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Published in final edited form as: Biol Blood Marrow Transplant. 2012 Aug 11;18(11):1629–1637. doi: 10.1016/j.bbmt.2012.08.004

Lack of Antidonor Alloantibody Does Not Indicate Lack of Immune Sensitization: Studies of Graft Loss in a Haploidentical Hematopoietic Cell Transplantation Swine Model

Raimon Duran-Struuck 1, Abraham Matar 1, Rebecca Crepeau 1, Ashley Gusha 1, Marian Schenk 1, Isabel Hanekamp 1, Vimukthi Pathiraja 1, Thomas R Spitzer 2, David H Sachs 1, Christene A Huang 1
PMCID: PMC3575102  NIHMSID: NIHMS439214  PMID: 22892552

Abstract

Loss of chimerism is an undesirable outcome of allogeneic hematopoietic cell transplantation (HCT) after reduced-intensity conditioning. Understanding the nature of cellular and humoral immune responses to HCT after graft loss could lead to improved retransplantation strategies. We investigated the immunologic responses after graft loss in miniature swine recipients of haploidentical HCT that received reduced-intensity conditioning. After the loss of peripheral blood chimerism, antidonor cellular responses were present without detectable antidonor antibody. Reexposure to donor hematopoietic cells after graft loss induced a sensitized antidonor cellular response. No induced antidonor antibody response could be detected despite evidence of cellular sensitization to donor cells. In contrast, unconditioned animals exposed repeatedly to similar doses of haploidentical donor cells developed antidonor antibody responses. These results could have important implications for the design of treatment strategies to overcome antidonor responses in HCT and improve the outcome of retransplantation after graft loss.

Keywords: Hematopoietic cell transplantation

INTRODUCTION

Reduced-intensity conditioning (RIC) regimens have facilitated hematopoietic cell transplantation (HCT) in patients who previously would not have been considered suitable candidates, owing to the toxicity of myeloablative preparatory regimens [1]. Although the risk of complications related to conditioning is decreased with RIC, the risk of graft loss is increased. Graft loss, whether secondary to rejection, the inability of the stem cells to engraft due to the lack of “space” [2], or poor graft quality [3], can have different but important immunologic consequences in the host. Rejection of the donor graft implies an active immunologic process in which donor cells sensitize the host (through cellular and/or humoral mechanisms). Conversely, if the loss of donor cells is not immunologic but rather related to a deficiency in stem cell “fitness” or quality [3], there may be no immunologic consequences (eg, sensitization). Factors related to graft loss include donor–recipient MHC mismatch, degree of host myeloablation, level of immunosuppression post-HCT, degree of host immunocompetence related to immediate preparatory regimens, level of T cell depletion of the donor graft, and presensitization to donor antigens, as is seen in patients with aplastic anemia [4–6]. To date, few clinical studies have assessed immune responses of patients after graft loss and reexposure of donor antigen.

In this study, we investigated the immunologic responses after graft loss in the Massachusetts General Hospital MHC-defined miniature swine, a clinically relevant large-animal model of HCT [7]. Recipients underwent RIC and received cytokine-mobilized peripheral blood mononuclear cells (PBMCs) haplomismatched at both MHC I and MHC II. The RIC regimen consisted of CD3 immunotoxin, 100 cGy of total body irradiation (TBI), and 45 days of cyclosporine A (referred to as the “ITC” regimen hereinafter). Seventeen ITC-conditioned haplo-HCT recipients were engrafted with donor stem cells and maintained moderate to high (30%–70%) donor-derived chimerism in all hematopoietic lineages [8,9]. Four ITC-conditioned haplo-HCT recipients that did not engraft and lost peripheral blood chimerism are discussed in this report. Cellular and humoral antidonor MHC responses were studied before and after reexposure to donor antigen. Antidonor immune responses were compared in HCT recipients and naïve animals exposed to donor antigen. Our findings provide insight into the immunologic responses after graft loss and may serve as a guide for modifying preparatory regimens when subsequent retransplantation of immune-sensitized hosts is considered.

