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The Journal of Immunology, 2004, 172: 1463-1471.
Copyright © 2004 by The American Association of Immunologists

Production of Donor T Cells Is Critical for Induction of Donor-Specific Tolerance and Maintenance of Chimerism1

Hong Xu, Paula M. Chilton, Yiming Huang, Carrie L. Schanie and Suzanne T. Ildstad2

Institute for Cellular Therapeutics, University of Louisville, Louisville, KY 40202


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Nonmyeloablative conditioning has significantly reduced the morbidity associated with bone marrow transplantation. The donor hemopoietic cell lineage(s) responsible for the induction and maintenance of tolerance in nonmyeloablatively conditioned recipients is not defined. In the present studies we evaluated which hemopoietic stem cell-derived components are critical to the induction of tolerance in a total body irradiation-based model. Recipient B10 mice were pretreated with mAbs and transplanted with allogeneic B10.BR bone marrow after conditioning with 100–300 cGy total body irradiation. The proportion of recipients engrafting increased in a dose-dependent fashion. All chimeric recipients exhibited multilineage donor cell production. However, induction of tolerance correlated strictly with early production of donor T cells. The chimeras without donor T cells rejected donor skin grafts and demonstrated strong antidonor reactivity in vitro, while possessing high levels of donor chimerism. These animals lost chimerism within 8 mo. Differentiation into T cells was aborted at a prethymic stage in recipients that did not produce donor T cells. Moreover, donor Ag-driven clonal deletion of recipient T cells occurred only in chimeras with donor T cells. These results demonstrate that donor T cell production is critical in the induction of transplantation tolerance and the maintenance of durable chimerism. In addition, donor T cell production directly correlates with the deletion of potentially alloreactive cells.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
A major goal of research in transplantation has been to establish safe methods for inducing donor-specific transplantation tolerance, thereby avoiding the expense and toxicity of immunosuppressive agents. We previously demonstrated that conditioning with 700 cGy of total body irradiation (TBI)3 is sufficient to achieve engraftment of MHC-disparate allogeneic mouse marrow in 100% of recipients (1). The effective TBI dose can be significantly reduced if antilymphocyte globulin (2), T cell-specific mAbs (3), cytotoxic drugs (1, 4), or costimulation-blocking agents are added (5). As we approach the minimum threshold of conditioning to establish and maintain tolerance, a dissociation between chimerism and tolerance has emerged that could allow an understanding of the early events that influence the induction of tolerance.

To date there has been considerable debate about which donor cell types are responsible for the induction and maintenance of tolerance in vivo. There is compelling evidence for cross-talk between dendritic cells (DC) and subsets of T cells in up- or down-regulating the immune response that is a major focus of research at present (6, 7). In vitro, tolerogenic DC (Tol-DC) can induce regulatory T cells (Treg), and conversely, Treg can promote the development of Tol-DC (8). The in vivo correlate to this is still being defined. Recent reports have demonstrated that the presence of hemopoietic chimerism does not always reliably confer tolerance (9, 10, 11). Elwood et al. (10) reported that solid organ transplant recipients with microchimerism exhibit a similar frequency and severity of rejection episodes as those without detectable donor chimerism. The dissociation of chimerism and tolerance has also been observed in experimental models of macrochimerism. In a nonradiation-based protocol using antilymphocyte serum and rapamycin, donor class II Ag-positive cells, but not T and/or B cells, were required for tolerance induction (12, 13). In a radiation-based tolerance model, the production of both class II Ag-positive cells and T cells was required for tolerance induction, because when class II knockout (KO) mice and T cell KO mice were used as bone marrow donors, chimerism was achieved, but tolerance was not associated with this chimerism. One limitation to these studies was the relatively short duration of follow-up (11, 13). Thus, the contributions of specific donor cellular subsets in inducing transplantation tolerance remain controversial.

In the present studies nonmyeloablatively conditioned mice were monitored for donor cell-specific lineage production, and this was then examined for its role in the induction of tolerance in this model. Our data demonstrate that pretreatment of the recipient with anti-{alpha}{beta}-TCR and anti-CD8 mAbs can reduce the TBI requirement for establishing mixed chimerism. However, chimerism remained stable only in those chimeras with donor T cell production. Moreover, donor-specific tolerance was observed only in chimeras with donor T cell production, suggesting a critical role for donor T cells in the induction of tolerance. In striking contrast, animals without donor T cell production were not tolerant despite the presence of significant levels of donor chimerism. Both cohorts of chimeras produced donor-derived DC, including the CD11c+/CD11b-/B220+ immunoregulatory Tol-DC subset. Differentiation into T cells was aborted at a very early stage (prethymic) in recipients that did not produce donor T cells. Additionally, donor Ag-driven deletion of recipient T cells occurred only in the chimeras with donor T cell engraftment, suggesting that clonal deletion is the likely mechanism for tolerance induction in this model. As we better understand how hemopoietic stem cell (HSC) chimerism induces tolerance and which cell types are important in this process, immunomodulatory strategies to potentiate the induction of chimerism and tolerance will emerge.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animals

Male C57BL/10SnJ (B10, H-2b), B10.BR/SgSnJ (B10.BR, H-2k), and BALB/cJ (BALB/c, H-2d) mice were purchased from The Jackson Laboratory (Bar Harbor, ME). Animals were housed in the barrier facility at the Institute for Cellular Therapeutics and cared for according to National Institutes of Health animal care guidelines.

