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. Author manuscript; available in PMC: 2015 Jun 15.
Published in final edited form as: J Immunol. 2014 May 7;192(12):6092–6101. doi: 10.4049/jimmunol.1302771

Preemptive donor apoptotic cell infusions induce IFN-γ-producing myeloid derived suppressor cells for cardiac allograft protection1

Jane Bryant *,§, Nadine M Lerret *,§, Jiao-jing Wang †, Hee-Kap Kang *, James Tasch *, Zheng Zhang †, Xunrong Luo *,†,
PMCID: PMC4082999  NIHMSID: NIHMS586268  PMID: 24808363

Abstract

We have previously shown that preemptive infusion of apoptotic donor splenocytes treated with the chemical cross-linker ethylcarbodiimide (ECDI-SPs) induces long-term allograft survival in full MHC-mismatched models of allogeneic islet and cardiac transplantation. The role of myeloid derived suppressor cells (MDSCs) in the graft protection provided by ECDI-SPs is unclear. In this study, we demonstrate that infusions of ECDI-SPs increase two populations of CD11b+ cells in the spleen that phenotypically resemble monocytic-like (CD11b+Ly6CHI) and granulocytic-like (CD11b+Gr1HI) MDSCs. Both populations suppress T cell proliferation in vitro, and traffic to the cardiac allografts in vivo to mediate their protection via inhibition of local CD8 T cell accumulation and potentially also via induction and homing of regulatory T cells. Importantly, repeated treatments with ECDI-SPs induce the CD11b+Gr1HI cells to produce a high level of IFN-γ and to exhibit an enhanced responsiveness to IFN-γ by expressing higher levels of downstream effector molecules ido and nos2. Consequently, neutralization of IFN-γ completely abolishes the suppressive capacity of this population. We conclude that donor ECDI-SPs induce the expansion of two populations of MDSCs important for allograft protection mediated in part by intrinsic IFN-γ dependent mechanisms. This form of preemptive donor apoptotic cell infusions has significant potential for the therapeutic manipulation of MDSCs for transplant tolerance induction.

Keywords: Allogeneic cardiac transplantation, ECDI (ethylene carbodiimide), Apoptosis, Myeloid derived suppressor cells, Interferon γ, Indoleamine 2, 3 dioxygenase, Graft rejection, Tolerance

Introduction

In clinical transplantation, life-long immunosuppression is often necessary to control graft rejection but could be associated with life-threatening complications including opportunistic infections, malignancies and cardiovascular diseases (1). Therefore, to establish and maintain donor-specific transplant tolerance is highly desirable and remains an unmet need. Previous work in our lab has demonstrated that infusions of donor splenocytes (SPs) treated with the chemical crosslinker 1-ethyl-3-(3′-dimethylaminopropyl)-carbodiimide (ECDI) induce long-term donor-specific tolerance to islet allografts (2-4), and when combined with a short course of rapamycin or anti-CD20, also to heart allografts and islet xenografts (5), respectively. A first-in-human clinical trial based on the same principle using peptide-coupled autologous cells in patients with multiple sclerosis was recently published and established the clinical feasibility, tolerability, and safety of this novel tolerance strategy (6). Ongoing preclinical studies in nonhuman primate models of allogeneic and xenogeneic transplantation are attempting to establish the efficacy of this tolerance strategy for clinical transplantation. Mechanistically, treatment of splenocytes with ECDI induces their apoptosis and phagocytosis by recipient splenic phagocytes (3, 7), and the subsequent graft protection appears to be dependent on the presence of Gr1+ cells and the enzyme indoleamine 2,3 dioxygenase (IDO)(8). However, the direct effect of donor ECDI-SPs on the Gr1+ cell populations and the source of IDO in this model remain unclear.

Myeloid derived suppressor cells (MDSCs) bearing the cell surface markers CD11b and Gr1 have been described in cancer and inflammation (9). While a barrier to promoting anti-tumour responses, they have been shown to play an important role in establishing allograft tolerance (10, 11). Murine CD11b+Gr1+ cells can be subdivided into monocytic or granulocytic MDSCs according to their surface expression of the components of the Gr1 antigen, Ly6C and Ly6G, and are defined as CD11b+Ly6CHILy6G- and CD11b+Ly6CINTLy6GHI cells respectively (12). A number of conditions including inflammation, infection and tumors can induce MDSCs (reviewed in (13)). They suppress T cell proliferation in both antigen dependent and independent manners through a variety of effector mechanisms including nitric oxide (NO), arginase and reactive oxygen species (ROS) (12, 14, 15); and they promote regulatory T cell (Treg) induction through production of IL-10, TGF-β and IDO (16, 17).

There is also evidence that most MDSC subsets require IFN-γ, both for their induction and effector function, through IFN-γ inducible genes such as inducible nitric oxide synthase (iNOS) (18). Studies in transplantation have demonstrated that IFN-γ can be protective via a variety of mechanisms including mediating the suppressive function of MDSCs (10, 19-21). The potential sources of such IFN-γ are thought to be macrophages, T, NK and/or NKT cells, and can also result from phagocytosis of apoptotic cells in the spleen (22). More recent studies have suggested that immature myeloid cells expressing Gr1 can themselves produce IFN-γ after pathogen infection for host protection (23).

In this article, we show that infusions of apoptotic donor ECDI-SPs induce an increase in two cell populations that resemble monocytic and granulocytic MDSCs. These cells are disinct in their phenotype and function, and protect cardiac allografts in part through IFN-γ production themselves and subseqeunt IFN-γ-mediated pathways of T cell suppression. Our study thus describes a novel therapeutic approach for the manipulation of these suppressor cell populations to facilitate the development of transplant tolerance.

Materials and Methods

Mice

Eight to ten week old male BALB/c (H-2d), C57BL/6 (H-2b) and SJL/J (H-2s) mice were purchased from the Jackson Laboratory and Harlan. All mice were housed under specific pathogen free conditions at Northwestern Univeristy (NU) and protocols were approved by NU IACUC.

