CCR5 Blockade in Combination with Cyclosporine Increased Cardiac Graft Survival and Generated Alternatively Activated Macrophages in Primates Free

Chemotactic cytokines or “chemokines” play a critical role in the activation of innate (1) and adaptive (2) immunity as well as in promoting ischemia/reperfusion injury (3). CCR5 is a major coreceptor for macrophage-tropic HIVs. Its natural ligands include RANTES (CCL5), MIP-1α, and MIP-1β (CCL3 and CCL4), as well as MCP-2 (CCL8). CCR5 is expressed on the surface of monocytes/macrophages, dendritic cells, activated T cells, and NK cells residing in both lymphoid and nonlymphoid tissues (4, 5). Cardiac transplantation is the last resort treatment for patients with end-stage heart failure. Although better immunosuppression has improved the prevention of acute rejection, long-term survival has been marred by the slow progressing graft damage. The predominant obstacle has been cardiac allograft vasculopathy (6).

Our previous studies have indicated that blockade of CCR5 prolonged allograft survival in a fully MHC-mismatched mouse model (7–9), and Schröder et al. (10) have demonstrated that CCR5 blockade modulates inflammation and autoimmunity in primates. Although a recent study demonstrated that CCR5 deficiency favors the generation of alternatively activated macrophages (AAMs), which played an important role in protecting cardiac allograft (11), the mechanism of action is still unknown.

In this study, we investigated the mechanism by which the CCR5 inhibitor, maraviroc (MVC), combined with cyclosporine (CsA) protects cardiac allograft survival in primates. These results coincided with reduced allograft infiltration by Th1, Th17, and macrophages but with increased numbers of Th2 and CD4⁺Foxp3⁺ regulatory T (Treg) cells. Furthermore, the population of macrophages was skewed from classically activated macrophages (CAMs) to AAMs. The AAM bias produced by MVC/CsA combination was dependent on upregulation of peroxisome proliferator-activated receptor γ (PPARγ) nuclear receptor.

MVC (Pfizer Pharmaceuticals) is a compound with specific high-affinity binding to human and rhesus monkey CCR5. Receptor binding studies reported the IC₅₀ value of MVC in the range of 3.3–7.2 nM by inhibition of CCR5 binding to MIP-1α, MIP-1β, and RANTES. Additionally, calcium flux and cAMP levels were inhibited with IC₅₀ at 4–30 nM. The MVC dose of 300 mg delivered orally twice daily exceeded the IC₅₀ level needed to block primate CCR5. A PPARγ-specific antagonist, bisphenol A diglycidyl ether (BADGE), was purchased from Sigma-Aldrich (St. Louis, MO). IL-4 and IL-13 were obtained from Sanofi Synthelabo (Labège, France), and rosiglitazone was from Cayman Chemical. The following Abs were used for immunohistochemistry: mouse anti-human CCR5 (clone eBioT21/8; eBioscience, San Diego, CA), mouse anti-human Arg1 mAb (Hycult Biotech), mouse anti-human Mrc1 mAb (Biocompare, San Francisco, CA), mouse anti-human PPARγ (Santa Cruz Biotechnology, Santa Cruz, CA), mouse anti-human C3d (Biocompare), mouse anti-human CD68 (Dako, Carpinteria, CA), rat anti-human IL-4, RORγt, and mouse anti-human IFN-γ, Foxp3, CD4, CD8 (eBioscience), FITC or CY3-conjugated donkey anti-mouse or anti-rat IgG (Biotium, Hayward, CA).

Outbreed male rhesus monkeys were obtained from the Guangzhou Institute of Zoology of the Chinese Academy of Sciences (Guangzhou, China). Monkeys were housed in the primate facility at the Experimental Animal Center of Tongji Medical College according to the University Research Animal Resources guidelines. Males weighing 3.5–5 kg were selected as organ donors and recipients. The selection of donor–recipient pairs and animals for third-party cells were based (12, 13) on the MHC incompatibility as defined by the MLR response. In particular, a stimulation index >3 assured that each donor–recipient pair was mismatched; pairings were arranged for the MVC/CsA group with a stimulation index of 6.8 ± 2.4; MVC group, 6.2 ± 2.8; CsA group, 5.9 ± 1.2; control group, 7.5 ± 3.9; MVC/CsA plus BADGE group, 8.4 ± 3.5; and BADGE group, 5.6 ± 1.9 (all p > 0.05).

