Posttransplantation administration of donor bone marrow cells in conjunction with transient, peri-operative immunosuppression using rabbit anti thymocyte globulin (RATG) can induce prolonged and specific unresponsiveness to skin and vascularized allografts in rodent, dog and outbred primate models (1–4). Recent studies have shown that the administration of donor bone marrow is accompanied by a reduction in the frequency of cytolytic T cell precursors (5,6), requires the presence of CD95-ligand (Fas-ligand) on donor cells (7) and is abrogated by depletion of CD8+ cells from the cell infusions (5). These observations have led to the hypothesis that donor cells effect the deletion of cytolytic T cell precursors (CTLp) by inducing apoptosis in the alloreactive T cells that interact with them, and that the process is dependent on the CD8 molecule (8). A mechanism of this type has been previously described by Miller and colleagues (9,10), who showed that the addition of viable cells syngeneic with the stimulator cells in a cell-mediated lympholysis assay resulted in functional deletion of reactive allospecific effector T cells. The same group also showed that the effect can occur in vivo after administration of lymphoid cells syngeneic to the stimulating alloantigen (11). There is evidence to implicate the CD8 molecule in the regulatory effect of allogeneic donor bone marrow cells. Blocking the CD8 molecule on the alloantigen bearing cells with monoclonal antibodies, or using cells lacking functional CD8 molecules, abrogated deletion of cytolytic T cell precursors by allogeneic cells (12–14). Such results are consistent with studies demonstrating the CD8 accessory molecule can act as an inhibitory ligand (13,15,16).
Although we have postulated that the inhibition of graft rejection by donor bone marrow cells is mediated by a CD8 dependent mechanism (5), it has not been shown whether the CD8 molecule is essential in transplant tolerance induced by donor bone marrow cells when they are infused into antithymocyte serum treated transplant recipients.
Monaco and colleagues have shown that donor spleen cell infusion can produce an effect that is functionally identical to that of bone marrow, providing that a dose two times stronger is used (17,18). The cellular composition of splenocytes is vastly different from that of bone marrow cells. Therefore, we also postulate that the suppressive effect mediated by spleen cells could be the product of a different cell subpopulation and/or a different molecular mechanism. The experiments described in this report strongly support this idea. The prolongation of graft survival by donor bone marrow cells in this model is dependent on the presence of the CD8 molecule on the MHC incompatible donor cells, whereas extension of graft survival by donor spleen cells is not CD8 dependent.
MATERIALS AND METHODS
Six- to eight-week old C3H/HeJ, (C57BL/6 x A)F1 (B6AF1), C57BL/6-CD8atm1Mak (CD8KO) (19), C57BL/6J, B10.D2(R107), and BALB/cJ mice were purchased from the Jackson Laboratory, Bar Harbor, ME. On arrival, the mice were placed in isolator cages and fed Purina rodent chow ad libitum.
On the day of transplantation (Day 0), a 1.0×1.5 cm piece of skin was removed from the thorax and replaced with abdominal skin from the donor. The graft was fixed in place using 8–10 interrupted 6–0 polyfilament sutures. In addition, a piece of skin removed from the recipient’s abdomen was grafted on the opposite side of the thorax as an autograft control. The grafts were then covered with gauze impregnated with white petrolatum and secured into place using a plaster body cast. Animals were scored every other day for rejection beginning at day 7. A graft was scored as rejected if it had shrunken to 20% of the initial size or was covered with a scab over 100% of the original area. Antithymocyte serum was created as previously described (7).
Donor cell preparation and sorting.
