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TRANSPLANTATION
From the Division of Haematology, School of Clinical
Laboratory Sciences, University of Nottingham, Nottingham, United
Kingdom, and the Sir William Dunn School of Pathology, University of
Oxford, Oxford, United Kingdom.
Graft-versus-host disease (GVHD), a major complication after
allogeneic transplantation, can be abrogated by the Campath (anti-CD52) monoclonal antibody. The induction of acute GVHD requires host antigens
to be presented to donor T cells by antigen-presenting cells (APCs).
Recent evidence has suggested that only host APCs can interact with
donor T cells in the induction of GVHD. Because CD52 has been reported
to be expressed on DCs, we reasoned that pretransplant Campath-1G might
have a direct effect on circulating DCs in addition to any effects on
donor T cells. Using direct immunostaining, we demonstrated expression
of CD52 on DCs and that Campath-1G killed purified DCs in
vitro. In vivo Campath also depleted DCs. Twenty-four hours after the
first dose of Campath-1G, circulating DCs were reduced by a mean of
79% (range, 44%-96%). By day 0 after 5 doses of Campath-1G and
chemoradiotherapy conditioning, DCs became undetectable in 7 of 9 cases, whereas in 6 of 7 patients receiving conditioning therapy
without Campath-1G, host DCs were still detectable. The reconstitution
of circulating DCs after transplantation was not affected by Campath-1G
and in both groups DC1 (CD11c+) recovered more rapidly than
DC2 (CD11c High-dose chemoradiotherapy followed by the
allogeneic transplantation of hematopoietic stem cells from either bone
marrow (BM) or peripheral blood stem cells (PBSCs) is widely used in the treatment of malignant and nonmalignant hematologic diseases. However, allogeneic transplantation is frequently associated with the
development of acute graft-versus-host disease (GVHD), which is thought
to occur as a result of donor T-cell activation damaging host tissues
particularly affecting the liver, skin, and gastrointestinal tract.1,2
Current therapeutic approaches to the prevention of acute GVHD after
transplantation include immunosuppression with agents such as
cyclosporin and methotrexate. However, despite these measures GVHD
still remains a significant cause of morbidity and
mortality.3-5 Alternative strategies for the prevention of
acute GVHD have focused on the depletion of donor T cells from the
graft before infusion into the host.6-8 Although it has
been recognized for many years that T-cell depletion decreases the risk
of GVHD development,6,9 it also increases the risk of
leukemia relapse in chronic myeloid leukemia10 and graft
rejection.11 A number of different approaches to T-cell
depletion have been used including CD34+ cell
selection12,13 and ex vivo treatment of the graft with monoclonal antibodies. For this purpose, Campath-1M, which recognizes the CD52 antigen, has been widely used.9,14,15 CD52 is
highly expressed on human lymphocytes and monocytes and is a good
target for cell lysis by antibody with human
complement.16,17 Although ex vivo treatment with
Campath-1M is effective in preventing acute and chronic GVHD, the
clinical benefit in reduction in GVHD is offset by an increased risk of
graft rejection by residual host T cells and by an increased risk of
leukemic relapse in CML due to the loss of a graft-versus-leukemia
effect.10 Graft rejection in the setting of
T-cell-depleted grafts can be reduced by increasing the intensity of
pretransplant immunosuppression by using the rat IgG2b antibody
Campath-1G to deplete residual host T cells.15 In vivo
treatment with Campath-1G has also been used to prevent acute GVHD,
particularly following unrelated donor
transplantation.15,18-20 Originally, the Campath-1G was
given both before and after transplantation with the intention that the
pretransplant antibody would prevent rejection and the posttransplant
antibody would suppress GVHD by the elimination of donor T cells via
antibody-dependent cell cytotoxicity (ADCC).20 More
recently, pretransplant treatment alone has been reported to be highly
effective in the prevention of acute GVHD following either unrelated
transplantation21 or transplantation from HLA-matched
siblings.22 In fact, we found that pretransplant treatment
with Campath-1G from day Induction of acute GVHD requires host antigens to be presented to donor
T cells by antigen-presenting cells (APCs) in the context of the major
histocompatibility complex molecules. Recent evidence in a murine
allogeneic BMT model showed that only host CD11c+APCs can
present antigen to donor CD8+ T cells and that these cells
are critical in the induction of GVHD despite the presence of numerous
donor APCs,23 suggesting that donor DCs cannot efficiently
present host antigens necessary for the development of acute GVHD.
