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,
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,,,
Departments of
*
Biomedical Engineering and
Immunology, and
The Glickman Urological Institute, Cleveland Clinic Foundation, Cleveland, OH 44195;
Department of Pathology, Case Western Reserve University School of Medicine, Cleveland, OH 44106;
¶ Department of Medicine, University of Pennsylvania, Philadelphia, PA 19104; and
|| Transplant Research Center, Brigham & Women’s Hospital and Children’s Hospital Boston, Harvard Medical School, Boston, MA 02115
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Abstract |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionDisclosuresReferences |
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B6.H-2bm12 is a spontaneous mutation of the I-Ab molecule resulting in a 3-aa substitution in the third hypervariable region of the A
chain (6, 7). B6.H-2bm12 skin allografts induce reactive T cells in C57BL/6 recipients and are acutely rejected. In contrast, C57BL/6 mice do not acutely reject B6.H-2bm12 heart allografts, although many of the grafts develop transplant-associated vasculopathy, indicating the induction of a donor-reactive immune response to the graft (5). Mechanisms accounting for the absence of B6.H-2bm12 heart allograft acute rejection despite the development of a potentially pathogenic alloimmune response remain unknown. Such mechanistic insights could prove to be fundamental in the design of therapies aimed at prolonging survival of solid-organ transplants that would otherwise reject.
The number of alloantigens presented to a recipient of a B6.H-2bm12 heart graft is certainly less than that presented to recipients of complete MHC-disparate heart grafts. As a consequence, the T cell repertoire reactive to B6.H-2bm12 alloantigens is less than that reactive to fully MHC-disparate allografts. Nevertheless, the number of anti-B6.H-2bm12 T cell precursors present in the C57BL/6 mouse is sufficient to lead to skin graft rejection. Recent work by our laboratory and by others showed that the frequency of induced effector T cells, rather than the number of naive T cell precursors, can largely determine rejection vs acceptance of a graft (8, 9, 10). Moreover, these studies showed that the threshold number of T cells required to destroy a transplant is dependent on the tissue mass of the transplanted organ. Larger numbers of graft-reactive T cells are needed to reject a heart vs a skin graft, providing an explanation for the preferential rejection of skin vs heart grafts in situations in which low numbers of effector T cells are induced posttransplant.
It therefore becomes essential to understand those factors that determine the size of the effector T cell pool following a given transplant stimulus. The peak size of the effector T cell pool is influenced by the ability of the responding precursor cells to optimally expand during Ag priming. For example, studies performed in infectious disease models have shown that extremely low precursor frequencies can expand to up to 30% of the T cell compartment in response to certain stimuli (11, 12, 13). Such clonal expansion is influenced by the number of Ag-expressing dendritic cells (DC),4 the presence of effective costimulatory signals, the presence of amplifying signals/cytokines (i.e., IFN
and/or IL-12) produced by the innate immune system, and potentially, by inhibitory signals that could lead to apoptosis or clonal exhaustion. In the present study, we investigated the relationship between priming and regulation of alloreactive T cells to B6.H-2bm12 cardiac allografts in C57BL/6 recipients. We provide evidence that regulatory T cells play an important inhibitory role in controlling the size of the effector T cell pool following transplantation of B6.H-2bm12 heart grafts into C57BL/6 recipients. The findings have important implications for therapies aimed at prolonging allograft survival in other model systems.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionDisclosuresReferences |
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B6.C-H-2bm12 (B6.H-2bm12) and C57BL/6 (H-2b) mice were obtained through Dr. C. Reeder at the National Cancer Institute (Frederick, MD). Adult males, 7–12 wk of age, were used throughout this study.
Skin transplantation
Full-thickness trunk skin transplantation was performed using a modification of the protocol of Billingham and Medawar (14). Briefly, trunk skin was excised from B6.H-2bm12 donor mice, the s.c. fat was removed, and 12-mm-diameter circles of full-thickness skin were prepared using a punch. The skin allograft was placed in a slightly larger graft bed prepared over the chest of the recipient and secured with Vaseline gauze and adhesive bandage. After 7 days, the bandage was removed, and each graft was examined daily and was considered rejected when 70% or more of the graft tissue was destroyed as assessed by visual examination.