Assessment of in vivo depletion of CD8+ and {alpha}{beta}-TCR+ T cells and coating of {gamma}{delta}-TCR+ T cell

The mAbs anti-{alpha}{beta}-TCR (H57-597), anti-{gamma}{delta}-TCR (UC7-13D5), and anti-CD8 (53-6.7) were diluted in saline to 1 ml in previously titrated doses and injected i.v. through the lateral tail vein. The anti-CD8 and anti-{alpha}{beta}-TCR mAb are depleting, whereas the anti-{gamma}{delta}-TCR mAb is nondepleting. To document depletion, peripheral blood was obtained 3 days after mAb treatment from treated mice and was stained with PE-conjugated anti-{alpha}{beta}-TCR (H57-597), anti-{gamma}{delta}-TCR (GL3), and anti-CD8 (53-6.7). Staining was also performed with secondary mAbs of mouse anti-hamster IgG-PE or mouse anti-rat IgG2a-FITC to assure that cells were depleted or coated with mAbs. One hundred micrograms per mouse of each mAb was the dosage required to deplete CD8+ and {alpha}{beta}-TCR+ as well as to saturate {gamma}{delta}-TCR+ T cells in normal recipients.

Chimera preparation

Recipient B10 mice were pretreated i.v. with mAbs of anti-{alpha}{beta}-TCR, anti-{gamma}{delta}-TCR, and anti-CD8, alone or in combination, 3 days before bone marrow transplantation (BMT). All mAbs used in vivo were produced and purified in our laboratory. On day 0, recipients were conditioned with 100, 200, or 300 cGy of TBI ({gamma}-cell 40; Nordion, Ontario, Canada). Animals were transplanted with 15 x 106 untreated B10.BR bone marrow cells via lateral tail vein injection 4–6 h after irradiation. Each experiment was repeated at least three times.

Donor bone marrow was prepared by a modification of the method previously described (1, 14). Briefly, B10.BR donor mice were euthanized, and tibias and femurs were harvested. Bone marrow was expelled from the bones with medium 199 (Life Technologies, Grand Island, NY) containing 10 µg/ml gentamicin (Life Technologies), referred to hereafter as chimera medium (CM). The marrow was then gently prepared as a single-cell suspension using a 3-cc syringe and an 18-gauge needle. The cells were filtered through sterile nylon mesh with 100-µm pores, centrifuged at 1000 rpm for 10 min at 4°C, and resuspended in CM. A cell count was performed, and the cells were diluted to a final concentration of 15 x 106 bone marrow cells/ml CM.

Characterization of chimeras

Recipients were characterized for chimerism using flow cytometry to determine the relative percentages of donor-derived PBL 1 mo after BMT and then monthly. Peripheral blood was obtained through tail vein bleeding and was stained with Abs specific for MHC class I Ags of donor (PE-conjugated anti-H2Kk, 36-7-5, mouse IgG2a) and recipient (FITC-conjugated anti-H2Kb, AF6-88.5, mouse IgG2a) origin. Briefly, 50 µl of whole blood was incubated with Abs for 30 min at 4°C in the dark. The blood was then incubated at room temperature for 6 min with ammonium chloride lysing buffer to eliminate erythrocytes and was washed twice. The analysis was conducted on a FACSCalibur (BD Biosciences, Mountain View, CA) with CellQuest software (BD Biosciences). Multilineage engraftment was assessed by four-color staining for FITC-conjugated, anti-donor-specific Ab (H2Kk) and different fluorochrome (PE, PerCP, and allophycocyanin)-conjugated lineage makers, including T cells (anti-CD4, RM4-5; anti-CD8{alpha}, 53-6.7; and anti-TCR-{beta}, H57-597), B cells (anti-B220, RA3-6B2), NK cells (anti-NK1.1, PK136), DC (anti-CD11c, HL3), and myeloid cells (anti-GR-1, RB6-8C5; and anti-MAC-1, M1/70). The following mAbs were used to analyze T cell development: anti-CD24 (30-F1), anti-CD25 (PC61), and anti-CD44 (IM7). Nonspecific background staining was controlled using isotype control Abs directed against irrelevant Ags conjugated with the same fluorochrome as the experimental Ab (i.e., anti-TNP mouse IgG2a mAb, conjugated with FITC, served as an isotype control for FITC-conjugated anti-H2Kb mouse IgG2a). All mAb were obtained from BD PharMingen (San Diego, CA).

Flow cytometric analysis of TCR V{beta} families

Peripheral blood (80–100 µl) from unmanipulated control mice and mixed chimeras 1–6 mo after reconstitution was stained with anti-V{beta}5.1/2-FITC (MR9-4), V{beta}6-FITC (RR4-7), V{beta}8.1/2-FITC (MR-5-2), or V{beta}11-FITC (RR3-15) vs anti-host H2Kb-PE, anti-CD8-PerCP, and anti-CD4-allophycocyanin (all from BD PharMingen) for 45 min at 4°C. A minimum of 50,000 gated events was collected within the total lymphoid gate the same day of staining. Samples were kept on ice before acquisition. Background staining was determined by FITC-conjugated isotype mAbs.