Preparation and injection of ECDI-SPs

Donor (BALB/c) splenocytes were treated with ECDI as previously described (2). Briefly, spleens were processed into single cell suspensions and eyrthrocytes were lysed. Splenocytes were incubated with ECDI (Calbiochem, 150mg/ml per 3.2 × 108 cells) on ice for 1 hour followed by washing. 108 cells were injected i.v. into recipient C57BL/6 (B6) mice on day-7 and day +1 with reference to transplantation on day 0.

Heterotopic cardiac transplantation

Abdominal heart transplantation was performed as previously described (24). BALB/c (donor, H-2d) to C57BL/6 (recipient, H-2b) combination was used. In some cases, SJL/J mice (H-2s) were used as donor for third party heart allografts. Briefly, the donor heart was excised en bloc via median sternotomy. The ascending aorta and pulmonary artery of the donor were anastomosed end to side to the recipient abdominal aorta and inferior vena cava, respecitively. Direct abdominal palpation of the heart beating was used to assess graft survival. Rejection is defined by loss of palpable cardiac impulses.

Antibody treatment

In experiments in which cardiac recipient B6 mice were treated with the anti-Gr1 antibody, the antibody (clone RB6-8C5, BioXCell) was injected i.p. at a dose of 200 μg on day -8, followed by 100 μg on days -7, -3, -1, and +1 with reference to transplantation on day 0 (10, 25-27).

Cell isolation and culture

For isolating cells from the spleens and the cardiac allografts, animals were sacrificed, and spleens and cardiac grafts were excised and digested at 37°C with collagnase type IV (Worthington) for 25 minutes, follwed by lysis of erythrocytes. For graft cells, leukocytes were further enriched by density centriguation (Lympholyte, Cedarlane) to remove the myocytes. For functional assays, Gr1+ cells were enriched by negative selection using the following biotinylated antibodies (clones): B220 (RA3-6B2), CD4 (GK1.5), CD8 α (53-6.7), Ter119 (erythroid cells), CD49b (DX5) and CD25 (7D4), and streptavidin-conjugated magnetic beads according to manufacturer's instructions (Miltenyi Biotec). Cell purity was determined by flow cytometry and ranged from 70-90%. For indicated assays, Gr1HI and Ly6CHI cells were further purified by fluorescence activated cell sorting (FACS) to a purity of >98%. All cells cultures were performed in RPMI 1640 (Gibco) supplemented with 10% FCS (Gibco), 1% PenStrep, 1% L-Glutamine (Gibco), 1% HEPES (Lonza), 0.5% Gentamicin (Gibco) and 0.1% β-Mercaptoethanol (Gibco). Splenic CD4+ or CD8+T cells were isolated by negative selection first using the following biotinylated antibodies (clones): CD11b (M1/70), Gr1 (RB6-8C5), Ter119, CD49b (DX5) and B220 (RA3-8B2), followed by positive selection for CD4 or CD8.

Proliferation assays

Splenic CD8+ T cells were purified as described above, labelled with 0.5mM CFSE (Life Technologies) and plated at 1×105 per well as responder cells for in vitro stimulation. T cells were activated by either anti-CD3/28 dynabeads per manufacturer's instructions (Invitrogen), or by 5×105 irradiated donor (BALB/c) APCs. FACS sorted suppressor cells were added at a 1:1 ratio with the CFSE-labeled T responder cells, and incubated at 37°C for 96hours. T cell proliferation was measured by CFSE dilution. For indicated studies, FACS sorted suppressor cells were either pretreated at room temperature for 30 minutes with 10μg/ml anti-IFN-γ (clone XMG1.2, BioXCell) prior to addition to the proliferation assays, or added to the proliferation assays in the presence of 5mM L-NMMA or D-NMMA (Cayman Chemical,) or 2mM 1-Methyl-DL-tryptophan (1-MT) (Sigma-Aldrich) or vehicle (2% carboxymethylcellulose) For suppression assays by graft Gr1HI and Ly6CHI cells, CFSE labeled responder CD8+ T cells were plated at 1×104 per well, co-cultured with 1×104 anti-CD3/28 dynabeads or 5×104 BALB/c APCs, and 1×104 FACS sorted suppressor cells from the graft. T cell proliferation was determined by CFSE dilution after 96 hours.

Flow cytometry

Cells were stained with fluorochrome-conjugated antibodies for 30 minutes on ice, washed, read on the Canto II (BD) and analysed using FlowJo v6.4.7 (TreeStar). For intracellular staining, cells were also fixed and permeabilised after surface staining using cytofix/cytoperm buffers according to manufacturer's instructions (BD Biosciences), and stained with fluorochrome conjugated antibodies for cytokine detection. The following antibodies (clones) were used: Gr1-PE (RB6-8C5), CD11c-APC (HL3) and CD80-FITC (16-10A1), all from BD Biosciences; Ly6C-eFluor450 (HK1.4), CD11b-eFluor780 (M1/70), F4/80-PerCPCy5.5 (BM8), MHCII-PeCy7 (MS/114.15.2), IL-12-PerCPCy5.5 (C17.8), IL-10-FITC (Jes5-16E3), IFN-γ-PeCy7 (XMG1.2), CD4-eFluor450 (GK1.5) and CD8-PerCPCy5.5 (53-6.7), all from eBioscience; Ly6G-PeCy7 (1A8) from Biolegend and CCR2-APC (475301) from R&D Systems. For Annexin V staining, cells were incubated with APC-conjugated Annexin V (1:20, eBioscience) for 10 min at room temperature followed by immediate analysis by flow cytometry.

Protein measurement and cytokine detection

Tissue cytokines were analysed by 32-Plex multiplex assays (Millipore). Tissues were homogenized to obtain cell lysates, centrifuged at 13,000 rpm for 2 minutes, and the soluble portion was collected and analysed by the multiplex assays per manufacturer's instructions. Results were normalized to the amount of total protein as measured by the Bradford assay (Pierce Biotechnology).