On the day of bone marrow injection (day +7), the plaster cast was removed and the indicated quantities of donor bone marrow cells were injected i.v. in the tail vein. Bone marrow cells were prepared by removing the hind leg bones of mice and flushing out the bone marrow cells using a syringe filled with complete RPMI 1640 containing 5% fetal calf serum and 1 U/ml heparin. The cells were washed in white Hanks balanced salt solution without phenol red, then resuspended at the appropriate concentration for injection in a volume of 0.2 ml PBS. Spleens were harvested from donor strain mice, disaggregated through a stainless steel mesh in osmolarity adjusted (320 mosm) phosphate buffered solution (PBS). The single cell suspension was then passed through a 30-micron nylon mesh filter. Spleen cells were injected i.v. into recipients through the tail vein on day +7 relative to transplantation. Purified subsets of spleen cells were infused in numbers corresponding to their presort proportions. To deplete CD8+ cells, spleen cells were incubated for 15 min on ice with microbeads conjugated to a monoclonal rat anti-mouse CD8α (Ly-2) antibody (Clone 53–6.7, CD8a microbeads, Miltenyi Biotec, Auburn, CA). The suspension was subsequently passed over a column containing a ferrous matrix (MACS, Miltenyi Biotec, Auburn, CA) in a magnetic field. To enrich for CD8+ cells, incubation was as above, but the suspension was passed over a MACS enrichment column. After the CD8 negative (CD8−) cells were washed away, the enrichment column was removed from the magnetic field, and the CD8+ cells were then eluted from the column using PBS supplemented with 0.5% bovine serum albumin. CD8+ cells were also enriched by negative selection. In this instance, donor spleen cells were treated with ammonium chloride (0.15 M) solution to lyse red blood cells and were then incubated with magnetic microbeads conjugated to rat anti-mouse CD4 antibodies (Clone GK1.5), MHC class II (Clone M5/114.15.2) and CD11b (Mac-1α, Clone M1/126.96.36.199) respectively in the same fashion as the CD8α microbeads. The suspension was subsequently passed over a MACS depletion column, and the CD8+ cells were collected from the column effluent.
Because CD8+ cells were present at much lower concentrations in bone marrow, CD8+ cells were enriched by magnetic sorting as above, but were further enriched by fluorescent activated cell sorting using a FACSVantage flow cytometer (Becton Dickinson, Hialeah, FL) to a final purity of 90%.
Flow cytometry analysis.
Various subsets of donor cells (5×105 cells) were analyzed by flow cytometry using the following monoclonal antibodies (all obtained from Pharmingen, Torrey Pines, CA): biotin-conjugated rat anti-mouse CD8α (Ly-2), fluorescein isothiocyanate (FITC) conjugated rat anti-mouse CD8α (Ly-2), FITC conjugated hamster anti-mouse CD3ε (clone 2C11), biotin conjugated rat anti-mouse CD11b (Mac-1), biotin-conjugated rat anti-mouse B220 (CD45R) and phycoerythrin-conjugated rat anti-mouse DX5 (pan NK cell). Cells were analyzed with a FACSscan flow cytometer (Becton Dickinson, Hialeah, FL).
Statistical Analysis. Survival of skin allografts were measured in days, and compared between groups by the Log-rank test.
The effect of depletion or enrichment of CD8+ cells in donor cell infusions.
As previously shown (18), treatment of (C57BL/6 X A)F1 (here after called B6AF1) mice with ATS on days −1 and + 2, relative to skin transplantation from a C3H donor followed by administration 2.5×107 donor bone marrow cells, results in an antigen-specific prolongation of graft survival compared with animals that were immunosuppressed with ATS alone (Fig. 1, MST=49 days vs. MST=27 days respectively, P =0.003). This protective effect did not extend to third party grafts (not shown). Similarly, the administration of 5.0×107 donor spleen cells also conferred extended graft survival (MST=44 days, Fig. 2) that was not significantly different from that conferred by bone marrow cells (compare Figures 1 and 2, P =0.3).
To determine whether CD8+ cells in the donor bone marrow infusion play a role in the prolongation of graft survival, we performed experiments in which the proportion of CD8+ donor bone marrow cells was enriched by magnetic sorting followed by fluorescence activated cell sorting from <0.5% to >90%. Infusion of these cells resulted in significant prolongation of graft survival (MST=41 compared with no infusion MST=27, P =0.04, Figure 1) at a dose that was 250-fold lower (1×105 cells per recipient) than the unsorted bone marrow. The survival was not significantly different than the group that received unsorted BMC (P =0.40). A dose of 1×105 cells was chosen because we have found that CD8+ cells constitute 1.3±0.6% of total bone marrow cells in male C3H mice (7). Therefore, 2.5×107 bone marrow cells could contain 1.75×105 to 4.75×105 CD8+ cells. In a separate experiment, a dose of 5×104 CD8+ cells failed to extend allograft survival relative to controls that received no BMC (MST=36 days vs. 37 days, respectively).