These results also suggested that the depletion of host APCs before
allogeneic transplantation might abrogate acute GVHD without requiring
prolonged posttransplant immunosuppression.23 DCs are
potent APCs that play an important role in initiating the immune
response, as a consequence of their potent ability to induce T-cell
activation and proliferation.24 Because the CD52 antigen
has been reported to be expressed on DCs,25 we reasoned
that the administration of in vivo Campath to the recipient prior to
allogeneic transplantation might result in the depletion of host DCs
prior to the infusion of donor cells and that this could contribute to
the reduction of acute GVHD.21,22
We have therefore assessed the effect of in vivo pretransplant
Campath-1G on circulating DCs before and after the completion of
conditioning chemoradiotherapy. We have also analyzed the effect of
pretransplant Campath-1G on the origin and kinetics of reconstitution of DCs after transplantation.
Samples
For patients receiving pretransplant Campath-1G, Campath-1G was given
at a dose of 10 mg daily from day Patients who did not receive Campath-1G received either a matched
sibling allogeneic transplant with cyclophosphamide 60 mg/kg for 2 doses and fractionated TBI (12 Gy; n = 3) or an autologous transplant
with BEAM conditioning (n = 4).
In both groups peripheral blood DCs were enumerated before
starting Campath/conditioning, and on day 0 following completion of
conditioning (with or without Campath therapy). In the Campath group
peripheral blood DCs were also measured on day Monoclonal antibodies
DC enumeration in peripheral blood Peripheral blood was stained without further separation to minimize selective loss. Erythrocytes were lysed after staining using Ortho-mune lysing reagent (Ortho Diagnostic Systems, Raritan, NJ) according to the manufacturer's instruction. DCs were identified as negative for the cocktail antibodies of phycoerythrin (PE) conjugated specific for lineage markers on T cells (CD3), B cells (CD19, CD20), monocytes (CD14), neutrophils (CD16), hematopoietic stem cells (CD34), and natural killer (NK) cells (CD16, CD56) and positive for FITC-conjugated anti-HLA-DR. DC1 and DC2 were identified using allophycocyanin-conjugated anti-CD11c. Three- color analysis was performed using a FACSCalibur flow cytometer (Becton Dickinson). The number of total white blood cells (WBCs) in the samples was determined using an automated WBC counter. The absolute number of total DCs and their subsets was calculated from the WBC count multiplied by the proportion of each subset among the WBC, as determined by flow cytometric analysis.CD52 expression by DCs To identify the expression of the CD52 antigen by DCs, a peripheral blood sample from a healthy volunteer donor was used. FITC-conjugated Campath-1G was used to stain the DC population together with PE-conjugated cocktail antibodies (anti-CD3, CD14, CD16, CD19, CD20, CD34, and CD56), allophycocyanin-conjugated anti-CD11c, and cychrome (PEcy5)-conjugated anti-HLA-DR. Four-color analysis was performed using a FACSCalibur flow cytometer. The expression of CD52 on T cells was also determined using Campath-1G as a positive control. A FITC-conjugated rat IgG2b was also used as an isotype negative control.The effect of Campath-1G on DCs in vitro Mononuclear cells were separated from normal peripheral blood by density gradient centrifugation technique using Histopaque (Sigma, St Louis, MO). Cells were stained with PE-conjugated anti-CD3, CD14, CD16, CD19, CD20, CD34, and CD56 and FITC-conjugated anti-HLA-DR. The HLA-DR+, lineage-minus (HLA-DR+/Lin ) cells were sorted using
EpicALTRA High-Performance Cell Sorting System (Beckman Coulter,