Quantitative RT-PCR

Total RNA was extracted using the RNeasy kit (Qiagen) according to manufacturer's instructions. Total RNA was reverse transcribed to cDNA using the High Capacity RNA-to-cDNA kit (Applied Biosystems). RT-PCR amplifications were performed using Taqman Universal Master Mix II and Taqman gene expression assays (Applied Biosystems). The reactions were run at 50°C for 2 minutes, followed by 95°C for 10 minutes and 40 cycles of 95°C for 15 seconds, and 60°C for 1 minute. Reactions were run on the 7500 Real Time PCR System and data analyzed using 7500 v2.0.1. Delta CT values for each duplicate sample were calculated with reference to 18S.

Graft histology and immunohistochemistry

Grafts were snap frozen in OCT compound with liquid nitrogen. All sections were 8 μm thick. Frozen sections were blocked with Avidin/Biotin blocking kit (Vector Laboratories) followed by staining with anti-mouse Foxp3 mAb (1:400, rat IgG2a, κ clone FJK-16s; eBioscience) or anti-mouse CD8 (1:250, rat IgG2a, κ clone 53-6.7, BD Biosciences). Samples were then stained with biotinylated goat anti-rat Ig for Foxp3 (1:200, goat Ig clone polyclonal; BD Biosciences) or biotin-SP-AffiniPure donkey anti-rat Ig for CD8 (1:250, Jackson ImmunoResearch Inc.). Visualization of Foxp3 and CD8 was performed with Vectastain ABC kit (Vector Laboratories) and DAB substrate kit (BD Biosciences).

Statistical Analysis

Significance between groups was calculated by student's t tests, one way ANOVA or the Wilcoxon Rank Sum test as appropriate. A p value of<0.05 was considered significant. All statistical analysis was performed using Prism v5.0 (Graphpad).

Results

Infusions of donor ECDI-SPs expand two cell populations in the spleen that resemble monocytic and granulocytic-like MDSCs

To determine if allogeneic ECDI-SPs infusions could expand cells bearing features of MDSCs, naive B6 mice were injected with either one or two doses of BALB/c ECDI-SPs, and the spleens were harvested for phenotypic analysis for CD11b+Gr1+ cells on the indicated day (Fig. 1A). Clone RB6-8C5 was used to stain for Gr1. As cells expressing the Gr1 antigen can be sub-divided according to their relative expression of Ly6C, a compoenet of the Gr1 antigen (28), clone HK1.4 was used to further stain for Ly6C. As shown in Fig. 1B and 1C, one dose of BALB/c ECDI-SPs induced a significant increase in the percentage as well as the total numbers of the CD11b+Gr1INTLy6CHI (hereafter referred to as “Ly6CHI”) cells and the CD11b+Gr1HILy6CINT (hereafter referred to as “Gr1HI”) cells. The total number of the Gr1HI cells was further increased after an additional dose of BALB/c ECDI-SPs (Fig. 1C). In contrast, injection of syngeneic B6 ECDI-SPs did not significantly increase either population (Fig. 1B and 1C). Phenotypic analysis (Fig. 1D) revealed that the Ly6CHI cells expressed CCR2 and F4/80, a marker profile consistent with that of monocytic-like MDSCs, whereas theGr1HI cells expressed Ly6G (stained with clone 1A8) and exhibited a high side scatter resembling granulocytic-like MDSCs (Fig. 1D). Other markers associated with the MDSC phenotype such as CD124 and CD115 were also measured, but were not expressed by either population (data not shown). Comparison of cells from naïve untreated versus BALB/c ECDI-SPs treated animals revealed that markers of these two populations were not changed by a single injection of BALB/c ECDI-SPs (data not shown) or two injections of BALB/c ECDI-SPs (Fig. 1D). Collectively, these data suggest that treatment with allogeneic ECDI-SPs expands two cell populations in the spleen that resemble monocytic and granulocytic-like MDSCs.

Figure 1. Infusions of ECDI-SPs expand two cell populations in the spleen that resemble monocytic and granulocytic-like MDSCs.

Figure 1

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(A) Timeline of ECDI-SPs injections and spleen harvest for analysis. (B) Top panels: Gating strategy. Bottom panels: Dot plots showing the Gr1INTLy6CHI cells and the Gr1HILy6CINT cells in the spleen from naïve untreated B6 mice (Naïve), B6 mice injected with B6 ECDI-SPs once (×1) or twice (×2), or B6 mice injected with BALB/c ECDI-SPs once (×1) or twice (×2). Dot plots were all gated on CD11b+ cells as shown in the gating strategy. (C) Total numbers of the Ly6CHI and Gr1HI cells in the spleen from naïve untreated mice (Naïve), B6 mice injected with B6 ECDI-SPs once (×1) or twice (×2), or B6 mice injected with BALB/c ECDI-SPs once (×1) or twice (×2).(D) Histograms comparing several markers of the Ly6CHI and Gr1HI cells in naïve untreated mice (Naïve) or mice injected with BALB/c ECDI-SPs twice (×2). The Ly6CHI and Gr1HI gates were identical to those shown in the dot plots in (B). Data for (B) – (D) were obtained from 3-11 animals per group from 3 independent experiments. Significance was determined using t tests, *p<0.05, **p< 0.01.

ECDI-SPs expanded Ly6CHI and Gr1HI cells suppress T cell proliferation

To further characterize ECDI-SPs expanded Ly6CHI and Gr1HI cells and to determine their similarities to MDSC populations, in vitro T cell suppression assays were performed. Here, we primarily focused on CD8+ T cells as they are the dominant effector cells in the graft rejection in our model (8). As shown in Fig. 2A, both subsets significantly inhibited CD8+ T cell proliferation in response to anti-CD3/28 stimulation, with the Gr1HI cells suppressed more efficiently. The un-expanded Ly6CHI and Gr1HI cells from naïve untreated mice were also tested and showed a similar pattern of suppression of CD8+ T cell proliferation (Fig. 2A, right bar graph), albeit slightly less potently than the expanded Ly6CHI and Gr1HI cells from ECDI-SPs treated mice. MDSCs are known to exert their effects in part through cytokine production (29). We next examined the cytokine profiles of the ECDI-SPs expanded Ly6CHI and Gr1HI cells after two infusions of ECDI-SPs. Neither IL-12 nor IL-10 was produced by either population (data not shown). Interestingly, IFN-γ was produced by the Gr1HI cells (Fig. 2B), and they were the predominant source of IFN-γ within the CD11b+ fraction (Fig. 2C). Furthermore, this IFN-γ production was not detected in Gr1HI cells from naïve untreated mice (Fig. 2B and 2D), was expressed at a very low level after one dose of ECDI-SPs, but increased significantly after the second dose of ECDI-SP (Fig. 2D).