MATERIALS AND METHODS

Animals

Animals were selected from our herd of MGH MHC-defined miniature swine [10,11]. The donors ranged in age from 4 to 8 months, and the recipients were age 9–12 weeks and weighed 9–13 kg at the time of HCT. Donor and recipients were chosen to differ by 1MHChaplotype at both MHCI and MHCII. Recipients expressed swine leukocyte antigen (SLA)ad, and donors expressed SLAac, hereinafter referred to as AD and AC, respectively. To facilitate the detection of chimerism, the donors were positive for pig allelic antigen (PAA), a nonhistocompatibility cell surface antigen present on all differentiated hematopoietic cells in animals that express this gene allele [12]. The recipients were PAA-negative.

HCT Protocol

The HCT protocol consisted of a combination of irradiation, T cell depletion, and hematopoietic cell infusion from a single-haplotype MHC-mismatched donor, with 45 days of cyclosporine cover for the peri-infusion/postinfusion period. Serum cyclosporine (CyA) levels were maintained between 400 and 800 ng/uL. T cell depletion was achieved using pCD3 immunotoxin (CD3-IT) [13] for 4 days before HCT. This recombinant CD3 immunotoxin selectively binds porcine CD3 and contains a diphtheria toxin subunit, which results in protein synthesis inhibition and depletion of CD3+ cells [14]. Animal 18862 did not receive CD3-IT.

Donor animals were cytokine-mobilized for 5–7 days with recombinant porcine IL-3 and porcine stem cell factor (Immerge Biotherapeutics, Cambridge, MA), each at a dose of 0.1 mg/kg for the first 30 kg of body weight and 0.05 mg/kg for each additional kg, as reported previously [8]. Peripheral blood mononuclear cells (PBMCs) were collected by leukapheresis (COBE BCT, Lakewood, CO) beginning on the 5th day of cytokine therapy and continuing until the target cell number was attained. After the initial leukapheresis, 5–12 × 109 PBMCs per kg (total of 5–12 × 1010 PBMCs, as animals averaged 10 kg in weight) were infused i.v. daily. Enteral CyA (Sandimmune) was administered via a gastrostomy tube starting at 1 day before mobilized PBMC infusion and continuing for 45 days. CyA whole blood levels were maintained at 400–800 ng/mL for the first 30 days before being tapered over the subsequent 15 days to 200 ng/mL, at which point CyA was discontinued.

Donor Antigen Exposure after HCT

Intravenous delivery of unselected PBMCs

Nonmobilized leukocytes were collected by leukapheresis from the original hematopoietic cell donor and then infused i.v. into the recipient at a normalized dose (CD3+ T cells) to include 5 × 107 donor T cells/kg of recipient body weight. Animals that underwent HCT received a donor leukocyte infusion (DLI) from the same HCT donor animal.

Subcutaneous PBMCs

Donor PBMCs were collected from donor whole blood. Eighty million donor PBMCs were washed with PBS and injected s.c. with a 19-gauge needle (in the inguinal region) with or without complete Freund’s adjuvant. Cells were suspended in a total volume of 5 mL PBS before injection. The animals that underwent HCT were immunized with PBMCs from the same donor.

Skin grafting

A vascularized skin flap [9] was transplanted from the inguinal region of the donor and placed on the neck of the recipient. The skin graft was monitored daily for color, temperature, and texture change.

Assessment of Chimerism

PAA is expressed on hematopoietic cells from PAA+ donor animals but not in PAA− recipients. Peripheral blood, bone marrow, and thymic donor chimerism were assessed by flow cytometry (FACScan; BD Biosciences, San Jose, CA), as described previously [8,12,15]. The following swine-specific antibodies were used: CD3ε (898H2-6-15; mouse IgGaK) [16], CD4 (74-12-4; mouse IgG2bK), CD8α (76-2-11; mouse IgG2aK), CD172 (74-22-15; mouse IgG1K) [17–20], CD5 [21], and PAA (1038H-10-9; IgMK), For assessment of chimerism, PAA staining was used to distinguish donor-origin and recipient-origin cells [20]. Monocyte and granulocyte chimerism was determined by gating on CD172+ mononuclear cells and granulocytes, respectively.