Heterotopic cardiac transplant
Cardiac transplants were performed using the method of Corry et al. (15). Briefly, donor and recipient mice were anesthetized with phenobarbital. Donor hearts were harvested and placed in chilled lactated Ringer’s solution while the recipient mice were prepared. The donor heart was anastomosed to the recipient abdominal aorta and vena cava using microsurgical techniques. Upon completion of the anastomoses and organ reperfusion, the heart grafts resumed spontaneous contraction. The strength and quality of cardiac impulses were graded daily by palpation as previously described (9). Rejection of cardiac grafts was considered complete by the cessation of impulse and was confirmed visually by laparotomy for each graft. Cardiac isografts in C57BL/6 recipients functioned for >100 days. The significance in allograft survival between recipient groups was analyzed by log-rank test, and p < 0.01 was considered a significant difference between groups. Heart allografts were harvested, snap frozen, and stored at –80°C until use.
Antibodies
The following Abs were used for immunohistology of graft tissue: rat anti-mouse CD4+ (GK1.5) mAb, rat anti-mouse CD8+ (53-6.7) mAb, and biotinylated goat anti-rat polyclonal Ab (BD Pharmingen). ELISPOT assays were performed using IFN-
-specific mAb (BD Pharmingen) and biotinylated anti-IFN- mAb (Vector Laboratories). Rat anti-mouse CD25 mAb PC61 and hamster anti-CTLA-4 mAb UC10-4F10-11 were purified from spent culture supernatant by protein G chromatography. Cardiac allograft recipients were treated with 0.5 mg of PC61 i.p. on day –1, followed by 0.25 mg every other day on days 1–9 or with 0.5 mg of UC10-4F10-11 on day 0 followed by 0.25 mg every other day on days 2–10 posttransplant. In some experiments, mice were thymectomized on day –14 followed by treatment with 0.25 mg of PC61 on days 0, 2, 4, 6, and 9 posttransplant.
DC isolation and expansion
DC were isolated and cultured as previously described (16). Briefly, bone marrow cells were flushed from the femurs and tibia of B6.H-2bm12 mice and cultured for 5 days in medium containing 10 ng/ml GM-CSF and 10 ng/ml IL-4. After the culture, the cells were washed and centrifuged through a 14.5% metrizamide gradient at room temperature for 30 min. The interface cells were harvested and washed three times, and 2.5 x 106 cells were injected i.v. into C57BL/6 recipients 3 days before heart transplantation with a B6.H-2bm12 graft.
Histology and immunohistochemistry
Heart grafts were retrieved from recipients at various times posttransplant, embedded in OCT compound (Sakura Finetek), and frozen at –80°C. Sections were cut at 8 µm and mounted onto slides. For immunohistochemistry, sections were fixed in acetone for 10 min and air-dried. Slides were immersed in PBS for 10 min and then in 0.03% H2O2 for 10 min to eliminate endogenous peroxidase activity. The slides were then stained for 1 h with 5 µg/ml anti-CD4 mAb (GK1.5) or anti-CD8 mAb (53-6.7) in 0.05 Tris-HCl with 1% BSA. Control slides were incubated with rat IgG as the primary Ab. After three washes in PBS for 5 min each, slides were incubated for 20 min with biotinylated goat anti-rat IgG diluted 1/300 in PBS. After three washes in PBS, slides were incubated with streptavidin-HRP (DakoCytomation) for 20 min and washed another three times. To prepare the substrate-chromagen solution, a 10-mg tablet of 3,3'-diaminobenzidine (Sigma-Aldrich) was dissolved in 15 ml of PBS plus 12 µl of 30% H2O2. The solution was applied to the slides, which were incubated for 3–7 min and then rinsed in dH2O to stop the reaction. The slides were counterstained with hematoxylin for 3 min and rinsed with tap water. The slides were dehydrated, viewed under light microscopy, and the images were captured using ImagePro Plus (Media Cybernetics).
