To determine if acute rejection resulting from NK cell depletion was caused by B-cell responses, serum levels of alloantibody were measured 10 days after transplantation. P815 (H-2d) mastocytoma target cells ubiquitously express high levels of MHC I and were used to detect anti–H-2d immunoglobulin in recipient serum. Alloantibodies were detected in untreated rejecting control recipients but not in tolerogen-treated recipients given either isotype control mAb or anti-NK1.1 mAb (Fig. 3D). The absence of anti–donor MHC-II–specific antibodies, however, could not be excluded from our assay because the P815 target cells used in our assay lack MHC-II. The number of graft-infiltrating B cells and peripheral B cells were not significantly different between the two groups (Fig. 3E). These findings showed that NK cell depletion did not directly antagonize the tolerizing effect of anti-CD40L mAb treatment on responding B or T cells.
CD11bloCD27hi NK Cells are Enriched in Tolerized Versus Untreated Rejecting Allografts
The absence of TEa T-cell activation or enhanced alloantibody production in response to alloantigen led us to hypothesize that NK cells served a regulatory function that did not directly alter alloantigen-specific lymphocyte responses. Thus, NK cells may favor tolerance by regulating the local inflammatory microenvironment or infiltrating cells in the allograft. Peripheral NK cell subsets and maturation states have been broadly characterized in mice using the markers CD11b and CD27 (27). CD11b and CD27 expression have also been linked to the cytokine-secreting potential of NK cells and their ability to become activated (28). It is not known which of these NK cell subsets migrate to allograft tissues under conditions of rejection or tolerance. Using multiparameter flow cytometry, we found NK cells in the allograft during rejection and tolerization, with more NK cells in grafts from tolerized recipients (Fig. 4A). The distribution of CD11b and CD27 was similar among NK cells in naive mice compared with mice who underwent transplantation receiving DST plus anti-CD40L mAb (Fig. 4B). However, under conditions of untreated rejection, there was a lower frequency of CD11bloCD27hi NK cells and a higher frequency of CD11bhiCD27hi NK cells in the blood and allografts of rejecting recipients compared with tolerant or naive recipients (Fig. 4B,C). The frequency of CD11bhiCD27lo NK cells remained unchanged. To investigate if there were changes in cytokine production between NK cells from tolerized and untreated rejecting recipients, total CD3−NK1.1+ cells were sorted from the spleens and allografts of patients who underwent transplantation 5 days after transplantation, and their cytokine expression measured by reverse-transcriptase polymerase chain reaction (RT-PCR). However, no significant changes in IFN-γ, tumor necrosis factor α, transforming growth factor β, or IL-10 cytokine transcript profiles were observed (Fig. 4D). Global cytokine levels of IL-1β, IL-4, IL-6, IL-10, IL-17, IFN-γ, and tumor necrosis factor α determined by Luminex assay on sera showed no significant differences between tolerized recipients receiving anti-NK1.1 or mIgG2a isotype control at posttransplantation day 10 (not shown). CD27lo and CD27hi NK cell subsets were further sorted from mice who underwent transplantation 5 days after transplantation, but no significant differences in their cytokine profiles were observed in the spleen or allograft under untreated rejecting versus tolerized conditions (not shown). NK cells were fluorescence-activated cell sorter sorted on CD3−NK1.1+ cells based on the expression of CD11b and CD27. CD11bhiCD27lo and CD11bloCD27hi NK cells were tested for their ability to rescue graft survival after total NK cell depletion. Sorted NK cells were adoptively transferred to day 10 transplant recipients who were previously depleted with one dose of anti-NK1.1 at day −1. Both subsets were able to prolong graft rejection 60 days or more (n=4), suggesting that these markers do not uniquely identify regulatory NK cells.
NK Cell–Depleted Recipients Have Increased Monocyte and Macrophage Infiltration
It was possible that NK cells regulated other infiltrating cell populations in the allograft tissue. To study this, we focused on characterizing the graft-infiltrating cells. Immunohistochemical staining of grafts at day 13 revealed that MHC II+ F4/80+ macrophages constituted most graft-infiltrating cells in the NK cell–depleted recipients (Fig. 5A). Immunohistochemical analysis of allograft myocardium showed no significant difference in macrophage infiltration between anti-NK1.1 mAb or isotype control–treated recipients until 10 days after transplantation. A twofold (P<0.005) and a fourfold (P<0.005) relative increase in F4/80+ macrophage number was observed in anti-NK1.1 mAb–treated recipients at 10 and 13 days, respectively (Fig. 5B). NK cell–sufficient allografts contained MHC II+ cells around vessel walls and throughout the myocardium, but only a minority of these cells expressed F4/80, suggesting they were dendritic cells and not macrophages. Posttransplantation day 10 infiltrating F4/80+ cells in NK cell–depleted grafts costained for I-A/I-E, F4/80, and CD86, consistent with the profile of activated macrophages (Fig. 5C). No other significant changes in the percentage of CD11c+dendritic cells, CD11b+Ly6C+ monocytes, or CD11b+Ly6G+ granulocytes could be observed in the allograft after anti-NK1.1 treatment 10 days after transplantation.