All recipients underwent heterotopic intra-abdominal cardiac allograft transplantation as described previously (14). Graft functions were assessed by ultrasonic wave and electrocardiogram once daily until graft demise. Protocol biopsies were performed whenever an examiner appreciated decreased graft contractility. Graft failure was defined as loss of electrocardiographic, palpable, and visible graft activity. In the main experimental group, recipients (n = 5) were treated with 300 mg MVC delivered by oral gavage twice daily and 5 mg/kg/d CsA injected hypodermically after transplantation until graft was explanted. CsA was dose adjusted to maintain therapeutic levels of 150–350 ng/ml. The cardiac allografts were biopsied on days 9, 30, and 45 and harvested on day 240 posttransplantation. There were three other groups treated with 1) placebo (n = 5), 2) MVC alone (n = 5), and 3) CsA alone (n = 5). Both the cardiac allografts treated with MVC alone and with CsA alone were biopsied on day 9.

To investigate whether PPARγ contributes to the effect of MVC treatment, a PPARγ antagonist, BADGE, was dissolved in DMSO and diluted with PBS and then injected hypodermically to allograft recipients (30 mg/kg) once a day for 30 d beginning 9 d before transplantation (15).

PBMCs obtained from rhesus blood were isolated by a standard Ficoll-Hypaque gradient method. Monocytes were isolated from mononuclear cells by adherence to plastic for 2 h in macrophage serum-free medium designed for the culture of human monocytes and macrophages (Invitrogen) at 37°C in a humidified atmosphere containing 5% CO₂. Nonadherent cells were removed by three washings with PBS. The remaining adherent cells (>85% monocytes) were incubated in serum-free medium.

Cardiac allograft tissues were stained with H&E and elastic van Gieson. Classification of cellular infiltrates was done according to International Society of Heart and Lung Transplantation criteria for acute allograft rejection. For immunohistochemical staining, 5-μm sections of cryostat frozen tissue were applied to poly-l-lysine microscope slides (Sigma-Aldrich) and fixed with cold acetone. Endogenous peroxidase activity was blocked with 5% H₂O₂ in PBS for 30 min. Next, the sections were treated with Dako universal block (0.25% casein in PBS) for 30 min to prevent nonspecific binding of primary Abs and incubated overnight (at 4°C) with the primary Ab specific for Arg1, Mrc1, or PPARγ. Bound primary Abs were detected by a 45-min incubation with a bridging Ab followed by a further incubation for 30 min with 1:20 dilution of avidin peroxidase–antiavidin peroxidase complex (ABC detection system; Vector Laboratories, Burlingame, CA) (16). Slides for immunofluorescence staining with Abs specific for CCR5, CD4, CD8, CD68, IFN-γ, IL-4, ROR-γt, Foxp3, Arg1, Mrc1, or C3d were fixed with acetone and blocked serially with 5% donkey serum, nonfat dry milk and Fc blocker (Accurate Chemical, Westbury, NY). After washing, the slides were incubated with primary Ab, washed again, incubated with donkey anti-mouse or anti-rat IgG conjugated to either FITC (green fluorescence) or CY3 (red fluorescence) and DAPI (Molecular Probes). After washing, slides were viewed under an epifluorescent microscope (DMR; Leica Microsystems, Wetzlar, Germany). Binding specificity was determined by using an isotype-matched Ab. To evaluate cell infiltration, five fields were randomly selected from one section to count the number of positive cells in each field.

Total RNA was isolated from allografts or PBMCs using a Qiagen Mini Kit (Qiagen, Hilden, Germany). Contaminating genomic DNA was removed by digestion with RNase-free DNase (Qiagen). Integrity of isolated RNA was verified by analytical agarose gel electrophoresis. Two micrograms of DNase-treated RNA was used to synthesize the first strand of a cDNA synthesis kit. The primers for PPARγ, Arg1, and Mrc1 were designed using Primer Express software as follows: Arg1 (XM_001103609), forward, 5′-GGACTGGACCCATCTTTCA-3′, reverse, 5′-ATTACCCTCCCGAGCAACT-3′; Mrc1 (NM_001193925), forward, 5′-TCCTGGGTGACGCCCTGCATA-3′, reverse, 5′-TGTTCCCCAGGGTCCACGTCT-3′; PPARγ (NM_001032860), forward, 5′-CTCCTTTGACATCAAGCCCTTC-3′, reverse, 5′-TTTGGTACTCTTGAAGTTTCAGGTC-3′; 18s rRNA (FJ436026), forward, 5′-TGACCACGGGTGACGGGGAAT-3′, reverse, 5′-GTGGGTAATTTGCGCGCCTGC-3′. PPARγ, Arg1, and Mrc1 were amplified in a separate PCR mixture to 18s rRNA, which was used as internal standard. The PCR mixture for PPARγ, Arg1, and Mrc1 contained 1× SYBR Green Master Mix (PE Biosystems), cDNA template, and optimized primer concentrations, made up to a final volume of 25 μl with nuclease-free water. All PCR reactions were performed using an ABI Prism 7900 sequence detection system (Applied Biosystems, Foster City, CA). The ΔΔC_T method was used to quantify the data, and the results were also normalized to nontransplanted cardiac tissue or PBMCs.