Figure 2. ECDI-SPs expanded Ly6CHI and Gr1HI cells suppress CD8+ T cell proliferation.

Figure 2

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(A) Suppression of CD8+ T cell proliferation. 1×105 naïve CFSE-labeled B6 CD8+ T cells were stimulated with anti-CD3/28 dynabeads in the presence or absence of FACS-sorted splenic Ly6CHI or Gr1HI cells from naïve B6 mice or B6 mice treated with BALB/c ECDI-SPs ×2. Representative dot plots were gated on CD8+ cells. “% proliferation” was calculated asCD8+ T cells with diluted CFSE over total CD8+ T cells. Data shown were obtained from 7 independent experiments and statistical significance was determined by paired t tests, *p<0.05, **p<0.01, ***p<0.005. (B) Production of IFN-γ by the splenic Ly6CHI or Gr1HI cells from naïve mice or mice injected with BALB/c ECDI-SPs ×2. Top panels: Gating strategy. Bottom panels: IFN-γhistograms gated on the Ly6CHI or Gr1HI cells as shown from the gating strategy. (C) Production of IFN-γ in the spleen of mice injected with BALB/c ECDI-SPswas restricted to the splenic Gr1HI(Ly6INT) cells. The dot plot on the left was gated on live CD4-CD8- cells as shown in the gating strategy in (B). Data from (B) and (C) are representative of 5-7 mice from 4 independent experiments. (D) The splenic Gr1HIIFN-γ+ cells increased in numbers with repetitive injections of BALB/c ECDI-SPs. Data shown were pooled from 3-4 animals per group, with significance determined by 1 way ANOVA with Bonferroni correction, **p< 0.01.

IFN-γ and IFN-γ inducible genes are critical for the suppressive function of ECDI-SPs expanded Gr1HIIFN-γ+ cells

We have previously demonstrated that pharmacological inhibition of IDO resulted in the loss of protection of cardiac allografts provided by infusions of donor ECDI-SPs, and that depletion of Gr1+ cells in the same model was concomitant with a reduction of IDO+ cells in the cardiac allografts (8). However, the exact source of IDO remained unclear. IDO is known to be an IFN-γ-inducible gene in other cell populations such as dendritic cells (30). Given that ECDI-SPs induce Gr1HI cells to produce IFN-γ, we next sought to determine if IDO could be produced by these Gr1HIIFN-γ+ cells. CD11b+Gr1HI cells from untreated mice or mice treated with ECDI-SPs were FACS sorted to a purity of >98%. Quantitative RT-PCR analysis of this cell population revealed that the Gr1HI cells from untreated animals did not express ido, but they did modestly increase their ido expression after incubation with recombinant IFN-γ (p= 0.008). However, after treatment with two doses of ECDI-SPs, not only was the baseline ido expression significantly higher than that of the control cells, there was a further significant increase after IFN-γ stimulation (p= 0.039) (Fig. 3A). INOS is another IFN-γ-inducible gene and a known effector molecule of MDSCs. As shown in Fig. 3B, in response to IFN-γ stimulation, the Gr1HI cells from ECDI-SPs treated mice showed a significant increase in the expression of nos2, the murine equivalent of inos, (p= 0.033). This increase in response to IFN-γ was not observed in the Gr1HI cells from untreated animals (Fig. 3B). Expression of arginase 1 and heme oxygenase-1, two additional molecules associated with MDSC function, was similarly examined but did not appear to be induced by either treatment with ECDI-SPs or IFN-γ stimulation (data not shown). Collectively, these data indicate that compared with Gr1HI cells from naïve untreated mice, ECDI-SPs expanded Gr1HI cells are induced to produce IFN-γ themselves, and have a further enhanced capacity to up-regulate the expression of IFN-γ-inducible genes. These characteristics may therefore form a positive feedback loop to enhance their suppressive function in vivo in addition to their increase in numbers by the ECDI-SP treatment.

Figure 3. IFN-γ and IFN-γ inducible genes are critical for the suppressive function of ECDI-SPs expanded Gr1HI IFN-γ+ cells.

Figure 3

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(A) Quantitative RT-PCR analysis of ido gene expression by the CD11b+Gr1HI cells. The CD11b+Gr1HI cells were obtained from control untreated mice or mice treated ECDI-SPs ×2. When indicated, cells were further stimulated for 18 hrs with 1 μg/ml recombinant murine IFN-γ. ΔCT values were calculated with reference to 18S and fold increase was determined relative to control. (B) Quantitative RT-PCR analysis of nos2 gene expression by the CD11b+Gr1HI cells. (C) Inhibition of IDO or iNOS partially abrogated suppression of T cell proliferation by the CD11b+Gr1HI cells expanded by ECDI-SPs treatment. CD11b+Gr1HI cells were obtained from mice treated with ECDI-SPs ×2, and suppression assays were set up as in Fig. 2A with the addition of 2mM 1-MT or 5mM L-NMMA or respective controls (CMC as the vehical control for 1-MT and D-NMMA as the control for L-NMMA) as indicated. (D) Neutralization of IFN-γ completely abrogated suppression of T cell proliferation by the CD11b+Gr1HI cells expanded by ECDI-SPs treatment. CD11b+Gr1HI cells were obtained from mice treated with ECDI-SPs × 2, and pretreated with 10μg/ml anti-IFN-γ (or isotype rat IgG1κ) prior to being added to suppression assays set up as above. Data for (A) to (D) were obtained from 2-4 independent experiments with 5-6 mice per group each experiment. Significance was determined by t tests, *p< 0.05, **p< 0.01, ***p< 0.005.