Flow cytometry
Spleen cells were obtained from anti-CD25 mAb-treated or control B6 recipients on day +17 or +18 posttransplantation of B6.H-2bm12 heart allografts. The cells were washed twice with staining buffer (Dulbecco’s PBS with 2% FCS/0.2% NaN3), and 1 x 106 cell aliquots were incubated on ice in 150 µl of rat serum (Rockland). After 30 min, the cells were washed twice and stained with fluorochrome-labeled mAb at 10 µg/ml. After 30 min on ice, the cells were washed five times, resuspended in staining buffer, and analyzed by two-color flow cytometry using a FACScan and CellQuest software (BD Biosciences). Sample data were collected on 20,000 gated cells, and cells staining positive for CD4+CD25+ and CD4+rat IgG+ were expressed as the percentage of CD4+ cells.
ELISPOT assays
Priming of donor-specific T cells to IFN--producing cells was quantified by ELISPOT assays as previously described (8, 10). Briefly, ELISA spot plates (Unifilter 350; Whatman) were coated with 2 µg/ml IFN--specific mAb and incubated overnight at 4°C. The plates were blocked with 1% BSA/PBS and then washed four times with PBS. Spleen cell suspensions from graft recipients were prepared on day 7 posttransplant and used as responder cells. Spleen cells from C57BL/6 and B6.H-2bm12 mice were prepared and treated with mitomycin C for use as stimulator cells in the assay as described above. Responder and stimulator cells (1:2) were cultured in serum-free HL-1 medium (BioWhittaker) supplemented with 1 mM L-glutamine. After 24 h of cell culture at 37°C in 5% CO2, cells were removed from the plate by extensive washing with PBS. Biotinylated anti-IFN- (2 µg/ml) or anti-IL-4 (4 µg/ml) mAb was added, and the plate was incubated for 6 h at room temperature. The plate was washed three times with PBS/0.05% Tween 20, and streptavidin-conjugated alkaline phosphatase was added to each well. After 2 h at room temperature, the plates were washed with PBS, and NBT-5-bromo-4-chloro-3-indolyl substrate (Kirkegaard & Perry) was added for the detection of IFN--producing cells. The resulting spots were counted with an ImmunoSpot Series I analyzer (Cellular Technology) that was designed to detect ELISA spots with predetermined criteria for spot size, shape, and colorimetric density.
CD25+ cell depletion and adoptive transfer
For adoptive transfers experiments, CD25+ cells were depleted from spleen cell suspensions by magnetic cell sorting using a CD4+CD25+ T cell isolation kit (Miltenyi Biotec) following the manufacturer’s protocol. Briefly, splenocytes were isolated from naive B6 mice and washed in buffer (PBS, 0.5% BSA, 2 mM EDTA) after lysis of remaining erythrocytes. The cells were incubated with PE-labeled anti-CD25 mAb (Biotech; 10 µl/107 cells) for 15 min at 4–8°C. The cells were washed and resuspended, and 10 µl of anti-PE microbeads were added. Following another 15 min of incubation at 4°C, the cells were washed, resuspended in 500 µl of buffer, and loaded onto the MACS columns. Cells passing through the column were collected as the CD25– fraction. The CD25+ depleted and nondepleted cells were adoptively transferred into RAG–/– recipients of B6.H-2bm12 heart grafts (20 x 106 cells/mouse) on day 3 posttransplantation.
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Results |
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The acute rejection of single class II MHC-mismatched skin and cardiac B6.H-2bm12 allografts by C57BL/6 recipients was compared. All skin allografts were rejected between days 15 and 19 posttransplant (Fig. 1). In contrast, all cardiac allografts were maintained past day 25 posttransplant, and after day 30, 80% of the grafts continued to survive beyond day 100, with median survival time (MST) >100 days (Fig. 1). Histological inspection of long-term-surviving heart allografts indicated little-to-no cellular infiltration when examined at day 80 posttransplant (Fig. 2).
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Cardiac allografts retrieved from recipients that were not primed with DC had little cellular infiltration at day 7 posttransplant, and on day 21 this infiltration increased in some but not all allografts (Fig. 2). In contrast, cardiac allografts retrieved from DC-primed cardiac-grafted recipients on day 7 were heavily infiltrated with mononuclear cells, and this infiltrate was associated with marked myocyte necrosis. By day 21 posttransplant, however, the intensity of this cell infiltration into allografts was absent in most of the heart allografts analyzed.