Skin grafting

Skin grafting was performed by a modification of the method described by Billingham (15). Full-thickness tail skin grafts were harvested from the tails of B10.BR (H2k, donor-specific) and BALB/c (H2d, third-party) mice. Recipient mice were anesthetized with Nembutal (pentobarbital sodium injection; Abbott, North Chicago, IL), and full-thickness graft beds were prepared surgically in the lateral thoracic wall, preserving the panniculus carnosum. The grafts were covered with a double layer of Vaseline (Alba-Waldensian, Rockwood, TN) gauze and a plaster cast. Casts were removed on the seventh day; and grafts were scored by daily inspection for the first month and then weekly thereafter for the percentage of rejection, as reflected by petechial and eschar formation. At the time of cast removal, grafts were inspected for vascular perfusion, absence of infection, and technical success. Rejection was defined as complete when no residual viable graft could be detected.

One-way MLR

MLR were performed as previously described (16). Briefly, splenocytes were made into single-cell suspensions, lysed free of RBC, washed, and resuspended in DMEM supplemented with 5% FBS, 1 mM sodium pyruvate, 2 mM L-glutamine, 10 mM HEPES buffer solution, 0.137 M L-arginine-HCl, 1.36 mM/0.027 M folic acid/L-asparagine, 100 U/ml penicillin, 100 U/ml streptomycin (all from Life Technologies), and 0.05 mM 2-ME (Sigma-Aldrich, St. Louis, MO). Responder cells (2.5 x 105) were cultured 1:1 with irradiated stimulator cells (2000 cGy) for 5 days at 37°C in 5% CO2. Each well was pulsed with 1 µCi of [3H]thymidine (DuPont-NEN, Boston, MA) 16 h before harvesting with an automated harvester (PHD Cell Harvester Technology, Cambridge, MA).

Statistical analysis

Data are presented as the average ± SD. One-tailed t test (two-sample, assuming unequal variances) was used to evaluate statistical differences. The difference between groups was considered significant at P < 0.05.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Pretreatment of the recipient with anti-{alpha}{beta}-TCR plus anti-CD8 mAb lowers the requirement of TBI for engraftment

In this study we characterized which specific cell populations in the host must be eliminated to enhance allogeneic bone marrow engraftment, with the goal of eliminating the requirement for cyclophosphamide and reducing the TBI dose as low as possible. To evaluate the efficacy of targeting different T cell populations in allogeneic engraftment, recipient B10 mice were pretreated with anti-{alpha}{beta}-TCR, anti-{gamma}{delta}-TCR, anti-CD8, anti-{alpha}{beta}-TCR plus anti-{gamma}{delta}-TCR, and anti-{alpha}{beta}-TCR or anti-{gamma}{delta}-TCR plus anti-CD8 mAbs 3 days before BMT. On day 0, mice were conditioned with 300 cGy of TBI and then transplanted with 15 x 106 untreated bone marrow cells from B10.BR donors 4–6 h later (Table I). Chimerism was followed by flow cytometric analysis monthly for up to 6 mo after BMT. With 300 cGy of TBI, allogeneic engraftment did not occur in animals preconditioned with anti-CD8 alone, anti-{gamma}{delta}-TCR alone, or anti-CD8 plus anti-{gamma}{delta}-TCR. In contrast, high levels of engraftment were established in animals preconditioned with anti-{alpha}{beta}-TCR alone, anti-{alpha}{beta}-TCR plus {gamma}{delta}-TCR, and anti-{alpha}{beta}-TCR plus anti-CD8 (87.5, 90, and 94%, respectively) 1 mo after BMT. Interestingly, long term engraftment (up to 6 mo) was only achieved in mice preconditioned with both anti-{alpha}{beta}-TCR and anti-CD8. The fact that both Abs in combination achieved the most durable engraftment in the highest proportion of recipients suggests that these agents may be targeting additional CD8+ cells in the recipient that are not {alpha}{beta}-TCR+.


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  		Table I. Percent engrafted in mice pretreated with indicated mAb or mAb combinations and conditioned with 300 cGya

 
A dose titration of TBI was performed to determine the minimal conditioning required in combination with preconditioning with the anti-{alpha}{beta}-TCR plus anti-CD8 mAb combination. B10 mice were pretreated with anti-{alpha}{beta}-TCR plus anti-CD8 on day -3 and transplanted with 15 x 106 untreated bone marrow cells from B10.BR donors after conditioning with 0, 100, 200, or 300 cGy of TBI. Ninety-four percent of mice conditioned with 300 cGy (n = 16) engrafted 1 mo after BMT (Fig. 1A). Seventy-five percent engrafted with 200 cGy (n = 16; Fig. 1A). Only 20% of mice engrafted when conditioned with 100 cGy (n = 5), and none engrafted without TBI (n = 6). The levels of donor chimerism observed in the engrafting recipients directly correlated with the amount of conditioning at 75.8 ± 7.7, 45.7 ± 12.6, and 5.0% 1 mo after BMT with 300, 200, and 100 cGy of TBI, respectively (Fig. 1B). Engraftment remained high (83.3% at 6 mo after BMT) in animals conditioned with 300 cGy of TBI, whereas the majority of engrafted animals conditioned with 200 cGy (eight of 12) lost their chimerism by 6 mo, as did all conditioned with 100 cGy of TBI (Fig. 1C). These results demonstrate that the TBI dose can be reduced from 700 to 300 cGy by depletion of both host {alpha}{beta}-TCR+ and CD8+ T cells in vivo. Moreover, these results further confirmed our previous finding that both {alpha}{beta}-TCR+ and CD8+ T cells in the host play critical and nonredundant roles in preventing engraftment of allogeneic bone marrow (17).