Briefly, responder cells were PBMCs from recipients in each group that had undergone cardiac transplantation 7 d earlier. Gamma-irradiated inactivated stimulator PBMCs (8 × 10⁵/ml; from donor or third-party rhesus) were cocultured for 14 h with the same number of responder cells (8 × 10⁵/ml) in complete medium in a humidified 5% CO₂ atmosphere at 37°C for 72 h, followed by pulsing with 0.5 μCi [³H]thymidine (Amersham Biosciences, Cleveland, OH). The cells were harvested with a semiautomatic cell harvester and counted on a beta scintillation counter. The MLR with autologous stimulator cells or with Con A (25 μg/ml; Sigma-Aldrich) substituting stimulator cells were used as negative and positive controls, respectively. An ELISPOT assay was used to analyze levels of IL-4, IL-13, and IFN-γ, and RT-PCR was used to investigate the expression of AAM markers in MLR.

ELISPOT assays were performed as previously described (17). Cells (2 × 10⁷) from a 48-h MLR were placed on 96-well plates that had been previously coated with mouse anti-human IFN-γ, IL-4, or IL-13 (BD Pharmingen) overnight. The cells were incubated for 24 h, washed, and stained with biotinylated mouse anti-human IFN-γ, IL-4, or IL-13. The spots were visualized with 3-amino-9-ethylcarbazole chromogen (Sigma-Aldrich). Visualization and analysis were performed using an ImmunoSpot Series I analyzer (Cellular Technology, Cleveland, OH). All assays were performed in triplicate and were repeated three times.

Blood samples were collected from recipient monkeys of each group; isolated cells were counted after lyses of erythrocytes. The mononuclear cells were cultured in 24-well plates in RPMI 1640 medium supplemented with 10% FBS, 200 ng/ml PMA (Sigma-Aldrich), 400 ng/ml ionomycin, and brefeldin A (Sigma-Aldrich) for 4 h. The cells were harvested and stained with FITC-anti–human CD4 at 4°C for 30 min. After washing with PBS, the cells were fixed, permeabilized, and stained with allophycocyanin-anti–IFN-γ, allophycocyanin-anti–IL-17, or PE-anti–IL-4 (eBioscience) at 4°C for 30 min. The frequency of Treg cells was determined using a nonhuman primate Treg cell flow kit (eBioscience), according to the manufacturer’s instructions. The frequencies of Th1, Th2, and Th17 and Treg cells were analyzed using a FACS cytometer equipped with CellQuest software (BD Pharmingen).

Serum alloantibody was measured by flow cytometry using frozen donor splenocytes as previously reported (18). Briefly, 5 × 10⁵ donor splenocytes were incubated with 50 μl recipient serum at 4°C for 30 min. After washing, cells were incubated with FITC-anti–human IgM or anti-human IgG (BD Pharmingen) at 4°C for 30 min and washed twice with a flow cytometry buffer and analyzed by a FACSCalibur. Results are expressed as the mean fluorescence intensity of the posttransplant minus pretransplant values. The response to autologous splenocytes served as negative controls.

The SDS sample loading buffer was used to lyse the tissues. The Bradford method was used to determine the protein concentration. Prepared protein samples were stored in a −80°C freezer. Twenty micrograms of each protein sample was loaded into each lane. After SDS-PAGE, proteins were transferred to nitrocellulose membranes and stained with Ponceau S. The staining was pictured as the sample loading reference, whereas the Ponceau S color was washed away with double-distilled water. The membrane was incubated with the mouse anti-human PPARγ (Santa Cruz Biotechnology) for 1 h at room temperature. HRP-conjugated goat anti-mouse IgG Ab (Santa Cruz Biotechnology) was added for 30 min. Additionally, all lanes were probed for actin as loading controls and protein was detected using an ECL detection kit (GE Healthcare).