To determine if IDO, iNOS or IFN-γ played a role in the suppression mediated by the Gr1HI expanded by ECDI-SP treatment, we performed similar suppression assays as described in Fig. 2. Addition of pharmacological inhibitors to IDO (1-MT) or iNOS (L-NMMA) to the suppression assays partially reversed Gr1HI cell-mediated suppression (Fig. 3C). Inhibition of arginase 1 did not affect T cell suppression (data not shown), consistent with the lack of its expression as determined by qRT-PCR. Most strikingly, pretreatment of Gr1HI cells with an anti-IFN-γ neutralizing antibody completely abrogated their suppressive function (p< 0.0001) (Fig. 3D). Collectively, these data indicate that ECDI-SPs expanded Gr1HI cells critically depend on the presence of IFN-γ and its downstream effector molecules to exert their suppressive function.

Cardiac allografts from recipients treated with ECDI-SPs show enhanced accumuation of Ly6CHI and Gr1HIIFN-γ+ cells

To determine how the Ly6CHI and Gr1HI cells behave after cardiac transplantation in recipients with or without ECDI-SPs infusions, allogeneic heterotopic cardiac transplants were performed. Spleens and grafts from untreated control or ECDI-SPs treated animals were harvested 7 and 21 days after transplantation. In the spleen (Fig. 4A), Ly6CHI cells significantly increased in percentages and total numbers at day 7 in transplant recipients compared with untransplanted recipients shown in Fig. 1B and Fig.1C. This increase was similar in both untreated control (white bars) and ECDI-SPs treated recipients (black bars), and sustained at day 21. Similarly, Gr1HI cells also significantly increased in percentages and numbers at day 7 compared with untransplanted recipients shown in Fig. 1B and Fig. 1C, but this increase appeared to be more profound in untreated recipeints (white bars) compared with ECDI-SPs treated recipients (black bars) at day 7 (p=0.059); and subsided in both to baseline at day 21 (Fig. 4A). Interestingly, the splenic Gr1HI cells from ECDI-SPs treated recipients were no longer positive for IFN-γ expression (Fig. 4A, right histograms).

Figure 4. Cardiac allografts from recipients treated with ECDI-SPs show enhanced accumuation of Ly6CHI and Gr1HI IFN-γ+ cells.

Figure 4

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(A) Spleen Ly6CHI and Gr1HI cells in control untreated recipients and recipients treated with ECDI-SPs on day 7 (D7) and day 21 (D21) post transplant. Representative dot plots gated on live CD11b+ cells are shown. Gating strategy is similar to that shown in Fig. 1B. Total numbers of spleen Ly6CHI and Gr1HI cells on D7 and D21 are shown in bar graphs. IFN-γhistograms gated on the Gr1HI cells from D7 and D21 spleens are shown on the right. Shaded histogram: isotype control; dashed line: spleen Gr1HI cells from control untreated recipients; solid line: spleen Gr1HI cells from ECDI-SPs treated recipients. (B) Same data as in (A) are shown for cardiac allograft-infiltrating Ly6CHI and Gr1HI cells from control untreated recipients and recipients treated with ECDI-SPs on D7 and D21 post transplant. Representative dot plots gated on live CD11b+ cells are shown. Total numbers of graft-infiltrating Ly6CHI and Gr1HI cells on D7 and D21 are shown in bar graphs. IFN-γhistograms gated on the graft-infiltrating Gr1HI cells from D7 and D21 cardiac allografts are shown. Shaded histogram: isotype control; dashed line: graft-infiltrating Gr1HI cells from control untreated recipients; solid line: graft-infiltrating Gr1HI cells from ECDI-SPs treated recipients. For (A) and (B), 4-6 independent experiments were performed with 4-8 mice per group. Significance was determined by t tests, *p< 0.05, ***p< 0.005. (C) Cardiac allograft cytokine and chemokine levels. Data shown are average of 3 mice per group from 2 independent experiments and significance was determined by 1 way ANOVA with Bonferroni correction, *p< 0.05, **p< 0.01, ***p< 0.005.

In the cardiac allografts however, a very different picture emerged (Fig. 4B): Ly6CHI cells were found in considerably higher percentages and total numbers in the protected grafts from ECDI-SPs treated recipients compared to grafts from untreated recipients at day 7 (Fig. 4B). In contrast, the percentages and total numbers of Gr1HI cells were significantly lower in the protected grafts in ECDI-SPs treated recipients compared to rejecting/rejected grafts from control untreated recipients at day 7. Importantly, the Gr1HI cells from allografts of ECDI-SPs treated recipients continued to express IFN-γ at both day 7 and day 21 (Fig. 4B, right histograms), contrasting to the Gr1HI cells from allografts of untreated recipients which were IFN-γ-. By day 21, the already rejected cardiac grafts from control untreated recipients were progressing to fibrosis, and considerably fewer graft-infiltrating cells, including Ly6CHI cells and Gr1HI cells, were recovered compared to the protected grafts from ECDI-SPs treated recipients.

We next measured cytokine and chemokine production of the graft tissue to determine if there was a correlation between cell infiltration, cytokine/chemokine production and graft outcome. Tissue lysates were prepared from D7 and D21 heart allografts. Multiplex analysis of heart graft tissue lysates revealed that IL-1β and M-CSF, factors important for MDSC induction, were significantly increased in the protected grafts compared to control grafts at day 21. IL-10, a factor known to be produced by MDSCs, was also significantly higher in protected grafts at day 21 compared to control grafts. CCL4, another soluble factor implicated in the recruitment of Tregs, was similarly increased in protected grafts at day 7 and day 21 compared to control grafts. On the other hand, factors associated with neutrophil accumulation including G-CSF, CXCL1 and CXCL2 were significantly hinger in control grafts at day 21 (Fig. 4C).