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  		FIGURE 1. Characteristics of engraftment and the level of donor chimerism in B10 recipients conditioned with increasing doses of TBI. B10 mice were pretreated with anti-{alpha}{beta}-TCR and anti-CD8 mAbs on day -3. On day 0 they were transplanted with 15 x 106 untreated bone marrow cells from B10.BR donors 4–6 h after conditioning with 100 (n = 5), 200 (n = 16), or 300 (n = 16) cGy of TBI. Unirradiated controls (n = 6) were also performed. Animals were analyzed for engraftment by flow cytometric analysis monthly for up to 6 mo after BMT. This figure shows the frequency of engraftment 1 mo after BMT (A), level of chimerism in animals that engrafted (percentage of donor cells in PBL) at 1 mo (B), and the kinetics of engraftment for up to 6 mo after BMT (C), as assessed by PBL typing. The results are the summary of three experiments.

 
Production of donor T cells is critical for the maintenance of stable mixed chimerism

The pluripotent HSC produces progenitors that differentiate into at least 11 different lineages. To evaluate the influence of multilineage production on the durability of engraftment and induction of tolerance, animals were followed for >=4 mo by four-color, flow cytometric analysis (donor class I vs lineage marker). In the engrafted recipients we found that the chimeras could be separated into two groups based on the presence of donor-derived T cells regardless of the TBI dose used for conditioning that absolutely predicted the induction of tolerance in vivo as well as the stability of chimerism (Fig. 2). In one group, although B cells, NK cells, monocytes, and DC of donor origin were detected, no donor-derived {alpha}{beta}-TCR+, CD4+, or CD8+ T cells were present (Table II and Fig. 2A). All chimeras conditioned with 100 cGy (n = 1) and 200 cGy (n = 12) of TBI and some of the chimeras conditioned with 300 cGy (n = 6) of TBI did not have donor T cell production (Table II and Fig. 2A). The second group of chimeras (Fig. 2B), which produced donor T cells, had all been conditioned with 300 cGy of TBI. These phenotypes did not change during the time course that chimerism was monitored.



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  		FIGURE 2. Detection of donor and host-derived cells of lymphoid and myeloid lineages in mixed allogeneic chimeras using four-color flow cytometry. Multilineage typing was performed 2–3 mo after BMT on animals that exhibited high levels of donor chimerism. The x-axis shows staining with fluorescein-conjugated Ab against donor class I Ag (H2Kk). On the y-axis, the mAb staining for the different cell lineage markers is shown: {beta}-TCR, CD8, CD4, NK1.1 (NK cells), B220 (B cells), Mac-1 (macrophages), Gr-1 (granulocytes), and CD11c (dendritic cells). The results of representative chimeras are presented from chimeras with no donor T cells (A) and those with donor T cell engraftment (B). Lineages of host origin were also evaluated in both groups of chimeras.

 

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  		Table II. Donor multilineage engraftment in recipients with or without donor T cell engraftmenta

 
The durability of chimerism strictly correlated with the production of donor T cells. In animals with donor T cell production, mixed chimerism was stable for >=6 mo (Fig. 3B). The level of chimerism was 82.7 ± 6.4% at 1 mo post-transplantation in this group. At 3 mo, donor chimerism was 87.9 ± 14.1% and remained stable at 6 mo (67.2 ± 18.8%). In striking contrast, the majority of animals without donor T cell production lost their chimerism gradually within 6 mo (Fig. 3A). Only four of these animals had low level donor chimerism at 6 mo, and this chimerism was lost by 8 mo. The initial percent chimerism ranged from 5.0–75.3% in this cohort. The level of chimerism significantly decreased over time in these mice. Although chimerism averaged 49.1 ± 19.5% at 1 mo and 30.7 ± 18.1% at 2 mo post-transplantation (p < 0.0.5), donor cells became undetectable in the blood between 3 and 8 mo in this group. These findings suggest that the early production of donor T cells is critical for the maintenance of stable chimerism.



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  		FIGURE 3. Production of donor T cells is critical for the maintenance of stable chimerism. Animals were analyzed for the level of donor chimerism by flow cytometric analysis monthly for up to 6 mo after BMT. Animals rendered chimeric by preconditioning with {alpha}CD8 plus {alpha}{beta}-TCR mAb and varying doses of TBI were divided into two groups according to results from PBL typing for multilineage engraftment: chimeras without donor T cell engraftment (A) and chimeras with donor T cell engraftment (B). This figure shows the kinetics of the level of donor chimerism in each individual animal for up to 6 mo after BMT in groups of chimeras without (A) and with (B) donor T cell engraftment. The results are a summary of four experiments.