For donor splenocyte infusions, we routinely obtained a purity of >90% CD8+ cells using positive selection with magnetic microbeads conjugated to a monoclonal rat antimouse CD8α antibody. These cells were infused at a dose such that the absolute numbers of CD8+ cells was the same as that received when unsorted spleen cells were administered (approximately 4×106 cells/recipient). Skin allografts on animals that were infused with CD8+ splenocytes (MST=29 days) did not survive longer than grafts on control animals that did not receive spleen cells (MST=27 days, P =0.28); graft survival was significantly worse than that obtained when unsorted spleen cells were infused (MST=44 days, P =0.0007, Figure 3). In contrast, recipients treated with spleen cells depleted of the CD8+ subpopulation to <0.5% (MST=56 days) exhibited enhanced allograft survival compared with the controls that received ATS treatment only (MST=27, P =0.0002). There was no statistically significant difference in survival times between CD8 depleted and unsorted spleen cells (Fig. 3, P =0.2).
Given that graft survival was not prolonged by CD8+ donor spleen cells, it is possible that the sorting procedure could have altered the function of the CD8+ cells, particularly because the beads used for sorting were not removed before infusion. Therefore, we repeated the experiment using CD8+ enriched cells sorted by negative selection with beads conjugated to rat anti-mouse MHC class II, anti CD11b, and anti-CD4. Recipients were infused with 5×106 CD8+ cells purified in this manner. As an additional control, unsorted spleen cells were subjected to a sham sort in which they were treated with magnetic beads conjugated to anti-CD8α monoclonal antibodies and passed through a ferrous wool column in the absence of a magnetic field. The negatively selected CD8+ spleen cells (MST=36) failed to enhance graft survival (Fig. 4, P =0.6 compared with controls receiving no spleen cells). The sham sorted spleen cells, however, conferred extended allograft survival (MST=49 days) compared with controls that received no spleen cells (MST=27 days, P =0.003). This was not a statistically significant difference from animals infused with untreated spleen cells (MST=44 days, P =0.70, Figure 4). These data support the hypothesis that donor subpopulations that can extend allograft survival are CD8+ in bone marrow and CD8− in the spleen.
The presence of the donor CD8 molecule affects prolongation of graft survival.
To corroborate the findings in the sorting experiments, and to directly investigate the role of the CD8 molecule, we used donor spleen cells and bone marrow cells from mice that were deficient in the expression of the CD8 molecule because of an induced mutation in the CD8α gene (CD8 “knockout mice”) (19). To perform these experiments, we changed the strain combination because the induced mutation was bred into C57BL/6 mice. The classical C3H to B6AF1 strain combination is matched at the MHC class I and class II loci except for H2-D. To maintain this difference, and to use the C57BL/6 knockout mice as skin and cell donors, B10.D2(R107) (H2: Kb, Ab, Eb, Dd) mice were used as recipients. These two strains are also matched at the MHC class I and class II loci except for H-2D, with the exception that C57BL/6 does not express H-2E.
Figures 5 and 6 depict the results of experiments in which B10.D2(R107) mice were treated on days −1 and +2 with 0.5 ml ATS relative to the day of transplantation with skin from a C57BL/6J mouse. On day +7, the mice received either 2.5×107 wild-type bone marrow cells or 5×107 wild-type splenocytes from C57BL/6J mice. A significant augmentation of skin graft survival was observed in recipients of wild-type donor bone marrow (MST=205) versus those that received no donor bone marrow (MST=70 days, P =0.004, Figure 5) and recipients of wild-type donor spleen cells (Fig. 6, MST=74 days) versus those that received no donor spleen cells (MST=29 days, P =0.0002). Therefore, in our hands, the C57BL/6 to B10.D2(R107) strain combination behaved in a manner similar to that of the classical C3H to B6AF1 strain combination in that the relationship between the various experimental groups was similar, albeit with generally longer survival times in all groups.
The use of bone marrow from CD8 knockout mice resulted in a significant diminution of graft survival relative to animals that received wild type donor bone marrow (MST=98 days and MST=205 days, respectively, P =0.04, Figure 5); graft survival was not significantly higher in recipients of CD8 KO bone marrow cells in comparison to those that received no bone marrow (MST=70, P =0.16). Therefore, the CD8 molecule is required for the optimal prolongation of allograft survival by donor bone marrow cells.