Fullerton, CA). Purified DCs were incubated for 20 minutes with 75 µg/mL Campath-1G in complete RPMI-HEPES supplemented with 10%
heat-inactivated fetal calf serum (First Link, Wolverhampton, United
Kingdom). Recombinant human granulocyte-macrophage colony-stimulating factor (GM-CSF; 10 ng/mL; (Pharmingen) and 50 ng/mL
recombinant human interleukin (IL)-3 (R & D Systems, Abingdon, United
Kingdom) were added to the culture medium because GM-CSF and
IL-3 are essential for the survival of DCs.26 Rat IgG2b
(75 µg/mL; Serotec) was also used as a control antibody in culturing
purified DCs without Campath-1G. Twenty percent of autologous serum was
added and cells were cultured for 24 hours. DCs were also cultured with
Campath-1G without autologous serum to determine whether complement is
needed for the induction of cell death. Cells were then washed twice
with phosphate-buffered saline (PBS) and resuspended with PBS. Cell
suspension was incubated with 20 µg/mL (final concentration) of
7-amino-actinomycin D (7-AAD, Sigma) at 4°C for 20 minutes in the
dark and analyzed for the numbers of live cells using FACSCalibur flow
cytometer according to the previous studies by Philpott et al
27
DC separation for microsatellite polymerase chain reaction Peripheral blood mononuclear cells (PBMCs) were separated from EDTA blood by a density gradient centrifugation technique. PBMCs were stained with PE-conjugated anti-CD3, CD14, CD16, CD19, CD20, CD34, and CD56. Cells were then labeled with anti-PE magnetic microbeads (Miltenyi Biotec, Bergisch Gladbach, Germany). Labeled cells were depleted on a high-gradient magnetic separation column (MiniMACS, Miltenyi Biotec). The negative fraction was then stained with anti-HLA-DR magnetic microbeads and positively selected in the same way. The immunomagnetic separation was performed according to the manufacturer's instructions. The entire procedure lasted 4 to 6 hours, during which cells were kept at 4°C. DCs were resuspended in 200 µL PBS. DNA was extracted using a QIAamp DNA Blood Mini Kit (Qiagen, Hilden, Germany). DNA samples were kept at 20 °C until
microsatellite polymerase chain reaction (PCR) was performed.
Microsatellite PCR Sequential samples were assessed for donor chimerism status using fluorescent microsatellite PCR for Rb1 or D6S264. One or both of these markers had previously been shown to be informative by analyzing DNA from donor-recipient pairs before transplantation. Forward (F) and reverse (R) primers specific for Rb128 and D6S26429 were Rb1(F) 5'CTCCTCCCTACTTACTTGT, Rb1(R) 5'AATTAACAAGGTGTGGTGGTACACG, D6S264(F) 5'AGCTGACTTTATGCTGTTCCT, D6S264(R) 5'TTTTCCATGCCCTTCTATCA with the forward primer being 5'end-labeled with FAM. Typically, 100 ng DC DNA was amplified in a 25-µL reaction volume using 1 U Amplitaq Gold (Applied Biosystems, Foster City, CA), one times PCR buffer II, and 0.125 mM dNTPs (Amersham Life Science, Cleveland, OH). Rb1 PCR had 35 cycles with annealing at 55°C and 2.0 mM MgCl2. D6S264 amplifications were conducted in 1.5 mM MgCl2 for 30 cycles and annealing at 55°C.Analysis of PCR products Fluorescent PCR products were diluted in molecular grade water (1/5 or 1/10) and electrophoresed through 4.2% sequencing gels (Thistle Scientific, Coatbridge, United Kingdom) on an ABI Prism 377 DNA sequencer (Applied Biosystems). Electrophoresis was for 5 hours at 40 W and 48°C with a GS-500 marker (Applied Biosystems) in each lane. Data were analyzed using Genescan Analysis software and quantitation of percentage of donor chimerism was as described by Miflin et al.30Statistics Statistical analysis (unpaired t test) was performed using the statistics package (GraphPad Prism software, West Wycombe, United Kingdom).