 
Subsets of DC have been shown to be immunogenic, whereas others are tolerogenic (8). To confirm that the lack of tolerance we observed was related to the lack of production of donor T cells and was not secondary to a specific lack of the tolerogenic subset of DC, the DC from the spleens of chimeras were evaluated for immunogenic (CD11c+/CD11b+/B220-) and tolerogenic (CD11c+/CD11b-/B220+) subpopulations by flow cytometry. Both DC subpopulations were represented in the donor and recipient lymphoid gates. No significant difference was observed between the two groups. In both groups, donor as well as host DC were present (data not shown; p > 0.3).

Production of donor T cells is critical for induction of donor-specific tolerance to skin grafts

Skin grafts were performed to assess donor-specific tolerance in vivo in the two groups. Each animal received a skin graft from donor-specific (B10.BR) and third-party (BALB/c, H2d) strains 2–3 mo after BMT. Grafts were assessed daily for the first 4 wk and weekly thereafter for evidence of rejection. Animals that failed to engraft donor stem cells at 1 mo after transplantation rejected both donor and third-party grafts promptly (median survival time (MST), 10 days for both grafts). The chimeras without donor T cell production rejected third-party skin grafts in a fashion similar to that in mice without chimerism. Surprisingly, however, the majority of chimeras without donor T cells also promptly rejected donor skin grafts as well (MST, 12.5 days), with a time course similar to the third-party grafts, despite the presence of significant levels of donor chimerism at the time of graft placement (Fig. 4). The level of donor chimerism in this group was 37.6 ± 27.4% (range, 7.1–72.7%) at the time skin transplantation was performed. Only one of 18 (5.6%) donor skin grafts survived >120 days in this group. In chimeras with donor T cell engraftment, donor-specific allogeneic skin grafts were permanently accepted (MST, >120 days) in seven of nine mice, and the survival of the other two grafts was prolonged, whereas third-party skin grafts were promptly rejected. Donor-specific skin graft survival in the group with donor T cell engraftment was significantly prolonged compared with that of chimeras without donor T cell production (p < 0.00005). These data show that production of donor T cells is critical for the induction of functional tolerance in nonmyeloablatively conditioned recipients.



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  		FIGURE 4. The production of donor T cells is critical for the induction of donor-specific tolerance to skin grafts. Animals were divided into three groups according to the results at 1 mo from PBL typing for donor chimerism and multilineage engraftment: 1) animals without engraftment, 2) chimeras without donor T cell engraftment, and 3) chimeras with donor T cell engraftment. Each animal received skin grafts from donor-specific (B10.BR) and third-party (BALB/c, H2d) strains 2–3 mo after BMT, and the grafts were then monitored up to 120 days. The level of donor chimerism at the time of skin transplantation and the MST of skin graft survival ± SD are shown. Donor-specific skin graft survival in the group with donor T cell engraftment was significantly longer than that in the group without donor T cell engraftment (p < 0.00005).

 
Only chimeras with production of donor T cells are functionally tolerant in vitro

Splenic lymphoid cells from chimeras with (n = 8) or without (n = 6) donor T cell production as well as from recipients that did not engraft (nonchimeras; n = 7) were assessed for donor-specific tolerance in vitro using one-way MLR assays directed against host, donor, and third-party irradiated stimulator cells. As shown in a representative one-way MLR assay (Fig. 5), chimeras with donor T cells exhibited tolerance to both host (B10) and donor strain (B10.BR) stimulators, but were reactive to MHC-disparate, third-party (BALB/c) alloantigen. Nonchimeras were reactive to both donor and third-party alloantigens, but were not reactive to host stimulators. Chimeras without donor T cells exhibited reactivity to donor and third-party stimulators. Moreover, proliferation in the presence of host stimulators and even in medium control wells was as high as that in donor and third-party wells. These results suggest the spleens from these animals contained both donor and host cells that were not tolerant to each other. Therefore, with medium alone, the host T cells proliferate in response to stimulation from the allogeneic donor cells of mixed chimeras. These data support the observation that donor cells are eventually rejected in chimeras without donor T cells. These in vitro data also confirm the presence of specific tolerance to donor strain alloantigens in chimeras with peripheral donor T cells and the absence of donor-specific tolerance in chimeras without peripheral donor T cells, consistent with the results of skin graft survival for these two groups of chimeras.



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  		FIGURE 5. Chimeras without donor T cells do not exhibit donor-specific tolerance in vitro. Lymphocytes from mixed chimeras with (n = 8) or without (n = 6) peripheral donor T cells as well as from recipients that did not engraft (nonchimera; n = 7), were cocultured with irradiated host (B10), donor (B10.BR), and third-party (BALB/c) stimulator cells in the MLR assay. Values are shown as the mean ± SD of triplicate cultures in a 1:1 responder to stimulator ratio from a representative experiment.

 
Production of donor T cells is critical for clonal deletion of donor-reactive TCR-V{beta} subsets

We hypothesized that donor T cell engraftment may be critical to induce or regulate the deletion of graft-reactive cells. To investigate whether clonal deletion is operational in our model, we measured the expression of superantigen-specific T cells, V{beta}5.1/2, V{beta}6, V{beta}8.1/2, and V{beta}11, in chimeras with or without donor T cell engraftment. Naive B10 and B10.BR splenocytes served as controls. The donor strain B10.BR mice express I-E, resulting in the deletion of V{beta}5.1/2+ and V{beta}11+ T cells (18, 19). As B10 mice do not express I-E, they do not delete these two V{beta} subfamilies (19, 20). Relative expression indicates the percentage of V{beta}-positive cells within the CD8+ or CD4+ T cell subsets of the host (H2Kb) lymphoid gate in peripheral blood. The host lymphoid gate used in the current study for clonal deletion analysis is more accurate than the total (host plus donor) lymphoid gate used previously (21, 22, 23) by avoiding an effect from the deleted donor V{beta}5.1/2+ and V{beta}11+ cells.