In parallel to histological analyses, T cell development to effector cells in unprimed and DC-primed recipients of B6.H-2bm12 cardiac allografts was compared by enumerating the number of alloreactive T cells producing IFN- on day 7 posttransplant in ELISPOT assays (Fig. 3). The number of alloreactive cells producing IFN- in cardiac allograft recipients that were not primed with DC was increased 
3-fold over the background response observed in naive mice (74 IFN--producing cells per 6 x 105 cells vs 22 per 6 x 105 cells). Priming with donor DC before receiving the cardiac allograft increased this 1.5-fold further on day 7 posttransplantation (110 IFN--producing cells per 6 x 105 cells; p < 0.01). As expected with a response to a single class II MHC disparity, the number of primed alloreactive T cells producing IFN- was considerably lower than observed in recipients of complete MHC-mismatched heart allografts at the time of rejection, typically >1000 per 6 x 105 cells. On day 21 posttransplantation, the numbers of alloreactive T cells producing IFN- were slightly lower in cardiac allograft recipients not primed with DC (61 per 6 x 105) and were detected at even lower frequencies (21 per 6 x 105; p < 0.05) in DC-primed allograft recipients. Overall, these results indicated a low-level T cell response to a single allogeneic class II MHC determinant that is poorly sustained over time posttransplant and is reflected by low-level cellular infiltration into the heart allografts that also subsides with time posttransplant.
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We next investigated whether negative signals might be restraining acute rejection of B6.H-2bm12 cardiac allografts. One obvious candidate was CTLA-4, which is implicated both as a negative regulator of naive and effector T cells as well as an effector mechanism expressed by CD4+CD25+ regulatory T cells (19, 20, 21, 22). Thus, we tested the effects of a blocking anti-CTLA-4 mAb on the survival of B6.H-2bm12 cardiac allografts. C57BL/6 mice were treated with control Ig or anti-CTLA-4 mAb every other day from days 0 to 10 posttransplant (Fig. 4). Blockade of CTLA-4 induced acute rejection of all B6.H-2bm12 cardiac allografts by day 15 posttransplant, demonstrating a role for CTLA-4 in tempering rejection in this allogeneic response.
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First, groups of C57BL/6 recipients of B6.H-2bm12 heart allografts were treated with control rat IgG or with rat anti-CD25 mAb from day –1 to day 9 posttransplant. When spleen cells were examined at day 17 posttransplant, treatment with anti-CD25 mAb led to an almost 75% decrease in CD4+CD25+ T cells (Fig. 5). There was no increase in CD4+ T cells staining positively with anti-rat IgG, indicating that the decrease was due to deletion of CD25+ T cells and not to blocking of CD25 by the Ab treatment.
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when compared with untreated B6 recipients (750 per 6 x 105 vs 41 per 6 x 105; p < 0.01) (Fig. 8). The increased alloreactive T cell response to B6.H-2bm12 cardiac allografts in recipients treated with anti-CD25 mAb was not accompanied by an increase in self-reactive T cells, because low numbers of spots (<8 per 6 x 105 cells) were observed when recipient cells were cultured with C57BL/6 spleen cells as stimulators in the ELISPOT assay (data not shown).
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Next, adoptive transfer experiments were conducted to further test the role of CD25+ cells in promoting acceptance of single class II MHC-mismatched grafts. B6.RAG–/– mice were transplanted with B6.H-2bm12 hearts, and 3 days later, the recipients were reconstituted with naive B6 splenocytes, which did or did not contain CD25+ cells. Adoptive transfer of CD25+ depleted cells precipitated acute rejection of all grafts by day 20 posttransplantation, whereas only one of five grafts was rejected in recipients reconstituted with undepleted B6 splenocytes (p < 0.01) (Fig. 9). At the time of rejection, cardiac allografts from recipients reconstituted with CD25-depleted spleen cells were heavily infiltrated with CD4+ T cells, whereas allografts from recipients reconstituted with whole spleen cells had little cell infiltration at all times examined (data not shown). Thus, the presence or absence of CD25+ cells influences the intensity of cellular infiltration and the acceptance vs rejection of vascularized allografts expressing a single class II MHC disparity.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionDisclosuresReferences |
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The short duration of the T cell response in recipients of B6.H-2bm12 cardiac allografts with or without donor DC priming suggested the presence of a highly regulated alloreactive response to the single class II-disparate grafts. Recent studies in rodent models have indicated the ability of CD4+CD25+ regulatory cells to inhibit allograft rejection (23, 24, 25, 26). The function of these regulatory T cells is inhibited by Abs to CD25 or to IL-2 (27, 28, 29). However, CD25 is also expressed by activated T cells during Ag priming including alloreactive T cells responding to allografts, and under specific conditions IL-2 binding to its receptor on activated T cells induces cell death (30). Recent studies by Sho et al. (2) have indicated that acute rejection of complete MHC-disparate heart allografts was inhibited by treatment with anti-CD25 mAb. In contrast, MHC-matched/multiple minor histocompatibility-disparate heart allografts normally accepted were rejected when recipients were treated with anti-CD25 mAb. Although it is unclear whether the Ab inhibited the activity of CD4+CD25+ regulatory T cells or acted directly on minor histocompatibility-reactive effector T cells, the Ab treatment was associated with decreased T cell apoptosis and sustained alloreactive T cell responses to the allograft.