NKG2D Blockade Increases Allograft Macrophage Infiltration and Rae-1γ Expression
The absence of alloantibody and CD4 T-cell responses after NK cell depletion suggested that NK cells directly regulate macrophage populations or their monocyte precursors. In addition to triggering effector responses, NK cell–activating receptors, such as NKG2D, have been recently shown to regulate host immune cells including CD8 T cells (10, 29). To determine if NKG2D blockade interfered with tolerance induction, recipients received HMG2D, an anti-NKG2D–blocking antibody, after transplantation. NKG2D blockade was not sufficient to cause acute rejection, but allografts analyzed by flow cytometry 10 days after transplantation contained a higher percentage of F4/80+ macrophages among infiltrating cells compared with recipients receiving isotype control (Fig. 6A,B). Additionally, F4/80+MHC-II+ cells expressed high levels of the NKG2D ligand Rae-1γ. HMG2D treatment further increased expression of Rae-1γ compared with recipients receiving isotype control antibody (Fig. 6C). Short-term adoptive transfer of CFSE-labeled NK cells in HMG2D-treated transplant recipients was performed at day 10 to determine if NK cells actively migrate to the allograft at this time point after transplantation. Twenty-four hours after injection, NK cells were found in the allograft, the spleen, and to a lesser degree, the peripheral lymph nodes (Fig. 6D). These observations suggest that under conditions of tolerance after transplantation, allograft-homing NK cells regulate macrophage infiltration in part by NKG2D-Rae-1γ receptor-ligand interactions.
Our study demonstrated the requirement for NK cells for preventing acute allograft rejection under conditions of costimulatory blockade–induced tolerance. Anti-NK1.1 mAb–treated recipients showed prolonged depletion of NK cells and rejected allografts within 2 weeks after transplantation. These observations are consistent with an islet transplantation study that found NK cells necessary for tolerance induction in anti-CD40L mAb–treated allograft recipients (17). Others have shown NK cells associate with regulating the expansion and activation of CD4+ and CD8+ T-cell responses (24–26, 30, 31). Rejection in our model was associated with increased macrophage infiltration in the graft but no significant alloantibody or CD4+ alloantigen-specific response. Numbers of Tregs were also unchanged between untreated and NK cell–depleted recipients. Despite the absence of robust alloantigen-specific responses, allograft rejection in anti-NK1.1–treated recipients may still be dependent on T cells. Rejection in our model was associated with enhanced macrophage infiltration. Graft-infiltrating monocytes and macrophages secrete proteases and reactive oxygen and nitrogen species, as well as inflammatory cytokines to mediate tissue injury (32). Macrophage depletion using liposomal clodronate attenuated histologic and functional parameters of acute rejection in a renal allograft model (33). One clinical study strongly correlated monocytic infiltration with renal dysfunction, whereas T-cell infiltration did not (34). Renal transplant patients receiving the T-cell–depleting antibody alemtuzumab showed undetectable levels of T cells for the first month after transplantation (35). These patients still experienced acute rejection episodes and allograft dysfunction that correlated with the early infiltration of monocytes and macrophage in the allograft. Thus, unregulated macrophage infiltration into allografts may be sufficient to mediate acute rejection despite attenuated or anergized alloantigen-specific T-cell responses.
NK cells may contribute to transplantation tolerance by directly suppressing or eliminating monocyte or macrophage populations during tolerogenesis. The expression of NK cell–activating receptor ligands in rejecting allografts suggests their role in facilitating acute rejection (36). However, a regulatory role for these ligands could apply during tolerance induction. NK cells regulate host immunity through activating receptors including NKG2D (24, 30). Blockade of NKG2D did not affect tolerance induction in our experimental model and did not prevent the migration of NK cells to the allograft. However, increased infiltration of macrophages was observed compared with untreated tolerized recipients after transplantation. Cytotoxicity-dependent mechanisms may be important in transplantation because Beilke et al. (17) demonstrated that NK cells require intact perforin to establish islet transplantation tolerance. A study of human macrophages showed that lipopolysaccharide stimulation up-regulated the messenger RNA levels of the NKG2D stress ligands ULBP1-3 and MHC class I chain A (37). Increased stimulation increased macrophage susceptibility to NK cell–mediated killing. Similar cytotoxic mechanisms were observed with NK cells co-cultured with immature human dendritic cells (38–40). Our data indicate that allograft-infiltrating macrophages in tolerized recipients expressed high levels of the NKG2D ligand Rae-1γ. Blockade of NKG2D further increased Rae-1γ expression on these cells. Although the contribution of other NKG2D-expressing cells cannot be excluded in our model, we demonstrated that NK cells migrate to the allograft after transplantation. The elevated expression of NKG2D ligand on macrophages may stimulate NK cell activation and promote cytotoxicity during the resolution of inflammation in the tolerized allograft.