A Kaplan–Meier curve was used to estimate graft survival time, and lesion scores were compared by a Mann–Whitney U test. Other results were analyzed by ANOVA followed by Bonferroni correction. All results were analyzed with SPSS 13.0 software and expressed as mean ± SD. A value of p < 0.05 was considered significant.

We examined the impact of CCR5 blockade on heart allograft survival by using selective inhibitor, MVC (Fig. 1A). After cardiac allograft transplantation, animals were treated with MVC alone or in combination with CsA. Control recipients treated with placebo alone acutely rejected heart allografts (median survival time [MST], 9 d). Continuous treatment with MVC alone or CsA alone resulted in a modest prolongation of allograft survivals (MST, 30 d; Fig. 1A). In contrast, allografts of recipients treated with an MVC/CsA combination showed long-term survival of allografts (MST, >240 d; p < 0.001 as compared with other groups).

These in vivo results were confirmed by histological examination (Fig. 1B, 1C). On day 9 postgrafting, biopsies of allografts in control animals (scored by International Society of Heart and Lung Transplantation method) revealed acute rejection with significant infiltration by mononuclear cells in comparison with biopsies from CsA alone and MVC alone groups (day 30). At the same time, biopsies of allografts performed during an ongoing MVC/CsA therapy showed much less infiltration by mononuclear cells and the lowest International Society of Heart and Lung Transplantation scores (Fig. 1B). Furthermore, the combined therapy protected from damage, as most of the vessels were slightly affected when examined on day 240 after transplantation (Fig. 1C). Thus, addition of selective CCR5 blockade to CsA treatment inhibited infiltration of heart allografts protecting them from damage.

To explain the mechanism by which CCR5 inhibition protected heart allografts from damage, we examined the proliferative response of PBMCs in response to donor versus third-party alloantigens (Fig. 1D). Whereas PBMCs from MVC/CsA and placebo groups had similar responses against third-party stimulators, only PBMCs from the MVC/CsA group showed significantly reduced proliferation to donor stimulators (Fig. 1D). The same PBMCs from recipients treated with MVC plus CsA exhibited much higher levels of IL-4 and IL-13 and lower level of IFN-γ as compared with PBMCs from other groups (Fig. 1E). These experiments indicate that addition of CCR5 inhibitor to CsA dramatically changed the quality of the immune response toward donor alloantigens by promoting a Th2-type response and reducing a Th1-type response.

We also examined the impact of MVC therapy on Ab production in heart allograft recipients. The alloantibody production was measured by the FACS method and immunofluorescence staining. On day 30 posttransplant, recipients treated with CsA alone showed high levels of anti-donor IgM and IgG. In contrast, both anti-donor IgM and IgG were at the low levels in both MVC alone and MVC/CsA groups. However, the levels of anti-donor IgM and IgG became slightly elevated at day 240 posttransplant in the MVC/CsA group (Fig. 2A). The findings observed on day 30 postgrafting correlated with the reduced C3d staining in MVC alone and MVC/CsA groups when compared with heart allografts from the CsA alone group; the C3d staining was also increased on day 240 posttransplant compared with day 30 in the MVC/CsA group (Fig. 2B). These results suggest that CCR5 inhibition modulates not only T cell- but also B cell-mediated immune responses by reducing the production of donor-specific IgM and IgG Abs.

To further understand the impact of MVC therapy on the immune response, we analyzed the levels of different functional CD4⁺ T subsets in peripheral blood of recipients. FACS analysis on days 9 and 30 postgrafting showed that the ratios of CD4⁺IFN-γ⁺/CD4⁺ and CD4⁺IL-17⁺/CD4⁺ T cells were significantly decreased in double therapy group in comparison with other experimental groups. This decline continued since the level of CD4⁺IFN-γ⁺/CD4⁺ T cells was further reduced on day 240. In contrast, the ratio of CD4⁺IL-4⁺/CD4⁺ T cells was markedly increased in the MVC/CsA group, especially on day 240 after transplantation. Interestingly, no significant difference was observed at the ratio of CD4⁺Foxp3⁺/CD4⁺ cells in peripheral blood from MVC/CsA-treated recipients as compared with other groups (Fig. 3). These results indicate that MVC therapy changed the composition of functional CD4⁺ T cells, thus most likely influencing long-term survivals of heart allografts.