DC enumeration in peripheral blood of healthy volunteers To establish the reference range of DCs in peripheral blood, peripheral blood samples from 10 healthy men and 9 healthy women, aged 23 to 48, were stained. DCs were identified as HLA-DR+ and HLA-DR for lineage markers of T cells, monocytes, NK
cells, granulocytes, B cells, and CD34+ progenitor cells
(Figure 1A). The 2 subsets of DCs were
analyzed using antibody against the adhesion molecule CD11c. DC1 were
identified as CD11c+ cells, whereas DC2 were negative for
CD11c (Figure 1B,C). The mean absolute number of DCs in normal
peripheral blood using this method was
27.7 ± 13.3 × 106/L (mean ± 1 SD) with DC1 of
15.4 ± 7.5 × 106/L and DC2 of
12.3 ± 7.5 × 106/L.
CD52 expression on DCs To determine whether peripheral blood DCs express CD52 antigen, normal peripheral blood was stained as described. The CD52 antigen was found to be expressed on both DC1 and DC2 (Figure 1D,E). Studies of 5 healthy controls showed that a median of 81% of DC1 expressed the CD52 antigen (range, 59%-97%), whereas a median of 64% of DC2 expressed CD52 (range, 46%-91%). In this experiment T cells were also stained for CD52 expression as the positive control for this antibody (Figure 1F).Campath-1G causes DC death in vitro The effects of Campath-1G on DCs in vitro were studied by culturing purified DCs with Campath-1G. Staining with 7-AAD was used to analyze the numbers of dead, apoptotic, and live cells as shown in Figure 2. The proportion of dead, apoptotic, and live cells was compared in the cells cultured with Campath-1G and the control cells cultured with rat IgG2b. The results showed that after incubating purified with Campath-1G for 24 hours in the presence of autologous complement, the proportion of surviving cells cultured with Campath-1G (median = 36.5%) was significantly lower than the proportion of live cells after culturing with control antibody (median = 76.2%; n = 4, P = .002). Cells were also cultured with Campath-1G without autologous serum and we found that there was no difference in the proportion of surviving cells between cells cultured with control antibody and cells cultured with Campath-1G alone (median = 71.6%). This suggested Campath-1G could induce complement-dependent cytotoxicity in DCs.
Pretransplant Campath causes rapid depletion of circulating DCs The effects of pretransplant Campath-1G on circulating DCs were analyzed using flow cytometry for DC enumeration at different time points before transplant as described. In the Campath-treated group, pretransplant Campath-1G (10 mg) depleted circulating DCs by a mean of 79% (range, 44%-96%; n = 9) 24 hours after the first dose of antibody and before starting any chemoradiotherapy conditioning (Figure 3 and Table 1). At this time point the total number of circulating DCs had fallen from 12.6 ± 7.3 × 106/L to 2.4 ± 2.3 × 106/L. At day 0, following the completion of Campath-1G and the conditioning therapy, circulating DCs in the Campath group were undetectable in 7 of 9 cases (Table 1). In contrast, on day 0 peripheral blood DCs in the non-Campath group were depleted but could still be detected in 6 of 7 cases (P = .029).
DC reconstitution in peripheral blood is not delayed by pretransplant Campath Because Campath-1G administered before transplantation has been shown to circulate at the time of marrow infusion,22 it was possible that the regeneration of donor DCs would be delayed. We therefore examined the kinetics of circulating DC reconstitution at various time points up to 1 year after transplantation. There was a slow recovery of both DC1 and DC2 subsets in both patient groups and the levels of both total DC numbers and DC subsets remained depressed up to 12 months after transplantation (Figure 4). However, comparison between the 2 groups of patients suggested that the posttransplant recovery of both DC1 and DC2 was not affected by pretransplant Campath (Figure 4).
Peripheral blood DCs regenerating after transplant are of donor origin Peripheral blood DCs were isolated after transplant and analyzed for chimerism. At 8 weeks after transplant, 4 of 4 patients who had received pretransplant Campath had 100% donor DCs by variable number of tandem repeats (VNTR) analysis. Two of 2 patients with no pretransplant Campath treatment also had 100% donor DCs. However, at 12 weeks after transplant, one patient in the non-Campath group had mixed chimerism with 90% of DCs being of donor origin. This patient later had a relapse at 5 months, whereas at this time point all patients in the Campath group had DCs of donor origin.