To determine the effect of the absence of the CD8 molecule on donor spleen cells, B10.D2(R107) mice were injected with ATS on days −1 and +2 relative to transplantation with C57BL/6 skin. On day + 7 relative to transplantation, animals were allocated into three groups: 1) infusion of 5.0×107 wild type C57BL/6 donor spleen cells/mouse, 2) infusion of 5.0×107 CD8 knockout C57BL/6 donor spleen cells/mouse, and 3) no infusion (ATS controls). Figure 6 indicates that both CD8 knockout (MST=145 days, P =0.0002 vs. no SPC) and wild type donor spleen cells (MST=74 days, P =0.0002 vs. no SPC) were able to significantly augment graft survival compared with ATS controls. Moreover, there was no statistical difference in graft survival between groups treated with CD8 knockout or wild-type spleen cells (P =0.4), indicating that the CD8 molecule is not necessary for the induction of augmented graft survival by donor spleen cell infusions.
Flow cytometric analysis of the distributions of major cell lineages in both spleen and bone marrow, in normal and CD8 knockout mice, with monoclonal antibodies specific for CD11b, B220, CD3, and Dx5 (a pan NK marker) showed that, other than a small increase in the number of B220+ cells and a small reduction in the number of CD3+ cells in the mutant mice, there was no significant difference in the proportions of cells bearing the lineage markers (Table 1).
The experiments described in this report are part of a series of studies in which we are analyzing the critical molecular events involved in the induction of operational tolerance after donor bone marrow administration. We define “operational tolerance” to mean the statistically significant extension of graft survival beyond that of the negative control without the use of immunosuppressive therapy beyond the initial dose. Monaco, Wood, and colleagues established that the infusion of bone marrow into recipients pretreated with anti-lymphocyte serum or antithymocyte globulin results in the establishment of donor specific tolerance (1,18,20). More recent evidence suggests that graft specific T cells undergo deletion after administration of donor cells. Limiting dilution analysis has shown that donor specific T cells are functionally eliminated (5,6), and we have shown that the extension of graft survival by donor bone marrow is CD95/CD95-ligand dependent (7), suggesting a deletional process in which donor bone marrow cells actively induce apoptosis in graft specific T cells. Thomas and colleagues have further shown that the down-regulation of allospecific responses by bone marrow cells in vitro requires cell to cell contact, and also requires the presence of CD8+ CD2+CD16+ MHC class II− cells, a result that mirrors requirements for the extension of rhesus monkey renal allograft survival in vivo (5, 8, 21, 22). On the basis of the in vitro and in vivo concordance using fractionated cell populations, they hypothesized that the extension of graft survival by donor bone marrow is due to a veto mechanism of action, as described by Miller and colleagues (10, 23, 24).
The veto hypothesis, as applied to the extension of graft survival by donor bone marrow cells, postulates that the donor cells inactivate or delete graft specific T cells that recognize allogeneic antigens on their surface through the T cell receptor (21, 25). Such a process requires cell to cell contact, and likely involves transmission of molecular signals in both directions. This certainly would involve a number of molecules on the surface of both cells. The data showing that depletion of CD8+ cells from donor bone marrow abrogates the ability to extend graft survival in primates (5, 21) suggest the CD8 accessory molecule on cell subpopulations in donor bone marrow may be directly involved in the down-regulation of alloresponses by bone marrow cells. The studies described in this report were designed to directly address this issue and further addressed the question of whether cells derived from other sources, such as donor spleen, use different mechanisms. The latter possibility was suggested by recent experiments in our laboratory showing that tolerance induction by infusion of donor spleen, unlike donor bone marrow, was not CD95/CD95-ligand dependent (Goldstein DR, Chang T, Sweeney SD, Kirklin JK, Thomas JM, George JF, Transplantation, in press).