We next examined the phenotype and number of graft-infiltrating immune cells. At days 9 and 30 postgrafting, the numbers of graft-infiltrating CCR5⁺, CD4⁺, CD8⁺, and CD68⁺ cells were significantly lower in animals treated with MVC plus CsA than in other groups (Fig. 4). To further investigate the function of MVC, functional T cell subsets were quantified by the immunofluorescence staining. We found that the numbers of CD4⁺IFN-γ⁺ Th1 cells were markedly decreased on days 9 and 30 posttransplant in the MVC/CsA group as compared with other groups. The numbers of Th1 cells continued to decline, as they were significantly reduced on day 240 when compared with those on day 30 postgrafting. At the same time, the numbers of CD4⁺IL-4⁺ Th2 cells were elevated on day 30 posttransplant in the MVC/CsA group when compared with other groups and these numbers still became higher on day 240. Although the numbers of CD4⁺Foxp3⁺ Treg cells seem to be similar at days 9 and 30 postgrafting in all experimental groups, the numbers of CD4⁺Foxp3⁺ Treg cells climbed between days 30 and 240 in the chronic phase in the MVC/CsA group. Another interesting observation was much lower numbers of CD4⁺RORγt⁺ Th17 cells infiltrating heart allografts on day 9 after transplantation in the MVC/CsA group in comparison with numbers in a placebo group (Supplemental Fig. 1A, 1B). Thus, cardiac allografts themselves represented the most dramatic shift in the frequencies of functional Th1, Th2, Th17, and Treg cell subsets.

We next examined how enhanced Th2 polarization may influence allograft survival in MVC/CsA-treated recipients. Previous studies revealed a very high degree of plasticity among macrophages and their quick adjustment in response to changes in the microenvironment. In particular, macrophages display two different phenotypes with different functions, namely CAMs and AAMs (19, 20). It was already demonstrated that Th2 cytokines promote AAM generation (21). To identify AAM macrophages, we measured their presence in cardiac allografts and peripheral blood using Arg1 and Mrc1 markers. Our results showed significant differences of AAM markers on day 30 postgrafting at the graft site and in peripheral blood (Figs. 5, 6). Immunostaining of allografts at day 9 already indicated the increased numbers of Arg1-expressing cells in the MVC/CsA group, whereas the numbers of cells expressing Arg1 and Mrc1 markers became significantly elevated on day 30 postgrafting when compared with all other groups (Fig. 5A, 5B). As observed for Th2 and Treg cells, the levels of Arg1 and Mrc1 markers increased even further on day 240 postgrafting in the MVC/CsA group (Fig. 5A, 5B). Additionally, a double immunofluorescence staining of CCR5 with Arg1 or Mrc1 showed lack of CCR5 expression on AAM (Fig. 5C), suggesting that CCR5 inhibitor has no direct regulatory effect on AAM.

The above results were confirmed by the quantitative PCR method, as heart allografts demonstrated an identical picture of increased Arg1 and Mrk1 mRNA levels in MCV/CsA-treated recipients (Fig. 6, left panels). The same two markers revealed an identical trend in isolated PBMCs (Fig. 6, right panels). Thus, inhibition of CCR5 improves graft survivals in CsA-treated recipients by changing the functional composition of infiltrating grafts: reduced Th1 versus elevated Th2, Treg, and AAM cells.

Recent studies have demonstrated that PPARγ was identified in hematopoietic cells, including dendritic cells and T lymphocytes (22). Furthermore, PPARγ agonists have immunoregulatory functions to inhibit functional maturation and migration of DCs and to induce T cell energy (23–25). More recently, some studies also showed that PPARγ was required for maturation of AAMs and disruption of PPARγ impaired activation of AAMs (26, 27). Therefore, we examined the immunostaining for PPARγ, showing more intense staining of heart allografts in the MCV/CsA group than in the CsA alone or MCV alone group (Fig. 7A). These observations were confirmed by Western blots, as PPARγ protein was expressed at higher levels on days 30 and 240 (but not on day 9) in allografts from the MCV/CsA group (Fig. 7B, right three bands) when compared with allografts from controls on day 9 or from CsA alone or MCV alone on days 9 and 30 (Fig. 7B, first five bands). In fact, the number of PPARγ-expressing cells was increasing with time at the graft site in double MCV/CsA therapy group (Fig. 7C), and this pattern also coincided with an elevated PPARγ mRNA in allograft (Fig. 7D) and in isolated PBMCs (Fig. 7E). Thus, PPARγ expression correlated with the pattern of elevated AAMs in recipients treated with the double therapy, suggesting that CCR5 inhibition upregulated PPARγ and AAM levels.