In the current studies, treatment of C57BL/6 recipients of B6.H-2bm12 heart allografts with anti-CD25 mAb also promoted acute rejection of the grafts. This rejection was accompanied by a substantial increase in the number of alloreactive T cells primed to the allograft and intense and sustained T cell infiltration into the grafts. The removal of CD25+ T cells before transfer to RAG-deficient recipients of B6.H-2bm12 heart allografts resulted in the ability of the wild-type T cells to reject the grafts. Thus, CD4+CD25+ regulatory cells constrain the clonal expansion of B6.H-2bm12-reactive T cells, resulting in low alloreactive T cell responses and low grades of cellular infiltration into the allografts that is not sustained over time. Several recent studies have reported the expansion of CD4+CD25+ T regulatory cells in response to self and exogenous Ags and the restriction of the Ag-specific effector T cell response by the emergence of the regulatory T cells (31, 32, 33, 34, 35). Similarly, both CD4+CD25+ T regulatory cells and pathogenic effector CD4+ T cells are likely to be activated and expand in response to the B6.H-2bm12 alloantigen. The regulatory cells halt or attenuate clonal expansion of the effector CD4+ T cells during the course of the immune response to the allograft so that the numbers of effector T cells are not sufficient to mediate rejection of the cardiac grafts. The realization that a controlled balance between alloreactive effector and regulatory T cells will promote allograft survival has prompted the recent design of a novel strategy that decreases the effector compartment while maintaining the regulatory compartment and successfully establishes tolerance to MHC-mismatched allografts in murine models (36).
If discrepancies in the numbers of allograft-derived DC accounted for the difference in rejection vs survival of skin vs heart allografts from B6.H-2bm12 donors, then recipient priming with donor DC should have increased the size of the effector T cell response and overcome the failure to acutely reject the heart allografts. In the current studies, priming of C57BL/6 mice with B6.H-2bm12 DC did result in a modest increase in primed alloreactive T cells but did not significantly increase acute rejection of the heart allografts. Furthermore, similar numbers of alloreactive T cells are primed in response to both skin and heart allografts expressing single class I MHC or single minor histocompatibility disparities, arguing against differences in graft-derived DC as a factor influencing acute rejection vs acceptance (9, 10). The current results emphasize the strict control imposed on the expansion of alloreactive effector T cells in response to the B6.H-2bm12 cardiac allografts with or without additional alloantigen priming by DC. The rapid decrease in alloreactive effector T cell numbers observed from days 7 to 21 posttransplant in the spleens of B6.H-2bm12 cardiac allograft recipients primed with donor DC may be indicative of the potency of this regulation in response to the early increase in the effector T cell pool induced in the DC-primed recipients. Another aspect of these studies that warrants consideration and further investigation is that the interstitial DC in the B6.H-2bm12 cardiac allograft, but not DC from skin allografts or the bone marrow-derived DC used to prime the allograft recipients, may express tolerogenic properties that induce or promote this regulation. This regulation was expressed in DC-primed cardiac allograft recipients as well as in cardiac allograft recipients subsequently challenged with a donor skin graft, indicating the dominant nature of this regulation in response to B6.H-2bm12 cardiac allografts. Acute rejection of the cardiac allografts was only quickly and consistently observed when this regulation was removed.