We have shown that purified CD8+ donor bone marrow cells can induce tolerance at a dose that is reduced 250 times relative to unsorted bone marrow cells. This and the failure of bone marrow cells from CD8 knockout mice to induce tolerance strongly suggest that CD8 molecules on donor bone marrow cells play a direct role in the prolongation of graft survival. In contrast to donor bone marrow cells, donor splenocytes did not seem to require the presence of CD8+ donor cells because the removal of CD8+ cells resulted in no detectable effect on graft survival relative to unsorted spleen cells. The method used for the isolation of splenic CD8+ cells did not seem to affect the ability to confer graft survival. Treatment of unsorted spleen cells with bead conjugated anti-CD8 antibodies did not affect their ability to confer graft survival. Furthermore, splenic CD8+ cells that were isolated using either negative or positive selection methods lacked the ability to extend graft survival.
The experimental system used to test donor cells from CD8 knockout mice relied on a different strain combination than the sorting experiments. One notable difference in this strain combination is that survival times were generally longer, albeit the relative differences between experimental groups were similar for both strain combinations. ATS treated animals, for example, exhibited median graft survival times of 27 days in the C3H to B6AF1 strain combination, and 70 days in the C57BL/6 to B10.D2(R107) strain combination. The precise reason for these differences are unknown, but the most likely explanation is that there are fewer minor histocompatibility differences between the C57BL/6 and B10.D2(R107) strains, which are both C57 derivatives.
The differential dependence on the expression of CD8 for spleen and bone marrow cells could be a result of differences in the lineage or maturity of cells that effect the prolongation of graft survival. Such a concept is consistent with the observation that deletional mechanisms in immune responsiveness and development are frequently redundant. It is also consistent with the view that the veto phenomenon describes an activity that can be mediated by a number of different cell types (8). The possibility of involvement of different veto cell types also suggests the existence of multiple molecular pathways for functional deletion of graft reactive T cells by donor cells.
Wood and colleagues have previously reported that donor splenocytes capable of prolonging skin graft survival are MHC class II−, Thy-1−, and surface Ig−, which is consistent with our observation that splenocytes capable of prolonging graft survival are CD8− (18,26). Their studies of bone marrow subpopulations also showed similar results, indicating that the active subpopulations in both spleen and bone marrow are probably not T cells. The evidence presented in this report supports the hypothesis that donor spleen cells and bone marrow cells capable of prolonging allograft survival could be NK cells or dendritic cells, respectively, because both lineages have been reported to mediate deletion of allospecific cells under certain conditions, and there are subpopulations of each of these cell types that express CD8 on the cell surface (5). Our current efforts are directed towards the investigation of these possibilities and towards the elucidation of other molecular signals involved in the prolongation of graft survival by donor cells from bone marrow and spleen.
There have been clinical trials involving the use of donor derived bone marrow cells as a means of reducing clinical rejection and graft loss. Further information regarding the molecular mechanisms by which bone marrow cells, or cells from other sources, can mediate a salutary effect on graft survival will facilitate rational decisions as to how the clinical application of this technique can be improved.
1. Gozzo JJ, Wood ML, Monaco AP. Use of allogenic, homozygous bone marrow cells for the induction of specific immunologic tolerance in mice treated with antilymphocyte serum. Surg Forum 1970; 21: 281.
2. Thomas JM, Carver FM, Foil MB, Hall WR, Adams C, Fahrenbruch GB, Thomas FT. Renal allograft tolerance induced with ATG and donor bone marrow in outbred rhesus monkeys. Transplantation 1983; 36: 104.
3. Thomas JM, Carver M, Cunningham P, Park K, Gonder J, Thomas F. Promotion of incompatible allograft acceptance in rhesus monkeys given posttransplant antithymocyte globulin and donor bone marrow. I. In vivo parameters and immunohistologic evidence suggesting microchimerism. Transplantation 1987; 43: 332.
4. Hartner WC, De Fazio SR, Markees TG, Maki T, Monaco AP, Gozzo JJ. Specific tolerance to canine renal allografts following treatment with fractionated bone marrow and antilymphocyte serum. Transplant Proc 1987; 19: 476.
5. Thomas JM, Carver FM, Kasten-Jolly J, et al. Further studies of veto activity in rhesus monkey bone marrow in relation to allograft tolerance and chimerism. Transplantation 1994; 57: 101.
6. Wood ML, Orosz CG, Gottschalk R, Monaco AP. The effect of injection of donor bone marrow on the frequency of donor-reactive CTL in antilymphocyte serum-treated, grafted mice. Transplantation 1992; 54: 665.