To document the principal role of PPARγ in protecting the survival of heart allografts, recipients treated with MCV/CsA combination were injected with the PPARγ antagonist, BADGE. The results showed that inhibition of PPARγ prevented long-term survival, as recipients rejected heart allografts at a MST of 45 d (compared with the MVC/CsA group; p < 0.001; Fig. 8A). To understand the mechanism by which BADGE abrogates MVC/CsA-induced long-term graft survival, proliferative response of PBMCs from animals was assessed in an MLR assay in vitro. As we expected, the potent inhibitory effect on cell proliferation induced by in vitro MVC/CsA treatment was abrogated by the additional treatment by PPARγ inhibitor (Fig. 8B). Further exploration correlated the MLR assay with an increased expression of Arg1 and Mrc1 that was induced by the in vitro MVC/CsA treatment and then abrogated by the in vitro BADGE treatment (Fig. 8C). These findings match our results obtained by directly analyzing graft tissues. In particular, grafts from recipients treated with MCV/CsA and examined on day 45 had upregulated PPARγ, Arg1, and Mrc1; effects for all three molecules were reversed by addition of BADGE inhibitor (Fig. 8D). This dramatic reversal also correlated with changes at the graft site: the decreased allograft infiltration by CD4⁺IFN-γ⁺ Th1 cells was reversed by BADGE, whereas the infiltration of CD4⁺IL-4⁺ Th2, CD4⁺Foxp3⁺ Treg, and CD4⁺RORγt⁺ Th17 cells was not affected by BADGE (Supplemental Fig. 1C). Thus, PPARγ inhibitor treatment abrogates the long-term graft survival, inhibits the induction of AAM, and reverses the diminished Th1 response in MVC/CsA-treated recipients.

Finally, we designed the experiments to explain how PPARγ was activated by CCR5 blockade. Recently, Coste et al. (28, 29) have shown that IL-13 activated PPARγ via a PLA2 signaling pathway as well as increased macrophage mannose receptor expression and attenuated gastrointestinal candidiasis. Most importantly, IL-4 and IL-13 were shown to induce the activation of PPARγ regulating macrophage differentiation (30–33). Based on these published results, we studied the role of IL-4 and IL-13 on PPARγ activation in primate monocytes. Our results showed that PPARγ was activated in primate monocytes after 60 min of treatment with IL-4 or IL-13 (Supplemental Fig. 2A). Moreover, we demonstrated that PPARγ was indispensable for the activation of AAM as the upregulated AAM marker genes by IL-4/13 were abrogated with the use of BADGE (Supplemental Fig. 2B). These results suggest that IL-4 and IL-13 may upregulate PPARγ necessary for activation of AAM. However, further experiments are required to confirm whether CCR5 blockade promotes generation of AAM by the Th2 cytokine-dependent induction of PPARγ.

Our current study demonstrated that addition of CCR5 inhibitor (MVC) to CsA therapy induced long-term survival of cardiac allografts in primates. This effect coincided with the lack of graft infiltration by leukocytes and almost intact graft blood vassals, as well as with low numbers of proinflammatory Th1 and Th17 cells as well as increased numbers of Th2 and Treg cells. Finally, the marked upregulation of AAMs in peripheral blood and at the graft site was correlated with the activation of PPARγ, showing a complex interconnected mechanism of the inhibition produced by CCR5 inhibitor.

Studies published by Fairchild and colleagues (34–36) indicated that anti-donor Abs are an important mediator of acute allograft rejection, whereas CCR5 deficiency in recipients elevated the levels of donor-specific Abs during acute rejection of cardiac and renal allografts. In our nonhuman primate cardiac transplant model, inhibition of CCR5 with MVC did not increase the levels of anti-donor Abs and C3d deposition. This was consistent with the recent study by Schröder at el. (10), who reported no impact on anti-donor Abs in cardiac recipients by treatment with CCR5 blocker alone. The potential discrepancy may be explained by the use of different animals, namely rhesus monkeys versus mice. However, our study with the MVC/CsA combination showed a slightly increased donor-specific Ab response on day 240 posttransplant.