An additional factor that may facilitate the acute rejection of allografts is the induction of an alloreactive Ab response. The close homology of the I-Ab and I-Abm12 molecules is reflected by the reactivity of most anti-I-Ab Abs with I-Abm12 (37). This cross-reactivity suggests that, if induced, a humoral response to B6.H-2bm12 allografts by C57BL/6 recipients would result in autoreactivity. The absence of an Ab response to B6.H-2bm12 allografts may also be indicative of the absence of T cells activated through the indirect alloantigen presentation pathway, which is proposed to be a major factor initiating alloantigen-specific Ab responses (38, 39). Consistent with this, we have been unable to detect an indirect T cell response in C57BL/6 recipients of B6.H-2bm12 heart or skin allografts to peptides incorporating the 3-aa substitutions of I-Abm12 or by immunizing C57BL/6 mice with the peptides (S. Schenk, unpublished results). The response to B6.H-2bm12 allografts by C57BL/6 recipients appears to be stimulated entirely through the direct alloantigen presentation pathway, which may also limit the number of effector T cells primed in response to the allograft. The low-level T cell response to B6.H-2bm12 skin and cardiac allografts might be indicative of a restricted repertoire of alloreactive T cells to the I-Abm12 alloantigen. However, rigorous investigation of the TCR repertoires expressed by B6.H-2bm12-reactive T cells indicates a diverse population of T cells generated in response to the single class II MHC disparity (40).
Collectively, the data of the current report provide new insights into factors that control the size of the T cell repertoire posttransplant. The effector T cell response induced to B6.H-2bm12 grafts under normal conditions is relatively low in number, well below that required to reject heart allografts but high enough to reject a smaller skin allograft. The B6.H-2bm12-reactive T cells are primed to express a pathogenic phenotype but cannot be amplified effectively, and the development of these effector T cells is not sustained. The number of induced effector cells posttransplant is theoretically dependent on the precursor frequency, the proinflammatory signals that activate the innate immune system subsequently amplifying the adaptive response, and the presence or absence of factors that regulate T cell expansion and function. The low precursor frequency cannot solely account for the lack of rejection because the number of alloreactive T cells can be increased significantly. Our data clearly show that naturally developing regulatory T cells are capable of limiting the expansion of transplant-reactive T cells in this strain combination. Interference with the presence or function of CD4+CD25+ T cells leads to a significant expansion of proinflammatory antidonor T cells and precipitates rejection of the heart allografts. These results suggest that, following transplantation, both pathogenic and regulatory T cells are activated. If the alloreactive T cell repertoire is not too large, the regulatory T cells have the capability to control the expansion of the pathogenic effector T cells and limit the extent and duration of T cell infiltration into the allograft.
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Disclosures |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionDisclosuresReferences |
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Footnotes |
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1 This work was supported by National Institutes of Health Grants AI40459 and AI51620 (to R.L.F.), AI37691, AI41521, and AI43626 (to L.A.T.), and AI43578 (to P.S.H.). 
2 L.A.T. and R.L.F. share senior authorship of this work.
3 Address correspondence and reprint requests to Dr. Robert L. Fairchild, NB3-79, Lerner Research Institute, Cleveland Clinic Foundation, 9500 Euclid Avenue, Cleveland, OH 44195-0001. E-mail address: fairchr{at}ccf.org
4 Abbreviations used in this paper: DC, dendritic cell; MST, median survival time.
Received for publication August 31, 2004. Accepted for publication November 24, 2004.
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References |
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induced chemokine Mig. J. Immunol. 163:4878.-chains (CD25): breakdown of a single mechanism of self-tolerance causes various autoimmune diseases. J. Immunol. 155:1151.[Abstract]
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) or vascularized cardiac (
) allografts from B6.H-2bm12 donors. One group of C57BL/6 mice was primed with 2.5 x 106 bone marrow-derived DC from B6.H-2bm12 mice and 3 days later received a B6.H-2bm12 heart allograft (•). Cardiac allograft rejection was confirmed visually in all recipients by laparotomy. MST was not significantly different between both groups of heart allograft recipients.