Collectively, these data indicate that the Gr1HIcells induced by ECDI-SPs infusions and those induced by allogeneic cardiac transplantation may be distinct from each other, and that cardiac allografts from recipients treated with ECDI-SPs show enhanced accumuation of Gr1HIIFN-γ+ cells as well as Ly6CHI cells, and a graft environment consistent with their local induction and accumulation.

Cardiac allograft Ly6CHI and Gr1HI cells produce soluble mediators important for regulatory T cell induction and migration

To determine the functional significance of the Ly6CHI and Gr1HI cells from the protected cardiac allografts in recipients treated with donor ECDI-SPs, we first examined their ability to suppress T cell proliferation. We chose to examine this at day 21 post-transplant, a time point when the fate of the graft in ECDI-SP treated recipients was readily distinguishable from that of the graft in control untreated recipients. As shown in Fig. 5A left panel, neither population isolated from the cardiac allografts on day 21 suppressed in vitro CD8+ T cell proliferation stimulated by anti-CD3/28. MDSCs have been reported to also have the ability to exert antigen-specific suppression of T cell proliferation (12, 31). We next tested if these cells suppressed CD8+ T cell proliferation stimulated by donor APCs. As shown in Fig. 5A right panel, they also did not substantially suppress donor APC stimulated CD8+ T cell proliferation. As a control, in vitro suppression assays using splenic Ly6CHI and Gr1HI cells from the protected recipients on day 21 were also set up and showed similar inhibition of proliferation as observed in Fig. 2A (Supplementary Fig. 1). Interestingly, when we examined cytokine production in ithe supernatants from the suppression cultures by multiplex analysis, we found that co-culture with allograft, but not splenic, Ly6CHI and Gr1HI cellsled to a significantly higher production of IL-10 and CCL4, two soluble mediators implicated in the induction and homing of Tregs (Fig. 5B). Supporting this possibility, cardiac allografts retrieved from donor ECDI-SPs treated recipients showed a progressive increase of the number of Foxp3+ cells compared with those from control untreated recipients (Fig. 5C). This characteristic was unique to graft Ly6CHI and Gr1HI cells, as spleen Ly6CHI and Gr1HI cells from the same recipient mice in the same co-cultures did not lead to an increased production of IL-10 or CCL4 (Fig. 5B). Collectively, these data indicate that post-transplant the graft, but not the splenic, Ly6CHI and Gr1HI cells may function by establishing an environment conducive to local Treg induction and homing, pointing to the functional difference between the graft and the splenic Ly6CHI and Gr1HI cells.

Figure 5. Cardiac allograft Ly6CHI and Gr1HI cells produce soluble mediators important for regulatory T cell generation and migration.

Figure 5

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(A) Graft Ly6CHI and Gr1HI cells harvested on day 21 post transplant from ECDI-SPs treated recipients did not suppress CD8+ T cell proliferation stimulated by either anti-CD3/CD28 or BALB/c APCs. Results are representative of 3-5 individual experiments. Significance was determined by t tests. (B) IL-10 and CCL4 levels in supernatants from T cell proliferation assays (T cells + anti-CD3/CD28) co-cultured with spleen or allograft Ly6CHI or Gr1HI cells harvested on day 21 post transplant from ECDI-SPs treated recipients. Data shown are average of 3 independent experiments. Significance was determined by t tests, *p< 0.05, **p< 0.01. (C) Cardiac grafts from control untreated and donor ECDI-SPs treated recipients were retrieved on the indicated days post transplant, sectioned and stained for Foxp3. Bar graphs show average cell numbers per low power field counted by two different individuals from 12 – 18 different sections from 3-4 different cardiac grafts from each group. Magnification: ×40. **p< 0.01.

Anti-Gr1 antibody depletes both Ly6CHI and Gr1HI cells, concomitant with a significant increase in graft-infiltrating CD8+ T cells and graft loss

To test the in vivo protective role of the Ly6CHI and Gr1HI cells, we treated mice with the Gr1-depleting antibody (clone RB6-8C5). As shown in Fig. 6A, this antibody depleted both Ly6CHI cells and Gr1HI cells in vivo. Here, the Gr1HI cells were shown as SCCHILy6CINT cells (as shown in Fig. 1B and 1D) to avoid the use of the same clone (clone RB6-8C5) for cell surface staining. Given that these cells suppress CD8+ T cells ex vivo (Fig. 2A), we sought to confirm these findings in vivo. As schematically outlined in Fig. 6B, B6 mice were treated with BALB/c ECDI-SPs and anti-Gr1 antibody, and BALB/c cardiac transplants were performed. Consistent with our previous findings (8), in sharp contrast to recipients treated with ECDI-SPs alone, all recipients additionally treated with the anti-Gr1 antibody promptly rejected the cardiac allograft by day 10 (Fig. 6B). Of note, injection of BALB/c ECDI-SPs did not protect third party SJL cardiac allografts (Fig. 6B), consistent with our previous demonstrations of the antigen-specificity of this regimen (2, 8, 32). At day 10, grafts were harvested and graft-infiltrating cells were analyzed by histology and by flow cytometry. It was observed that allografts from recipients treated with ECDI-SP infusions only had preserved graft architecture and overall few graft-infiltrating cells (Fig. 6C, top image), and specifically few CD8+ T cells (Fig. 6D, left image), consistent with graft protection. However, with concomitant anti-Gr1 antibody treatment, there was a discernable loss of graft integrity with a significant increase in overall graft-infiltrating cells (Fig. 6C, bottom image) and specifically graft-infiltrating CD8+ T cells (p= 0.043) (Fig. 6D, right image). These data suggest that the presence of Ly6CHIand Gr1HI cells is critical for the protection of the cardiac allograft, possibly through their control of graft-infiltrating CD8+ T cells.

Figure 6. Anti-Gr1 antibody depletes both Ly6CHI and Gr1HI cells, concomitant with a significant increase in graft-infiltrating CD8+ T cells and graft loss.