As shown in Fig. 6, recipient CD4 and CD8 cells from chimeras with donor T cell production showed the same relative V{beta} expression as B10.BR mice, indicating that V{beta}5.1/2+ and V{beta}11+ subfamilies of both CD4+ and CD8+ T cells had been deleted (Fig. 6; p < 0.005). This negative selection was specific, as V{beta}6+ and V{beta}8.1/2 subsets were not deleted. Chimeras without donor T cells exhibited V{beta} expression similar to recipient B10 mice in CD8+ T cells, indicating that no deletion of V{beta}5.1/2+ and V{beta}11+ subfamilies of CD8+ T cells occurred (Fig. 6A). However, partial deletion of V{beta}5.1/2+ and V{beta}11 of CD4+ T cells was observed in chimeras without donor T cells (Fig. 6B). The levels of V{beta}5.1/2+ and V{beta}11+ CD4+ T cells in chimeras without donor T cells were significantly higher than those in chimeras with donor T cell production (p < 0.005). However, the reduced levels of V{beta}5.1/2+ and V{beta}11+ CD4+ T cells in chimeras without donor T cells were statistically significant compared with control B10 mice (Fig. 6B; p < 0.05), suggesting that partial deletion had occurred. This partial deletion was not sufficient to induce host hyporesponsiveness to donor alloantigen. At the time of analysis, animals still had significant levels of donor chimerism in the groups with (68.9 ± 15.8%) or without (48.9 ± 19.3%) donor T cell production.



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  		FIGURE 6. Relative V{beta}-TCR expression in chimeras with or without donor T cell production. Expression of V{beta}5.1/2 ({square}), V{beta}6 (), V{beta}8.1/2 (), and V{beta}11 ({blacksquare}) on PBL from unmanipulated hosts (B10), unmanipulated donors (B10.BR), chimeras with donor T cell engraftment, or chimeras without donor T cell engraftment was measured by FACS analysis. Relative expression represents the percentage of V{beta}-positive cells within the CD8+ (A) or CD4+ (B) T cell subsets of the host (H2Kb) lymphocytes in peripheral blood. Data from three experiments are depicted as the mean ± SD. V{beta}-TCR expression in either chimeric group was compared with that in B10 mice using one-tailed t test (two-sample, assuming unequal variances). Significant p values are indicated above the respective data bars (*, p <= 0.005; {psi}, p <= 0.05).

 
Early stage pre-T cells (CD24+/CD4-/CD8-) are not present in the thymus of chimeras without peripheral donor T cells

To evaluate where the block of T cell development was taking place in the chimeras without donor T cell production, we analyzed donor-derived MHC expression on pre-T cells in the thymus of naive and chimeric animals using flow cytometry. CD24 (heat-stable Ag) is a marker of T lineage commitment in early stages of T cell development in the thymus (24). It is expressed at high levels during the double-negative (CD4-/CD8-) stage of T cell development (25). A single-cell thymocyte suspension was prepared from chimeras with or without donor T cells and from naive B10 or B10.BR mouse controls. Cells were stained with CD4-allophycocyanin, CD8-PE, CD24-PerCP, and donor class I H2Kk-FITC. As shown in Fig. 7, donor CD24+ T cells were not detected in the double-negative (CD4-/CD8-) thymocytes in chimeras devoid of donor peripheral T cells. Their staining pattern was identical with that of naive recipient B10 mice. In contrast, donor CD24+/CD4-/CD8- thymocytes were present in chimeras with donor peripheral T cells. The staining pattern of these chimeras strongly resembled that of naive B10.BR controls. The donor pre-T cells from chimeras with donor T cells were present in the thymus at all stages of T cell maturation (24), from the most immature CD4-/CD8-/CD24+/CD44+/CD25- to the mature single-positive CD4+/CD8- or CD4-/CD8+ cells (data not shown).



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  		FIGURE 7. Analysis of the expression of donor MHC class I on CD24+/CD4-/CD8- thymocytes in chimeras with or without donor T cells. Thymocytes were prepared from chimeras with (n = 5) or without (n = 4) peripheral donor T cells and from naive control B10 or B10.BR mice. They were stained with CD4-allophycocyanin, CD8-PE, CD24-PerCP, and donor class I H2-Kk-FITC. Cells that were negative for both CD4 and CD8 were gated (A) and further analyzed for the expression of donor class I (H2-Kk) and CD24 (B) as an indication of pre-T cell commitment. The data shown are representative staining of one mouse from each group.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
A number of incompletely ablative approaches have been developed to establish mixed chimerism (1, 2, 26, 27). Conditioning of recipients with TBI plus cyclophosphamide (day 2) allowed mixed chimerism to be established with as low as 500 cGy of TBI (1). The addition of antilymphocyte globulin to the regimen allowed the TBI dose to be reduced to as low as 300 cGy of TBI (2). When we evaluated which recipient cells must be targeted by preconditioning the recipient with subset-specific mAbs, CD8+ and {alpha}{beta}-TCR+ cells were found to be the primary effectors (3, 28). In the present studies we combined anti-CD8 mAb pretreatment with anti-{alpha}{beta}-TCR to attempt to further reduce the TBI dose for conditioning, with the rationale that these specific host cellular subsets controlled the hemopoietic microenvironment for HSC engraftment. In studies that defined the threshold dose of TBI required for engraftment of syngeneic vs allogeneic marrow, Down et al. (29) were one of the first to propose that conditioning may be targeting specific host effector cells with differing radiation sensitivities. Our own data using mAbs in lieu of irradiation would strongly support that hypothesis, because the administration of specific mAb resulted in a significantly reduced requirement for TBI.