The induction of acute GVHD requires host antigen to be presented
in the context of HLA to donor T cells by APCs. Recent reports have
suggested that host APCs may be critical to donor CD8+
T-cell activation and the induction of acute GVHD,23
suggesting that depletion of host APCs before allogeneic
transplantation might abrogate acute GVHD. Because pretransplant
serotherapy with Campath-1G has been highly effective in the prevention
of both acute GVHD and graft rejection following either unrelated donor or sibling donor transplantation,21,22 we reasoned that
this effect might be mediated by a direct effect of Campath on host APCs. In this study, we focused on the effect of Campath-1G on DCs,
which are one of the most potent APCs. The CD52 antigen has been
reported to be expressed on CMRF-56+ peripheral
blood DCs25 and we initially studied the expression of
CD52 on circulating DCs. These studies confirmed that the majority of
peripheral blood DCs, enumerated as
HLA-DR+/Lin Studies of the effect of Campath-1G therapy on peripheral blood DCs demonstrated that pretransplant Campath reduced the number of circulating DCs by a mean of 79% (range, 44%-96%) after one dose. At day 0, following the completion of 5 doses of Campath-1G and the conditioning therapy, peripheral blood DCs were undetectable in 7 of 9 patients. In contrast, peripheral blood DCs in 6 of 7 patients receiving conditioning therapy for BMT without Campath could still be detected on day 0. The data demonstrate that in vivo Campath-1G causes rapid depletion of peripheral blood DCs and in conjunction with high-dose chemotherapy or chemoradiotherapy totally cleared DCs from the peripheral blood in the majority of patients. Furthermore, we also showed a direct in vitro effect of Campath-1G in killing purified DCs by complement-dependent cytotoxicity suggesting that the in vivo effect of Campath-1G was a direct effect and not merely due to pooling of DCs into the extravascular space. These effects of Campath on DCs may in part explain the lack of severe acute GVHD observed using pretransplant Campath.18,19,21 However, our studies have been confined to the effects on circulating DCs and we have not studied whether CD52 is expressed on tissue DCs or whether in vivo Campath-1G depletes these cells as efficiently as peripheral blood DCs. Furthermore, other mechanisms are probably involved in the suppression of acute GVHD by pretransplant Campath-1G because we have previously demonstrated that the antibody has a half-life of approximately 13 hours and we found that at the time of marrow infusion plasma levels of Campath were sufficient to deplete donor T cells by ADCC in the majority of cases.22 Thus we cannot be certain whether depletion of host DCs contributes to the prevention of acute GVHD by pretransplant Campath antibodies because it is impossible to separate this effect from T-cell depletion of the graft by residual Campath-1G. As well as effects on host DCs we considered whether Campath antibodies could delay donor DC reconstitution after transplant. Our results show that there is a slow recovery of both DC1 and DC2 following allogeneic transplantation. However, there were no significant differences between the Campath and non-Campath groups (Figure 4), which suggested that the regeneration of DCs after transplant was not adversely affected by pretransplant Campath-1G. Because host DCs persisting after transplant might induce acute GVHD, we considered it important to determine the origin of DCs regenerating after transplantation. In all patients who received pretransplant Campath-1G, peripheral blood DCs were totally of donor origin. None of these patients had evidence of GVHD at the time of analysis. We have not specifically studied DC chimerism in patients undergoing transplantation with established acute GVHD, but early reports from others suggest that this is associated with a pattern of mixed chimerism,35 which is in keeping with the role of residual host DCs in mediating GVHD. In summary, we found that peripheral blood DCs express the CD52 antigen and that Campath-1G induces DC death on in vitro culture. In vivo Campath when used in combination with conditioning therapy totally depletes circulating host DCs, which contrasts with conditioning regimens not including Campath where the majority of patients still had circulating host DCs at the time of transplantation. These observations may therefore in part explain the low incidence of acute GVHD in patients receiving pretransplant Campath-1G. Furthermore, these results support the concept that antibodies that specifically target host DCs may be of value in the prevention of acute GVHD without the prolonged immunosuppression caused by T-cell depletion.