Mixed Lymphocyte Reaction and Cell-Mediated Lymphotoxicity Assays

In primary mixed lymphocyte reaction (MLR), responders (4 × 105 cells) and stimulators (4 × 105 cells irradiated with 25 Gy) were cultured together. The responders and stimulators were plated at 200 μL/well in triplicate and incubated for 5 days at 37°C in 5% CO2 and 100% humidity. Proliferation of responder cells was assessed by measuring the uptake of H3 thymidine after 5 hours of incubation. Cell-mediated lymphocytotoxicity (CML) assays were performed as described previously [22]. In brief, CML tissue culture medium included fetal porcine serum. The medium consisted of basal DMEM (Life Technologies, Grand Island, NY) supplemented with 6% serum replacement medium (Sigma-Aldrich, St. Louis, MO). For both CML and MLR assays, PBMCs from noninbred Yucatan or Yorkshire pigs were used as third-party positive stimulators and targets. Stimulation indices were calculated by dividing the average counts per minute detected for a particular responder–stimulator pair by the average counts per minute for the same responder stimulated by self-stimulators.

Complement-Mediated Antibody-Dependent Cellular Cytotoxicity Assay

Allogeneic cytotoxic antibodies to donor PBMCs were detected by complement-mediated cytotoxic assays, as described previously [23,24]. In brief, target cells were diluted to 5 × 106 cells/mL and suspended in Medium199 (Cellgro, Herndon, VA) supplemented with 2% FCS. In 96-well U-bottom plates (Costar, Cambridge, MA), 25 μL of the appropriate target cell suspension was incubated with 25 μL of serum serially diluted from 1:2 to 1:1024 or controls for 15 minutes at 37°C, followed by a second incubation with 25 μL of appropriately diluted rabbit complement. Dead cells were identified by staining with 10 μL of 7-AAD for 30 minutes. Data were acquired, and the percentage of dead cells was assessed using a FACScan (BD Biosciences) and analyzed with WinList analysis software (Verity Software, Topsham, ME).

RESULTS

In the current study, we investigated the immune responses of animals that lost peripheral blood chimerism within 200 days of HCT (Figure 1 and Table 1). Animals 19138–19140 underwent the same ITC regimen, whereas animal 18862 received a modified regimen without porcine CD3 immunotoxin consisting of 45 days of CyA and 100 cGy of TBI at 2 days before HCT. These 4 animals lost peripheral blood chimerism within 80 days (animals 19139, 19140, and 18862) and 150 days (animal 19138) after HCT. All assays performed assessed the response of PBMCs.

Figure 1.

Figure 1

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(A) Groups 1 and 3 received the ITC preparatory regimen and 45 days of immunosuppression. Groups 1 and 2 received mobilized PBMCs at a dose of 5–10 billion cells/kg. A DLI of 50 million Tc/Eq/kg was given to animals in groups 1 and 2 without immunosuppression coverage. Group 3 received a skin graft 2 months after discontinuation of immunosuppression. (B) Donor-derived chimerism in animals 19138–19140 that received RIC (100 cGy TBI, 45 CyA, CD3 immunotoxin). Animal 18862 received 100 cGy TBI and 45 days of CyA without CD3 immunotoxin. Peripheral blood chimerism was lost in animals 19139, 19140, and 18862 by day 80 and in animal 19138 by day 150. Percentage donor chimerism was measured by the PAA+ marker (y-axis). Days post-HCT is shown on the x-axis. Group 1 comprises animals 19138, 19139, 19140, and 18862; group 2 includes animals 19937 and 19938; and group 3 includes animal 20312.

Table 1.

Summary of Cellular and Humoral Outcomes in Experimental Animals That Underwent HCT with Immunosuppression (Group 1), Animals That Received No Immunosuppression but Were Exposed to Donor Antigen (Group 2), and 1 Animal That Received Both a Preparatory Regimen and Donor Antigen, but Not at the Same Time (Group 3)

Group Animal RIC Regimen HCT First Antigen Exposure Antidonor Antibodies Sensitized Antidonor Cellular Responses Second Antigen Exposure Antidonor Antibodies Sensitized Antidonor Cellular Responses Major Contributor to Antidonor Response
(1) Experimental HCT 19138
19139
19140
18862		 Yes Yes i.v. (at time of HCT) No No i.v. No Yes Sensitized cellular alloresponses
No antibody produced after skin graft more than 1 year after HCT
(2) Naïve, i.v. only 19937
19938		 No Yes IV Yes No i.v. Yes No Alloantibody
(3) Conditioning only (no HCT) 20312 Yes No Skin (after D/C CyA) Yes Yes ND ND ND Alloantibody and sensitized cellular alloresponses
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ND indicates not done; D/C, discontinue.

