a role for CXCL6 in directing the recruitment of CXCR2-expressing PMN-MDSCs and e-MDSCs into the intestinal grafts after transplant.

3.5 Reduced peripheral blood MDSCs during ACR

The relationship between peripheral blood MDSC numbers and occurrence of ACR was investigated in samples from 9 ITx patients with biopsy-proven ACR. The total numbers of MDSCs in PBMCs from ITx recipients with ACR was significantly lower than the numbers in patients without ACR (Figure 6A). ACR occurring within the first 2 months after transplant had no significant effect on the numbers of MDSCs in the peripheral blood; however, the numbers of PMN-MDSCs during ACR tended to be lower than the numbers in patients without ACR (Figure 6B). In addition, the numbers of M-MDSCs and e-MDSCs, but not PMN-MDSCs, significantly decreased in PBMCs from ITx patients experiencing ACR 2 months or more after transplant compared with the numbers in patients without ACR (Figure 6B). In terms of clinical characteristics and demographics of the ITx patients, while de novo DSAs were detected at higher rates in patients with ACR than in patients without ACR, there was no significant correlation between patients with ACR and without ACR in the other factors (Table S4).

3.6 MDSCs suppress donor-specific T cell–mediated damage of intestinal epithelial organoids

A critical histologic criterion for the diagnosis of ACR is > 6 apoptotic bodies/10 sequential crypts,24, 25 which is proposed to reflect donor-reactive T cell–mediated epithelial damage.26 To directly investigate the role of MDSCs in ACR, we established an ex vivo coculture system, called “organoid formation assay,” using donor-reactive T cells and intestinal epithelial organoids from donor intestinal grafts. Donor-reactive T cells (CFSE^low T cells) and non–donor-reactive T cells (CFSE^high T cells) were sorted after 6 days of coculture with irradiated donor B cells (Figure 7A). Subsequently, epithelial organoids derived from donor intestinal grafts, were cocultured with CFSE^low T cells or CFSE^high T cells in the presence or absence of the sorted recipient MDSCs for 5 days. The number of viable epithelial organoids was significantly less in the presence of donor-reactive T cells compared with that in the presence of non–donor-reactive T cells (Figure 7B, C), suggesting that the epithelial damage was mediated by donor-specific T cell response. It is noteworthy that the number of viable organoids in the cocultures with MDSCs and donor-reactive T cells was comparable to that of organoids without T cells, suggesting that MDSCs suppress the damage to organoids by donor-reactive T cells (Figure 7B, C). These findings suggest that accumulated MDSCs suppress donor-reactive T cell response to the donor intestinal epithelium.