Figure 6

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(A) Ly6CHI and Gr1HI cells were both depleted by the anti-Gr1 (clone RB6-8C5) antibody. Mice were injected i.p. with 200μg of anti-Gr1 antibody and the spleen was harvested 24 hours later for analysis. The left panel shows representative dot plots gated on live CD11b+ cells from untreated vs. anti-Gr1 treated mice. Here, the Gr1HI cells were shown as SCCHILy6CINT cells to avoid the use of the same clone (clone RB6-8C5) for cell surface staining. The right panel shows total numbers of Ly6CHI and Gr1HI cells in the spleen from untreated vs. anti-Gr1 treated mice. Data shown are the average of 5 mice per group. (B) Depletion of Ly6CHI and Gr1HI cells by anti-Gr1 antibody abolished cardiac allograft protection provided by ECDI-SPs. BALB/c cardiac allografts were transplanted to B6 recipients treated with BALB/c ECDI-SPs, with or without anti-Gr1 antibody treatment. In one group, SJL cardiac allografts were transplanted to B6 recipients treated with BALB/c ECDI-SPs to demonstrate donor specificity of this regimen. Left panel: timeline of treatment with anti-Gr1 antibody and ECDI-SPs in reference to cardiac allograft transplantation. Right panel: survival curve of the cardiac allografts. (C) Anti-Gr1 antibody treatment resulted in a marked increase of graft-infiltrating cells and a concomitant loss of graft integrity. Cardiac grafts were retrieved from ECDI-SPs alone or ECDI-SPs + anti-Gr1 treated recipients on day 10 post-transplant, and stained by hematoxylin and eosin. Magnification: ×40. Data shown are representative of 4 allografts per group. (D) Anti-Gr1 antibody treatment resulted in a significant increase of graft-infiltrating CD8+ cells around the time of graft rejection (day 10 post transplant). Cardiac allografts were harvested on day 10 post transplant and enumerated for CD8+ T cells by flow cytometry (top panel) or sectioned and stained with anti-CD8 (bottom panel, magnification: ×40). Data shown are representative of or the average of 4-6allografts per group. For (A) and (D), significance was determined by t tests. For (B), significance was determined by Mantel Cox test. *p< 0.05, **p< 0.01.

Discussion

In the current study, we demonstrate that infusions of apoptotic donor cells in the form of ECDI-SPs expand two myeloid suppressor populations (Ly6CHI and Gr1HI) that function to inhibit T cell proliferation. Moreover, treatment with ECDI-SPs induces the Gr1HI population to produce a high level of IFN-γ, and enhances the responsiveness of this population to IFN-γ to produce downstream effector molecules including IDO and iNOS that mediate their ability to suppress T cell proliferation. Finally, these cells traffic to and protect cardiac allografts through inhibiting graft CD8 T cell accumulation, and potentially also through inducing a Treg rich environment.

Phenotypic analysis of Ly6CHI cells expanded in our model revealed similarities with monocytic MDSCs (M-MDSCs), however this population was distinct in its absence of expression of the IL-4 receptor (CD124) and the M-CSF receptor (CD115) (data not shown). Cell subset heterogeneity is a hallmark of MDSCs, with cell surface phenotype dependent on the model, stimulus and anatomic location. M-MDSCs can be further classified as M1 or M2, dependent on the polarizing conditions. M1 M-MDSCs are defined by their production of iNOS, TNFα, and expression of IFNγR, whereas M2 M-MDSCs produce arginase, IL-10 and express CCR2 (33). As shown in Figures 1D, ECDI-SPs expanded M-MDSCs appear to exhibit an M2-like phenocypte by their expressing CCR2. The function of these Ly6CHI cells also appears to correspond to that of M2 M-MDSCs in their ability to suppress CD8 effector T cells and produce factors for Treg induction (Fig. 2A and 5B) (33).

ECDI-SPs-expanded Gr1HI cells are Ly6GHI, Ly6CINT, SSCHI, and CCR2- (Fig. 1B and 1D). This phenotype is indicative of neutrophil-like cells or granulocytic MDSCs (G-MDSCs). Several recent studies have attempted to phenotypically and functionally differentiate between neutrophils, G-MDSCs, and a further population of neutrophil-like cells termed tumor-associated neutrophils (TAN) (34, 35). G-MDSCs can be sub-categorized according to their phenotype, with G2 G-MDSC expressing arginase, IL-10 and CCL2 (33). Graft infiltrating Gr1HI cells in the ECDI-SPs treated cardiac transplant recipients appeared to have an enchanced ability to induce IL-10 production in comparison to their splenic counterpart (Fig. 5B), indicating that the graft environment may skew the Gr1HI phenotype to resemble that of G2 G-MDSCs, which mirrors that of TAN from tumour bearing mice (36). Interestingly, the TAN phenotype has been shown to be dependent on tumor TGF-β, which may also perpetuate Treg development in conjunction with IL-10; and depletion of TAN resulted in a significant increase in local CD8+ T cell proliferation (36). On the other hand, bone marrow neutrophils did not exhibit these characteristics (35, 36). Collectively, our data indicate that ECDI-SPs expanded Gr1HI cells resemble G-MDSC or TAN rather than bone marrow neutrophils, a phenotype that may be further perpetuated by the graft micro-environment.

Flow cytometric analysis of Gr1HI cells revealed that these cells produced a high level of IFN-γ after repeated infusions of ECDI-SPs (Fig. 2C and 2D). Although not yet reported in transplant models, CD11b+Gr1+ cells producing IFN-γ has been previously demonstrated in models of infection. In 2002, two groups demonstrated that the immune response to N. asteroides and Salmonella infections induced CD11b+Gr1+ cells to produce IFN-γ (37, 38). This was corroborated recently in a Streptococcus infection model, where phenotypic analysis demonstrated that these cells were CD11b+Gr1+, and Ly6G+, Ly6CLO, CCR2- and CX3CR1+ (23). Interestingly, this expression profile closely resembles that of ECDI- SPs expanded Gr1HI cells. In each of these studies, the IFN-γ-producing cells were defined either as neutrophils (37, 38) or as immature myeloid cells based on their induced differentiation with addition of growth factors (23). However, only the immature myeloid cells could suppress T cell repsonses (23). Therefore, while more than one subtype of CD11b+Gr1+ cells may be capable of producing IFN-γ, it is clear that not all may be suppressive.