Antidonor Cellular Responses after Graft Loss Are Comparable to Naïve Alloresponses

After graft loss (weeks 14–16), antidonor cellular cytotoxic responses appeared similar to responses seen before HCT (Figure 2A and B). MLRs were performed to assess for evidence of cellular sensitization in these animals. When animals are sensitized to MHC-mismatched donor antigen, proliferative responses are maximal after only 2 or 3 days of stimulation, and by day 5 there is no (or minimal) further proliferation based on thymidine uptake (Figure 3A and B). In swine, normal alloproliferative responses (without previous donor antigen exposure) across MHC barriers are maximal after 5 days of stimulation in vitro (Figure 3B). Early (day 3) and standard 5-day proliferative responses of animals 19138–19140 to donor MHC-matched PBMCs were assessed to determine whether the animals exhibited a pattern of sensitization similar to that shown in Figure 3A. Similar to animals not previously exposed to donor antigen, the 4 animals that lost chimerism after ITC-conditioned haplo-HCT showed maximal proliferation after 5 days of stimulation, with little to no proliferation observed after 3 days of stimulation (Figure 3C–E).

Figure 2.

Figure 2

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CML results before and after HCT in animals that lost their hematopoietic cell graft. Chimera antiself (squares) versus antidonor (triangles) versus third party (diamonds) performed in animals that received a haplo-HCT and eventually lost peripheral blood chimerism. (A) Before HCT; (B) 14 weeks after HCT.

Figure 3.

Figure 3

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Antidonor cellular proliferative responses after loss of donor hematopoietic cell graft. (A) Representative animal demonstrating early cellular proliferative responses after exposure to donor antigen, suggestive of sensitization. Note the early proliferative responses (day 3). This animal (AD) received 2 s.c. injections (1 month apart) of MHC-mismatched (AC) PBMCs without immunosuppression. (B) Representative responses of an AD animal that was never exposed to donor AC antigen. Naïve animals exhibited maximal proliferative responses after 5 days, stimulation in culture. (C–E) Cellular responses after graft loss in animals 19138 (C), 19139 (D), and 19140 (E). Assays were performed on week 33 (231 days post-HCT). Note the pattern similarity to (B) but not to (A). The number on top of the bars are the stimulation index.

Lack of Antidonor Antibody Responses after Graft Loss

We next examined whether antidonor antibody was induced after haplo-HCT in ITC-conditioned animals that lost chimerism yet regained MLR responses similar to a naïve animal. Serum from animals 19138–19140 was assessed for the presence of alloantibodies based on both binding and antibody-mediated cellular cytotoxicity to donor cells. No serum antibody binding to donor PBMCs was detected by flow cytometry (data not shown). Using the more sensitive antibody-mediated cellular cytotoxicity assay, we confirmed the absence of detectable antidonor antibodies in serum (Figure 4A).

Figure 4.

Figure 4

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Antidonor antibody assessment after HCT detected by complement-mediated antibody-dependent cellular cytotoxicity assay. (A) Serum samples from recipient animals 19138–19140 (Table 1; group 1) after loss of the hematopoietic cell graft and peripheral blood chimerism. Serum samples from animals 19138–19140 were analyzed before HCT and after loss of peripheral blood chimerism. The positive control serum used for the cytotoxic assays comes from an AD pig that was grafted with a CC skin graft that was subsequently rejected within 7 days. On day +21 after graft placement, this control animal also received CC PBMCs s.c. (as a form of reimmunization). The target cells used in the antibody cytotoxicity assay were AC (donor type) PBMCs. (B) Antibody cytotoxicity after a DLI and 2 injections (animals 19138 and 19139) or 1 injection (animal 19140) of donor AC PBMCs delivered s.c. in an attempt to immunize the animals not developing alloantibodies (Figure 1A; group 1). (C) Antibody cytotoxicity responses of 2 animals (19937 and 19938) that did not receive immunosuppression after receipt of their first dose of donor antigen in the form of cytokine-mobilized PBMCs (Figure 1A; group 2). As early as 1 week after i.v. immunization, animals 19937 and 19938 developed antidonor antibodies. (D) Animal 20312 received an RIC regimen without antigen exposure (no HCT), to examine whether the preparatory regimen alone can prevent development of alloantibodies. At 2 months after discontinuation of immunosuppression, animal 20312 received a donor skin graft, and antidonor antibodies were tested (Figure 1A; group 3).