Cytotoxic-independent mechanisms of macrophage regulation should also be considered in assessing roles of NK cells in transplantation tolerance. Liver transplant models associating IL-4 treatment to prolonged graft survival noted increased levels of NK cell–derived IFN-γ and indoleamine-pyrrole 2,3-dioxygenase (41). NK cells may also directly compete with the migration of monocytes/macrophages toward chemokines including MIP-1α and inducible protein 10 (42). In contrast, NK cells also have the potential to secrete chemokines such as CCL3/MIP-1α and CCL4/MIP-1β that can recruit other inflammatory cells (13). The expression and contribution of these factors by NK cells under conditions of tolerance requires further study.
Our finding that NK cell subsets expressing CD11b and CD27 differ in the blood and grafts of rejecting versus tolerized mice further highlights the importance of defining functional NK cell subsets. Studies focusing on CD11b and CD27 expression broadly describe NK cell subsets, their maturation, and the expression of cytokines and cytotoxic effector function potential (27, 43). However, NK cell subsets defined by these cell surface markers variably express activating and inhibitory receptors as well as chemokine receptors responsible for homing (5, 43). NK cell effector function mediated by cytotoxic granule release, Fas-FasL binding, or inhibitory co–receptor-ligand interactions such as PD-1-PD-L1 (5) are additional potential mechanisms regulating immunity.
NK cell research has primarily focused on nonself or stress-induced responses in infection and tumor models. However, there is increasing evidence that NK cells act as regulators of adaptive immunity. Uterine NK cells favor IL-10 secretion, lack Fc-receptors, and are poorly cytotoxic. These tissue-resident NK cells protect the developing fetus from maternal immune responses (44). Gut mucosal NK cells regulate local immune responses by expressing IL-22 and limiting responses to intestinal pathogens (45). Similar regulatory roles may be expected in the allograft during rejection or in tolerance. A greater understanding of NK cell subsets and identification of NK cell regulatory mechanisms may uncover tolerance favoring pathways, and therapeutics that may be harnessed to improve transplantation outcomes.
MATERIALS AND METHODS
CD1d-deficient, BALB/c and C57BL/6 mice were purchased from The Jackson Laboratory. C57BL/6 TCR transgenic TEa mice (46) were maintained in our facility. All mice were housed in a specific pathogen-free facility, and all experiments used age-matched and sex-matched mice in accordance with protocols approved by the Mount Sinai Institutional Animal Care and Utilization Committee.
Monoclonal Antibodies and Treatment Protocols
For tolerance induction, the mice received 1×107 DST on day −7 and 250-μg CD40 ligand-specific mAb (MR1; BioXCell, West Lebanon, NH) intravenously on days −7, −4, 0, and +4 relative to transplantation. NK cells were depleted using a single intravenous dose of 100 μg NK1.1-specific mAb (PK136; BioXCell) on day 1. NK cell–sufficient controls received mouse IgG2a isotype control mAb (C1.18.4; BioXCell). NKG2D blockade was achieved using intraperitoneal dose of 200 μg anti-NKG2D–blocking mAb (HMG2D; BioXCell) every 3 days after transplantation.
Vascularized Cardiac Transplantation
BALB/c hearts were transplanted as fully vascularized heterotopic grafts into C57BL/6 mice as described (47). Graft function was monitored every other day by abdominal palpation.
Adoptive Transfer of TCR Transgenic CD4+ TEa Cells
CD4+ T-cell subsets were isolated from the spleens and lymph nodes of TCR transgenic TEa mice using the mouse CD4+ T-cell enrichment kit (STEMCELL Technologies, Vancouver, BC, Canada) according to the manufacturer’s protocol. Cells were then stained with 5-μM CFSE (Invitrogen, Carlsbad, CA), and 2×106 cells were adoptively transferred into C57BL/6 mice on the day of transplantation. CFSE-labeled TEa CD4+ T cells were evaluated day 5 after transplantation by flow cytometry and immunofluorescence microscopy.
Adoptive Transfer of NK Cells
NK cells were isolated from the spleens of C57BL/6 mice using NK cell isolation kit II (Miltenyi Biotec, Cambridge, MA). Isolated cells were then stained with 5-μM CFSE (Invitrogen), and 2×106 cells were adoptively transferred into C57BL/6 mice 10 days after transplantation.