The fact that a combination of anti-CD8 mAb and anti-{alpha}{beta}-TCR mAb was more potent for conditioning than either Ab alone could be due to one of two possible mechanisms. One explanation for this could be residual cell populations remaining after each mAb is administered. The presence of immunocompetent residual cells after mAb treatment that can mediate allorejection has been previously reported (30, 31). Rosenberg et al. (31) reported that there was a small number of residual CD8+ cells left in recipient animals despite the fact that >99% of the CD8+ cells had been removed, and further, that these residual cells mediated the rejection of allogeneic skin grafts. When two mAbs with overlapping cell specificities were administered, more residual alloreactive cells were targeted. Therefore, our data showing that the combination of anti-{alpha}{beta}-TCR and anti-CD8 is more effective in establishing allogeneic chimerism than either mAb used singly is in agreement with these reports. A second possibility is that more than one host cell type contributes to alloresistance, and that in addition to {alpha}{beta}-TCR+ T cells, a second CD8+/TCR- cell serves as an effector cell as well. Regardless of the mechanism, our data indicate that high dose conditioning plays more of a role in providing suppression of host-vs-donor reactive cells than making space in light of the host effector cells identified.

To our surprise we observed that the early production of donor T cells in the partially conditioned host is an absolute prerequisite for sustained chimerism and induction of allograft tolerance. Notably, the production of donor B cells, NK cells, granulocytes, or DC in chimeras that did not produce T cells did not contribute to the durability of chimerism or the induction of tolerance. Moreover, the presence of immunoregulatory or tolerogenic DC of donor and host origin was not sufficient to induce tolerance in those chimeras without donor T cell production in vivo. Although a feedback loop involving cross-talk between Tol-DC and induction of Treg, and vice versa, has been shown in vitro, an in vivo model has not yet been fully defined (8, 32). Our data confirm that donor T cell chimerism is critical to induce tolerance to achieve durable engraftment in vivo. One could hypothesize that a critical cell population in the host with a radiation sensitivity at 200–300 cGy of TBI is responsible for this dichotomy, or that cellular cross-talk between host and donor HSC or progenitors determines whether tolerance is induced at this threshold level of conditioning. However, there must be at least partial suppression of a host-vs-graft response against the donor bone marrow cells, as the donor chimerism persists for up to 8 mo in nontolerant animals despite a vigorous MLR in vitro and donor skin graft rejection.

Although a strict correlation between HSC chimerism and tolerance has historically been demonstrated in ablated recipients (14, 27, 33, 34), recent reports have challenged the relationship between chimerism and tolerance, especially in partially conditioned and immunosuppressed recipients. Recipients of solid organ grafts have a low level, systemic, donor-derived chimerism, or microchimerism (35, 36), that may contribute to the decreased requirement for immunosuppression over time. However, despite the presence of microchimerism, most solid organ allograft recipients do not become drug free. Moreover, a dissociation of the presence of microchimerism and the establishment of functional tolerance has been reported in some solid organ allograft recipients (9, 10, 37). Although low levels of donor cells were detected systemically in recipients of heart and liver allografts, microchimerism was present even during rejection (9, 10). It has therefore been argued that microchimerism is a result, but not a cause, of long term graft survival (38).

The dissociation of chimerism and tolerance has also been observed in models establishing macrochimerism. In an allogeneic mouse model similar to that used in this study, a small fraction of animals conditioned with anti-CD8 and anti-CD4 mAb plus 300 cGy of TBI failed to produce significant numbers of donor-type T cells, and donor chimerism declined quickly. In contrast, animals with donor T cell engraftment showed stable chimerism and accepted both primary and secondary donor-specific skin grafts (39). Similarly, production of donor T cells at levels <50% are correlated with a higher rate of graft rejection in humans (40). These data suggest that donor T cell chimerism was associated with the maintenance of long term allograft acceptance in irradiation-induced tolerance and are consistent with our own findings. However, in our own studies the duration of chimerism in animals without T cell production was significantly prolonged. A dissociation of chimerism and tolerance was also observed in a mouse model in which T cell split chimerism was established using different T cell KO mice as donors. When KO mice deficient in both CD4 and CD8 T cells or CD3{epsilon} transgenic mice lacking both T cells and NK cells were used as donors, high levels of donor chimerism resulted, but the animals were not tolerant to donor-specific grafts. Tolerance was restored when donor-type T cells were adoptively transferred to recipient mice initially given T cell-deficient bone marrow (11). The limitation to these studies was that the animals were followed for <=4 mo for durability of chimerism, and the mechanism responsible for the observation was unclear.