It is interesting to note that our previous attempts to induce transplant tolerance to islet allografts using donor ECDI-SPs were unsuccessful in IFN-γ deficient mice (3), indicating an obligatory role of IFN-γ in mediating tolerance induced by the ECDI-SPs based regime. We have previously shown that IFN-γ contributes to the contraction and deletion of allo-specific T cells (3). Our data here provide an additional role of IFN-γ in graft protection by promoting and perpetuating the function of MDSCs, in part via IDO and iNOS-mediated suppression (12, 39). These findings indicate the complex role of IFN-γ may play in transplantation tolerance, consistent with studies by others demonstrating a protective role of IFN-γ in transplant models via a variety of mechanisms. For example, exogenous IFN-γ prevented GVHD in a murine bone marrow transplantation model through post transplant inhibition of T cell proliferation (19, 20). IFN-γ stimulation of dendritic cells resulted in an upregulation of IDO production which mediated allograft protection in a rat liver transplantation model (21). Most relevant to our findings, a recent study by Garcia et al in a tolerant heart transplant model demonstrated that a monocytic suppressive cell population bearing the phenotype CD11b+Gr1+CD115+ mediated their suppression via signaling responses to IFN-γ, as Ifngr-/- Gr1+ monocytes could not provide graft protection as wild-type Gr1+ cells did (10). Interestingly, the source of the IFN-γ was not the Gr1+ monocyte themselves, as Ifng-/- Gr1+ monocytes were equally efficacious in graft protection as wild-type Gr1+ cells (10). Therefore, to the best of our knowledge, ours is the first report of IFN-γ production by MDSCs themselves potentially mediating the induction of transplant tolerance.

It is unclear at the moment what signals are transmitted to these cells by the injection of ECDI-SPs to allow the induction of IFN-γ in the Gr1HI cells, as these cells themselves do not directly interact and phagocyse the injected ECDI-SPs (Bryant and Luo, unpublished observation). On a broader base, determining the exact mechanism through which ECDI-SPs induce the expansion of both the Ly6CHI and Gr1HI cells is of clinical relevance, as it will potentially allow for a more direct and effective manipulation of this response therapeutically. The pathways through which MDSCs are differentiated and expanded have been extensively studied in other model systems. Inflammatory mediators arising from infections and tumors have been implicated by many studies, including IL-1β, PGE2, COX-2, GM-CSF, IL-6, VEGF, S100A8/9 and IFN-γ (13). Interestingly, the process of apoptotic cell uptake (efferocytosis) also induces PGE2 production (40). Apoptotic ECDI-SPs are phagocytosed in the recipient spleens by subsets of DCs (3), which may generate subsequent mediators associated with MDSC differentiation or expansion. This may provide a link between efferocytosis and the expansion of Ly6CHI and Gr1HI cells in our model. Further studies are ongoing to delineate this relationship.

Previous work in our lab demonstrated that IDO was critical for cardiac allograft tolerance after ECDI-SP infusions (8). In the current study, we demonstrate that this molecule can be produced by the Gr1HI cells and it mediates the T cell suppressive function of the Gr1HI cells. Although the role of IDO in positive allograft outcome has been extensively studied, its production by MDSCs has not been fully investigated. Two recent studies in human tumor MDSCs showed that IDO was directly produced by these MDSCs, and that it played a role in T cell suppression and Treg induction by this cell population (41, 42). Additionally, in allogeneic stem cell transplant recipients, monocytic-MDSCs were also shown to produce IDO, playing a role in T cell suppression (43). Our studies demonstrate that infusions of ECDI-SPs induce an MDSC population with a heightened ability for IDO production in response to IFN-γ, resulting in a positive graft outcome via both mediators through CD8+ T cell suppression and potentially Treg induction. Collectively, ours and published data demonstrating IDO production by MDSCs provide a link between T cell suppression and Treg induction by the same cell population, therefore point to an attractive target for therapeutic exploitation in transplantation.

Characterizing the ECDI-SPs expanded MDSCs before (Fig. 1C) and after (Fig. 4A and 4B) cardiac transplantation suggests that prior to transplantation, these cells are present in the spleen and function to suppress proliferation of T cells via IFN-γ dependent mechanisms; whereas after transplantation, they migrate to the cardiac allograft and protect the graft via possible induction and homing of Tregs. Migration of MDSCs has been best examined in tumors, and is thought to be mediated by multipe tumor-secreted factors, including IL-1 β (44) and M-CSF (17), two factors that we have also observed in our protected cardiac allografts (Fig. 4C). Furthermore, it has been well recognized that functional differences exist between the MDSCs isolated from the spleen and the MDSCs isolated from the site of tumors (45-47). Similarly, migration of MDSCs to transplanted allografts has also be described (10), although there the expression of the ligands for P- and E-selectins on the MDSCs was found to be crucial for the homing. Thus, findings of the spleen vs. the graft MDSCs in our model are consistent with these notions. However, the exact signals necessary for their trafficking to the allograft and the basis for their functional evolution warrant futher investigation.

In summary, we demonstrate that ECDI-SPs expand MDSCs that function to protect transplant allografts paritally through their own production of IFN-γ and its downstream effector molecules. These studies thus expand the recent advances in the understanding of regulatory myeloid cells in transplantation (48), and provide the basis for future therapeutic manipulations of the population size and functionality of MDSCs for the purpose of transplant tolerance induction.

Supplementary Material

1

Acknowledgments

We wish to acknowledge the Northwestern University Interdepartmental ImmunoBiology Flow Cytometry Core Facility for its support of this work.

Footnotes

1

This work was supported by the National Institutes of Health Training Grant T32 DK077662 (N.L.) and NIH Directors New Innovator Award DP2 DK083099 (X.L.).

Authorship: Contribution: J.B., N.L., X.L. designed research; J.B., J.W., N.L., J.T. performed experiments; J.B., N.L., Z.Z. and X.L. analysed results; J.B., N.L. and X.L. wrote the manuscript.

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