The fate of transplanted organs is determined at least partially by the types and numbers of effector T cells (37). It is generally understood that IFN-γ plays an important role in acute and chronic allograft rejection (38, 39). On the one hand, addition of MVC to CsA therapy reduced IFN-γ expression, whereas PPARγ antagonist abrogated this downregulation effect. On the other hand, the expression of Th2-type cytokines IL-4 and IL-13 was increased in the MVC/CsA group. These results suggested that the prolongation of graft survival observed in MVC/CsA-treated recipients may be attributed to the skewed Th1-type response to Th2-type response. The presence of Treg cells was recognized as one of the critical factors in induction of transplant tolerance (40). A recent study performed by Burrell et al. (41) demonstrated that Th17 cells may promote inflammatory response in transplantation. In our experimental study, cardiac allografts from recipients treated with MVC plus CsA were infiltrated with increased numbers of CD4⁺Foxp3⁺ Treg cells observed in the chronic phase and decreased numbers of CD4⁺RORγt⁺ Th17 cells in the acute phase. Indeed, the increased Foxp3 expression was present only at the graft site but not in the blood, a similar pattern reported in previously published studies where trafficking of Treg cells was regulated via chemokine receptors CXCR3, CCR4, and CCR8 (42, 43). Our observations suggest that the interruption of CCR5/CCR5 ligand axis may affect the balance of regulatory and effector T cells at site of allograft.

Cells of the macrophage lineage are a major component of the infiltrate in allografts undergoing T cell-mediated rejection (44). Macrophages are involved in the innate and adaptive immunity during allograft rejection playing a key role in the initiation and effector phases of the immune response (45, 46). Functionally, CAMs seem to be indispensable for an effective immune response against some pathogens known to have an excessive destructive response on tissues (21). In contrast, an AAM subset of macrophages has a well-established role in tissue homeostasis and repair with its potent anti-inflammatory properties (47, 48). The results of a recent study performed by Wang et al. (49) showed that AAMs could exert a dominant protective effect against in vivo renal injury. More importantly, a previous study demonstrated that AAMs inhibited proliferation of CD4⁺ T cells and induced transplantation tolerance (50). Our study showed an upregulation of the AAM numbers in both cardiac allografts and peripheral blood in MVC/CsA-treated recipients. These results suggested that CCR5 blockade reprograms macrophages to the alternatively activated phenotype (AAM), thereby favoring a “graft protecting” status rather than the progressive inflammation and destruction.

The high number of AAMs in allografts of MVC/CsA-treated recipients during the early and late phases raised a question how the CCR5 blockade could possibly mobilize this population. A previously published study strongly suggested that PPARγ was indispensable for maturation of AAM (25, 26). Our experiments found that PPARγ was upregulated in allografts of recipients treated with an MVC/CsA combination. Furthermore, both suppression of cellular proliferation and upregulation of Arg1 and Mrc1 markers induced by MVC plus CsA were all abrogated by inhibition of PPARγ. Based on these results, we concluded that blockade of CCR5 upregulated AAM though activation of PPARγ and this was associated with prolonged cardiac allograft survival.

We also tried to explain how PPARγ was activated by CCR5 blockade. One possible explanation suggested that CCR5 blockade activated PPARγ through the increased production of IL-4/IL-13 by Th2 cells, whereas increased PPARγ elevated the numbers of AAMs. Indeed, our findings confirmed that IL-4/IL-13 induced activation of PPARγ and that IL-4/IL-13 upregulated Arg1 and Mrc1 was abrogated by inhibition of PPARγ. These results were in agreement with the previous study performed by Coste et al. (28), which showed that IL-13-induced AAMs were dependent on the PPARγ signaling pathway. These results connected the CCR5 blockade with an expansion of AAMs, suggesting that similar events in vivo increased AAM levels after MVC therapy.

In summary, our present data demonstrate that treatment with MVC plus CsA generated AAMs and that this generation was dependent on the activation of PPARγ. This finding not only provides important insights into the role of CCR5 chemokine receptor for the activation of leukocyte subpopulation, but it also may be helpful for the development of new therapeutic strategies. We plan to further investigate the precise molecular and cellular mechanism involved in the regulation of the immune response after administration of CCR5 antagonist in combination with CsA.

The authors have no financial conflicts of interest.