Sensitized Cellular Responses Observed before and after Second Antigen Exposure via DLI

ITC-conditioned haplo-HCT recipients that lost peripheral blood chimerism did not appear to be sensitized to donor MHC, based on the lack of a detectable alloantibody response. Cellular proliferative responses to donor-matched PBMCs in these animals also appeared similar to the response seen in a naïve animal (not exposed to donor antigen). We next asked how these animals with apparently naïve alloresponses after haplo-HCT would react immunologically to another exposure to donor cells. We hypothesized that reexposure to donor antigen would induce responses similar to those observed in animals that had never been exposed to donor antigen. Animals 19138–19140 received their second antigen load i.v. in the form of a DLI. Naïve (unexperimented) animals that received PBMCs i.v. for the first time (and without immunosuppression) developed antidonor antibodies (animals 19937 and 19938; Figure 4C). After the DLI, animals 19138–19140 (who had lost their grafts) still demonstrated no antidonor alloantibodies (Figure 4B) despite exhibiting sensitized cellular proliferative responses (Figure 5A) after the DLI. In contrast, animals 19937 and 19938 (healthy animals without immunosuppression responding to a DLI) developed strong antidonor alloantibody responses while retaining normal (day 5) proliferative responses (Figure 5B and Table 1) to the donor after 2 i.v. infusions of donor cells. These proliferative responses were comparable to those seen in animals that had never been exposed to donor antigen (Figure 5C).

Figure 5.

Figure 5

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Cellular responses after the second exposure to donor antigen. (A) Sensitized MLR assays in animals 19138–19140 in cells harvested on days 3 and 5. MLRs were performed at 1 month after the second exposure of donor antigen (DLI). (B) MLRs of animals 19937 and 19938, which received i.v. PBMCs without immunosuppression. The third animal (naïve control) is an animal (from the farm) that was never exposed to donor antigen. The MLR of the naïve control was performed in parallel to MLRs of animals 19937 and 19938. All 3 animals are of the same haplotype as the experimental HCT recipients (animals 19138–19140). In brief, animals 19937 and 19938 received mobilized PBMCs without immunosuppression (antigen dose 1) and a subsequent DLI (antigen dose 2), all at equal doses as those delivered to experimental HCT animals. MLRs for animals 19937 and 19938 were performed at 1 month after DLI (after the second i.v. sensitization of donor antigen). (C) Animal 20312 received the ITC regimen without exposure to donor antigen. The MLR shown was performed at 1 month after a skin graft was placed (and rejected within 10 days). The graft was placed 2 months after discontinuation of immunosuppression. The number on top of the bars are the stimulation index.

We ruled out the possibility that the preparatory regimen could prevent the development of alloantibodies. Animal 20312 (Figures 4D and 5C) underwent the preparatory regimen and received donor antigen (in the form of a skin graft) only after immunosuppression had been discontinued for 2 months. The effects of the preparatory regimen did not prevent animal 20312 from developing humoral and cellular immune responses when donor skin antigen was given at 2 months after immunosuppression.

DISCUSSION

Animals that lost their HCT graft after ITC conditioning regained normal cellular proliferative and cytotoxic alloresponses to the donor without any detectable antidonor antibodies, similar to the alloresponses observed in animals that had never been exposed to donor antigen. Given these results, animals that lost chimerism might be expected to have immune responses to DLI comparable to those seen in naïve animals, but we have demonstrated that this assumption was unfounded. A second donor antigen exposure (without immunosuppression) in the form of a DLI in animals that lost chimerism induced an early cellular proliferative immune response different from that seen in animals that received a similar antigenic exposure without immunosuppression. Our findings do not directly prove that initial exposure to donor cells in animals that had lost their grafts induced an immune response leading to graft loss. However, our cellular and humoral assays indirectly suggest that this could be a possibility.