Anti–mouse CD16/32 was used to block Fcγ III/II receptors and fluorochrome-conjugated antibodies specific for CD45, CD3ε, CD4, CD8β, CD25, CD11c, NK1.1, Gr-1, CD19, CD44, CD11b (eBioscience, San Diego, CA), NKp46 (R&D Systems, Minneapolis, MN), and F4/80 (Serotec, Raleigh, NC) were used for staining. Cells were stained according to manufacturers’ protocols. An LSR II (BD Biosciences, San Jose, CA) was used for flow cytometry, and data were analyzed with FlowJo software (Tree Star, Ashland, OR).
P815 (H-2d) mastocytoma cells were blocked with anti-CD16/32 mAb (eBioscience) and incubated with serum diluted 1:100. Samples were then stained with R-phycoerythrin-conjugated secondary antibodies to mouse IgM and IgG (eBioscience). P815 cells incubated with naive serum followed by isotype control R-phycoerythrin conjugated secondary antibody were used to determine background (bg). This control documented cell autofluorescence within the assay. Cells were analyzed by flow cytometry and mean fluorescence index calculated by FlowJo.
Tissues were embedded in optimum cutting temperature compound (Sakura Finetek USA, Torrance, CA) and frozen at −80°C. Five-micrometer sections were prepared and fixed in cold acetone and blocked with 2.5% normal horse serum (Vector Laboratories, Burlingame, CA). I-A/I-E (clone M5/114.15.2), F4/80 (clone CI:A3-1), CD86 (clone GL1), B220 (clone RA3-6B2), CD4 (clone GK1.5), CD8β (clone H35-17.2), and Foxp3 (clone FJK-16S) antibodies were purchased from eBioscience or Serotec. Fluorescein isothiocyanate, Cy3, and Cy5-conjugated antirat secondary antibodies were purchased from Jackson ImmunoResearch (West Grove, PA). Quantification of graft-infiltrating cells was performed by counting 5 fields per tissue section, 3 tissue sections per graft. Tissues were prepared for paraffin embedding after overnight fixation in 4% paraformaldehyde. Five-micrometer paraffin-embedded sections were cleared using xylene, and antigens were retrieved using a 0.05% trypsin incubation at 37°C for 30 min. Sections were blocked with 2.5% normal horse serum (Vector Laboratories) and stained with biotinylated primary antibodies (eBioscience) followed by avidin horseradish peroxidase (eBioscience). Primary and secondary antibodies were stained using a 1:100 dilution. Staining was visualized using NovaRED Peroxidase Substrate Kit and hemotoxylin QS counterstain (Vector Laboratories) according to the manufacturer’s protocols.
Real-Time Polymerase Chain Reaction
Graft tissue was perfused with phosphate-buffered saline then minced and digested with collagenase D (Roche, Indianapolis, IN) according to manufacturer’s protocol. Splenocytes were obtained by pulverization. Single-cell suspensions were red blood cell lysed using ACK Lysing Buffer (Lonza, Walkersville, MD). NK cells, CD4 T cells, CD8 T cells were sorted on a MoFlo XDP Cell Sorter (Beckman Coulter, Miami, FL). Total RNA was extracted from cells using TRIzol reagent (Invitrogen) then used for reverse transcription using OligoDT primers (Invitrogen) and Omniscript RT Kit (QIAGEN, Valencia, CA) according to the manufacturer’s protocol. Quantitative real-time PCR was performed in duplicate using SYBR Green PCR Master Mix (QIAGEN) and LightCycler 2.0 Real-Time PCR System (Roche). Relative expression was calculated as 2(Ct cyclophilin A − Ct gene), where Ct is cycling threshold, with cyclophilin A RNA as the endogenous control.
Serum was harvested from mice day 10 after transplantation. Samples were tested for cytokine levels using a Th17 6-Plex and IL-4 set (Bio-Rad Laboratories, Hercules, CA). Samples were prepared according to manufacturer’s protocol and analyzed on a Bio-Plex 200 system (Bio-Rad). Cytokine concentrations were determined from experimental standard curves with a sensitivity less than 10 pg/mL for all cytokines measured.
Mixed Lymphocyte Reaction
BALB/c stimulator dendritic cells were enriched from total splenocytes using EasySep CD11c+ positive selection kit (STEMCELL Technologies). CD4+ and CD8+-responding T cells were enriched from C57Bl/6 splenocytes and lymph nodes by fluorescence-activated cell sorter sorting. CFSE (Invitrogen)-labeled T cells were cultured with BALB/c CD11c+ dendritic cells at a 1:1 ratio for 5 days in complete medium.
The authors thank Dan Chen, Peter Boros, Jianhua Liu, and Yansui Li (Mount Sinai) for their technical contributions.
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Keywords:© 2012 Lippincott Williams & Wilkins, Inc.
NK cells; Tolerance; Graft-infiltrating cells