In nonradiation-based chimerism protocols using antilymphocyte serum and rapamycin, it has been clearly shown that donor T cells are not required for the induction of tolerance (41). Allogeneic chimerism was achieved, but donor T cells were not produced. Interestingly, chimeras showed specific functional tolerance, as evidenced by acceptance of second-donor grafts and rejection of third-party grafts. The lack of requirement for donor T cells for tolerance induction in this model was also confirmed using T cell-deficient knockout mice as donors (42). Furthermore, in this nonradiation-based model, donor class II Ag-positive bone marrow cells were required for tolerance induction (12), and when class II-deficient KO mice were used as donors, tolerance did not occur. Taken together, these data and our own suggest that the critical donor hemopoietic cells that initiate the early steps in tolerance induction might be influenced by the conditioning approach used, an observation very important for clinical application.

To determine the mechanism required for the early events occurring in the induction of tolerance in our model, we evaluated whether donor T cell production is critical to induce clonal deletion of graft-reactive cells. In chimeras with donor T cell production, recipient T cell deletion of V{beta}5.1/2+ or V{beta}11+ T cells occurred in both CD8 and CD4 T cells as expected. Chimeras without donor T cell engraftment showed no reduction in the percentage of V{beta}5.1/2+ or V{beta}11+ CD8 cells. However, there was a partial reduction in the percentage of V{beta}5.1/2+ or V{beta}11+ CD4 cells compared with chimeras with donor T cell engraftment. Therefore, the ability to affect clonal deletion was directly correlated to the production of donor T cells and was not due to the presence of superantigen alone, nor did it correlate with the production of donor DC, which were present in both groups. This study provides the first evidence that donor T cell production after engraftment is associated with clonal deletion of donor-reactive T cells as well as the maintenance of durable engraftment of MHC-disparate hemopoietic stem cells after transplantation. It is only in this context that chimerism is associated with functional tolerance.

To evaluate T cell development in chimeras with or without donor T cells, CD4-/CD8- thymocytes were stained and analyzed for CD24 expression as a marker of T lineage commitment. There were no CD24+ donor early stage pre-T cells (CD4-/CD8-) in chimeras that did not produce donor T cells. In contrast, donor CD24+/CD4-/CD8- cells and all expected subsequent stages of T cell maturation were present in thymus in chimeras with donor T cell production. These data suggest that the block of T cell development occurs at an extrathymic stage in maturation. The lack of mature donor T cells in peripheral blood and even at a very early stage of pre-T cells in thymus might be due to the fact that donor-derived T lymphoid progenitors do not migrate to the recipient thymus, or that donor T lymphoid progenitors migrate to the thymus, but are blocked very early before commitment to the T lineage. Taken together, these data demonstrate a strong correlation between donor T cell maturation in the thymus and the induction of tolerance, most likely by clonal deletion.

Our findings suggest that some components of donor chimerism may be ancillary phenomena, but not necessarily the mechanism for tolerance induction. We hypothesize that as there are no donor-derived T cells present in the thymus, there are no donor Ags present to mediate negative selection of the alloreactive T cell clones. Thus, donor-specific tolerance could occur if clonal deletion is initiated in the host, regardless of the type of conditioning used or which donor cell populations engrafted. This would explain why different donor cell components are required for tolerance induction in radiation-based vs nonradiation-based tolerance models. Tolerance could instead depend on the components that allow induction of host clonal deletion, which may be influenced by the conditioning approach used as well as the types of donor lineage cells present in the host. It is possible that confirming evidence for an active process for clonal deletion may be a more reliable predictor of clinical tolerance to organ allografts than the presence of donor chimerism per se.

In light of the fact that production of donor T cells absolutely correlated with functional donor-specific tolerance in vivo and in vitro as well as efficient clonal deletion, our results emphasize the importance of donor T lineage-specific chimerism in the maintenance of stable mixed chimerism and induction of donor-specific tolerance after nonmyeloablative allogeneic BMT. These results provide indirect evidence that clonal deletion is the likely mechanism for tolerance induction in our BMT model. A clear definition of the requirements that influence the induction of chimerism and tolerance will allow in vitro and clinically relevant in vivo strategies to potentiate the ability to establish chimerism with minimal or no conditioning.


   Acknowledgments
 
We thank Carolyn DeLautre for manuscript preparation, the staff of the animal facility for outstanding animal care, and Loretta Doan for help with experimental design for the thymocyte studies.


   Footnotes
 
1 This work was supported in part by National Institutes of Health Grant R01HL63442-01, the Commonwealth of Kentucky Research Challenge Trust Fund, the Jewish Hospital Foundation, and the University of Louisville Hospital. Back

2 Address correspondence and reprint requests to Dr. Suzanne T. Ildstad, Institute for Cellular Therapeutics, Department of Surgery, University of Louisville, 570 South Preston Street, Suite 404, Louisville, KY 40202-1760. E-mail address: stilds01{at}gwise.louisville.edu Back

3 Abbreviations used in this paper: TBI, total body irradiation; BMT, bone marrow transplantation; CM, chimera medium; DC, dendritic cell; HSC, hemopoietic stem cell; KO, knockout; MST, median survival time; Tol-DC, tolerogenic dendritic cell; Treg, regulatory T cell. Back

Received for publication August 11, 2003. Accepted for publication November 19, 2003.


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