A second remarkable, clinically relevant finding is the observation that alloantibodies were never induced after graft loss or after DLI in ITC-conditioned haplo-HCT recipients. Antidonor alloantibodies are generally tested in the clinic to assess for sensitization [25,26]. Our data suggest that the absence of alloantibodies after graft loss might not be a reliable indicator of a lack of sensitization, and that decisions regarding retransplantation based on their presence may be misguided. This scenario must not be confused with patients who have been exposed to donor antigen before HCT (by, eg, transfusions) and who have already developed alloantibodies before HCT. The swine ITC HCT model is similar to transplantation scenarios in which patients who lack antidonor antibodies before HCT undergo RIC regimens and lose their donor hematopoietic cell graft. In these patients, the presence or absence of alloantibody is used as a diagnostic tool to assess the need for retransplantation.

The preparatory regimen might have had some impact in the immunologic outcomes observed. We previously reported that CD3-ITC delivered to animals that received the ITC regimen had a minimal impact on the level of donor T cell chimerism [8]. All animals that received the ITC regimen achieved stable mixed chimerism [8,9]. Recipients received fewer PBMCs (5–12 × 1010 cells instead of 15 × 1010 cells), because leukopheresis had to be discontinued early. The donors had an adverse response to the cytokines injected for stem cell mobilization (data not shown). It is possible that the delivery of a reduced (albeit still relatively high) cell dose in this very mild conditioning regimen, or the fact that the donors became sick, might have been a factor in the loss of chimerism and engraftment. Although our results are based on a relatively small cohort of animals, our data suggest that donor cells did not disappear passively and likely were rejected. The regain of antidonor cellular responses was associated with graft loss in these animals. It is possible that the incomplete depletion of host T cells in combination with a lower HCT dose might have been contributing factors in donor graft rejection [3,27,28].

Clinical implications extrapolated from these studies may affect decision making in patients requiring retransplantation. This may be important especially when considering the same donor, as in parent-to-child haplo-HCT [5]. On one hand, our results argue for the use of aggressive T cell–specific immune suppression (eg, anti-CD2 or antithymocyte globulin) to avoid the effects of cellular-sensitized responses, especially after the observed cellular responses on second antigen exposure. However, on the other hand, it can be argued that mild immunosuppression (with, eg, calcineurin inhibitors) could be sufficient, given that no sensitized responses were elicited right after graft loss. Further studies are needed to evaluate which of these 2 clinical approaches would be best.

Although induced alloantibody responses and early proliferative responses to donor cells are strong indicators of sensitization, as demonstrated in humans [29,30] and mice [31], our data indicate in a clinically relevant large-animal model that lack of antidonor alloantibody production does not reliably predict a lack of sensitization. These results in cases where sensitization after graft loss cannot be confirmed based on antibody or cellular responses suggest that immune suppression and/or T cell depletion may be warranted before retransplantation.

In conclusion, the present study confirms that an absence of sensitized cellular and humoral immune responses after graft loss does not necessarily indicate lack of immune response to donor cells. Our results have important clinical implications, and identification of better and more sensitive assays should be considered to identify sensitized patients before retransplantation.

Acknowledgments

We thank Aseda Tena and Joey Kurtz for their critical comments on the manuscript.

Footnotes

Authorship Statement: Raimon Duran-Struuck designed research, analyzed data, and wrote the manuscript. Abraham Matar performed research and analyzed data. Rebecca Crepeau performed research and analyzed data. Ashley Gusha analyzed data. Marian Schenk performed research. Isabel Hanekamp performed research and analyzed data. Vimukthi Pathiraja performed research. Thomas R. Spitzer wrote the manuscript. David H. Sachs designed research. Christene A. Huang designed research and wrote the manuscript.

Financial disclosure: This research was supported by National Center for Research Resources Grants K01RR024466 (to R.D.S.), R01AI084657, and P01CA111519. The authors have no conflicts of interest to disclose.

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