7. George JF, Sweeney SD, Goldstein DR, Kirklin JK, Thomas JM. An essential role for fas-ligand in transplantation tolerance. Nat Med 1998; 4: 333.
8. Thomas JM, Verbanac KM, Carver FM, et al. Veto cells in transplantation tolerance. Clin Transplant 1994; 8: 195.
9. Miller RG, Derry H. A cell population in nu/nu spleen can prevent generation of cytotoxic lymphocytes by normal spleen cells against self antigens of the nu/nu spleen. J Immunol 1979; 122: 1502.
10. Miller RG. An immunological suppressor cell inactivating cytotoxic T-lymphocyte precursor cells recognizing it. Nature 1980; 287: 544.
11. Martin DR, Miller RG. In vivo administration of histoincompatible lymphocytes leads to rapid functional deletion of cytotoxic T lymphocyte precursors. J Exp Med 1989; 170: 679.
12. Sambhara SR, Miller RG. Programmed cell death of T cells signaled by the T cell receptor and the alpha 3 domain of class I MHC. Science 1991; 252: 1424.
13. Hambor JE, Weber MC, Tykocinski ML, Kaplan DR. Regulation of allogeneic responses by expression of CD8 alpha chain on stimulator cells. Int Immunol 1990; 2: 879.
14. Zhang L, Shannon J, Sheldon J, Teh HS, Mak TW, Miller RG. Role of infused CD8+ cells in the induction of peripheral tolerance. J Immunol 1994; 152: 2222.
15. Kaplan DR, Hambor JE, Tykocinski ML. An immunoregulatory function for the CD8 molecule. Proc Natl Acad Sci USA 1989; 86: 8512.
16. Hambor JE, Kaplan DR, Tykocinski ML. CD8 functions as an inhibitory ligand in mediating the immunoregulatory activity of CD8+ cells. J Immunol 1990; 145: 1646.
17. Monaco AP, Wood ML. Studies on heterologous antilymphocyte serum in mice. VII. Optimal cellular antigen for induction of immunologic tolerance with antilymphocyte serum. Transplant Proc 1970; 2: 489.
18. Monaco AP, Wood ML, Maki T, Gozzo JJ. The use of donor-specific bone marrow to induce specific unresponsiveness (tolerance) to tissue allografts. In: Ildstad KJ, ed. Chimerism and tolerance. Austin: R.G. Landes, 1995
19. Schilham MW, Fung-Leung WP, Rahemtulla A, et al. Alloreactive cytotoxic T cells can develop and function in mice lacking both CD4 and CD8. Eur J Immunol 1993; 23: 1299.
20. Monaco AP, Gozzo JJ, Wood ML, Liegeois A. Use of low doses of homozygous allogeneic bone marrow cells to induce tolerance with antilymphocyte serum (ALS): tolerance by intraorgan injection. Transplant Proc 1971; 3 (1): 680.
21. Thomas JM, Carver FM, Cunningham PR, Olson LC, Thomas FT. Kidney allograft tolerance in primates without chronic immunosuppression: the role of veto cells. Transplantation 1991; 51: 198.
22. Thomas JM, Carver FM, Cunningham P, Olsen L, Thomas FT. Veto cells induce long-term kidney allograft tolerance in primates without chronic immunosuppression. Transplant Proc 1991; 23: 11.
23. Miller RG, Muraoka S, Claesson MH, Reimann J, Benveniste P. The veto phenomenon in T-cell regulation. [Review]. Ann N Y Acad Sci 1988; 532: 170.
24. Carlow DA, Teh SJ, van Oers NS, Miller RG, Teh HS. Peripheral tolerance through clonal deletion of mature CD4-CD8+ T cells. Int Immunol 1992; 4: 599.
25. Muraoka S, Miller RG. Cells in bone marrow and in T cell colonies grown from bone marrow can suppress generation of cytotoxic T lymphocytes directed against their self antigens. J Exp Med 1980; 152: 54.
26. George JF, Goldstein DR, Thomas JM. Donor bone marrow and transplantation tolerance: historical perspectives, molecular mechanisms and future directions. Int J Mol Med 1999; 4: 475.