The authors would like to thank Dr Adrian Robins and Dr Heather Judge from the Division of Molecular and Clinical Immunology, School of Clinical Laboratory Sciences, University of Nottingham, Nottingham, United Kingdom, for their kind help in purifying dendritic cells.
Submitted May 31, 2001; accepted November 21, 2001.
The publication costs of this article were defrayed in part by page charge payment. Therefore, and solely to indicate this fact, this article is hereby marked "advertisement" in accordance with 18 U.S.C. section 1734.
Reprints: Nigel H. Russell, Haematology Department, Nottingham City Hospital, Hucknall Rd, Nottingham NG5 1PB, United Kingdom; e-mail: nigel.russell{at}nottingham.ac.uk.
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© 2002 by The American Society of Hematology.
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A. Y. L. Ho, A. Pagliuca, M. Kenyon, J. E. Parker, A. Mijovic, S. Devereux, and G. J. Mufti Reduced-intensity allogeneic hematopoietic stem cell transplantation for myelodysplastic syndrome and acute myeloid leukemia with multilineage dysplasia using fludarabine, busulphan, and alemtuzumab (FBC) conditioning Blood, September 15, 2004; 104(6): 1616 - 1623. [Abstract] [Full Text] [PDF]
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F. F. Fagnoni, B. Oliviero, G. Giorgiani, P. De Stefano, A. Deho, C. Zibera, N. Gibelli, R. Maccario, G. Da Prada, M. Zecca, et al. Reconstitution dynamics of plasmacytoid and myeloid dendritic cell precursors after allogeneic myeloablative hematopoietic stem cell transplantation Blood, July 1, 2004; 104(1): 281 - 289. [Abstract] [Full Text] [PDF]
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 M. Y. Mapara and M. Sykes Tolerance and Cancer: Mechanisms of Tumor Evasion and Strategies for Breaking Tolerance J. Clin. Oncol., March 15, 2004; 22(6): 1136 - 1151. [Abstract] [Full Text] [PDF]
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R. D. Faulkner, C. Craddock, J. L. Byrne, P. Mahendra, A. P. Haynes, H. G. Prentice, M. Potter, A. Pagliuca, A. Ho, S. Devereux, et al. BEAM-alemtuzumab reduced-intensity allogeneic stem cell transplantation for lymphoproliferative diseases: GVHD, toxicity, and survival in 65 patients Blood, January 15, 2004; 103(2): 428 - 434. [Abstract] [Full Text] [PDF]
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 M. Zoller Tumor Vaccination after Allogeneic Bone Marrow Cell Reconstitution of the Nonmyeloablatively Conditioned Tumor-Bearing Murine Host J. Immunol., December 15, 2003; 171(12): 6941 - 6953. [Abstract] [Full Text] [PDF]
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 A. M. Woltman and C. van Kooten Functional modulation of dendritic cells to suppress adaptive immune responses J. Leukoc. Biol., April 1, 2003; 73(4): 428 - 441. [Abstract] [Full Text] [PDF]
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G. Ratzinger, J. L. Reagan, G. Heller, K. J. Busam, and J. W. Young Differential CD52 expression by distinct myeloid dendritic cell subsets: implications for alemtuzumab activity at the level of antigen presentation in allogeneic graft-host interactions in transplantation Blood, February 15, 2003; 101(4): 1422 - 1429. [Abstract] [Full Text] [PDF]
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 G. Mufti, A. F. List, S. D. Gore, and A. Y.L. Ho Myelodysplastic Syndrome Hematology, January 1, 2003; 2003(1): 176 - 199. [Abstract] [Full Text] [PDF]
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D. G. Maloney, B. M. Sandmaier, S. Mackinnon, and J. A. Shizuru Non-Myeloablative Transplantation Hematology, January 1, 2002; 2002(1): 392 - 421. [Abstract] [Full Text]
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Abstract
Introduction
