Role for CD4+CD25+ T Cells in Inhibition of Graft Rejection by Extracorporeal Photopheresis

CARDIAC REJECTION

Role for CD4ⴙCD25ⴙ T Cells in Inhibition of Graft Rejection by Extracorporeal Photopheresis James F. George, PhD, Christie W. Gooden, MD, Wing Hong Guo, MD, and James K. Kirklin, MD Background: Extracorporeal photopheresis (ECP) is used to treat recurrent severe rejection in clinical heart and lung recipients. The mechanisms of the salutary effects of ECP are poorly understood, but appear to involve regulation of T-cell–mediated alloreactive responses, possibly by induction of regulatory T cells. We created a mouse model of ECP to determine the effects of ECP on T-cell responses in vivo and the contribution of CD4⫹CD25⫹ T cells. Methods: In this study, 1 ⫻ 107 splenocytes were treated with 8-methoxypsoralen (8-MOP, 200 ng/ml), followed by ultraviolet A (UVA) irradiation (2 J/cm2, 350 nm), and then injected intravenously into syngeneic mice. Thirty minutes later, the treated animals received heterotopic cardiac allografts with no immunosuppression. Treated graft recipients were analyzed to determine the effect of ECP on graft survival, deletion of allospecific T cells, and the frequency and in vivo suppressive activity of CD4⫹CD25⫹ T cells. Results: ECP extends cardiac allograft survival in at least two different strain combinations. For CBA/Ca recipients of C57BL/6 allografts, median survival time (MST) in ECP-treated animals was 16 days vs 10 days in graft recipients treated with cells exposed only to 8-MOP (p ⫽ 0.04). The frequency of splenic CD4⫹CD25⫹ cells expressing FoxP3⫹ increased 2-fold in ECP-treated CBA/Ca mice (82.6 ⫾ 5.2%, n ⫽ 4) relative to untreated mice (44.9 ⫾ 4.5%, n ⫽ 4, p ⬍ 0.001). Adoptive transfer of 3 ⫻ 105 sorted CD4⫹CD25⫹ splenocytes from ECP-treated graft recipients to untreated cardiac allograft recipients 30 minutes after transplantation resulted in extended graft survival compared with animals that received the same number of CD4⫹CD25⫹ splenocytes from cardiac allograft recipients not treated with ECP (MST: 24 days vs 13 days, respectively, p ⫽ 0.001). Analyses of 5,6-carboxyfluorescein-succinimidyl-ester (CFSE)-labeled H-2Kb–specific T cells in the spleen and lymph node showed no evidence of peripheral deletion after ECP treatment. Conclusions: ECP extends graft survival even in fully histoincompatible strain combinations with no immunosuppression. It increases the frequency of FoxP3⫹CD4⫹CD25⫹ splenic T cells, and its effects can be transferred to untreated recipients using minimal numbers of CD4⫹CD25⫹ T cells, indicating that CD4⫹CD25⫹ T cells may play a key role in the immunomodulatory effects of ECP. J Heart Lung Transplant 2008;27: 616 –22. Copyright © 2008 by the International Society for Heart and Lung Transplantation.

Extracorporeal photopheresis (ECP) is approved for treatment of cutaneous T-cell lymphoma, and has demonstrated benefit for graft-vs-host disease (GVHD),1 a number of T-cell– dependent chronic inflammatory diseases2,3 and recurrent solid-organ allograft rejection.4 – 6

From the Department of Surgery, University of Alabama at Birmingham, Birmingham, Alabama. Submitted September 20, 2007; revised January 29, 2008; accepted February 17, 2008. Supported in part by a grant from Therakos Corp., Exton, Pennsylvania (to J.G.), and by the American Heart Association (Grant No. 0655318B). Reprint requests: James F. George, PhD, Department of Surgery, University of Alabama at Birmingham, 1530 Third Avenue South, Birmingham, AL 35294. Telephone: 205-934-4261. Fax: 205-975-0085. E-mail: [email protected] Copyright © 2008 by the International Society for Heart and Lung Transplantation. 1053-2498/08/$–see front matter. doi:10.1016/ j.healun.2008.02.015

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ECP is performed in a patient-connected, closed-loop system in which whole blood is removed from the patient and leukocytes are isolated by centrifugation. The isolated leukocytes are treated with 8-methoxypsoralen (8-MOP), followed by exposure to ultraviolet A (UVA) light (350 nm), resulting in formation of covalent bonds between the 8-MOP and pyrimidine bases in the cellular DNA, causing induction of apoptosis.7,8 The cells are immediately returned to the patient after the UVA treatment. During ECP treatment, approximately 1.5 liters of blood is processed, resulting in the infusion of an intravenous bolus of cells in which the apoptosis program has been activated. We have shown that this treatment reduces the frequency of rejection in cardiac allograft recipients at high risk for rejection with hemodynamic compromise.4 The purpose of this study was to determine the effect of ECP on unmodified graft rejection, the frequency and function of CD4⫹CD25⫹ T cells, and the frequency of

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alloreactive T cells. We developed a murine model of ECP and showed that ECP inhibits solid-organ allograft rejection in vivo by non-deletional inhibition of graftspecific T-cell responses, and that the mechanism of inhibition involves CD4⫹CD25⫹ T cells capable of actively suppressing allograft rejection in vivo.

effect on graft survival was determined by comparison of graft survival times to allograft recipients that had received adoptively transferred cells from animals that had received cells treated with 8-MOP only, and to allograft recipients that were not treated with ECP or adoptive transfer.

METHODS Animals C57BL/6J (H-2b, Stock No. 000664), BALB/cJ (H-2d, Stock No. 000651) and CBA/CaJ (H-2k, Stock No. 000654) mice were obtained from Jackson Laboratories (Bar Harbor, ME). CBA/Ca transgenic mice (hereafter called BM3; kindly provided by Dr Andrew Mellor, Medical College of Georgia) were created by microinjecting CBA/Ca oocytes with T-cell receptor (TCR)-␣ and -␤ gene constructs from the H-2Kb–specific T-cell clone BM3.3.9 This clone recognizes the octamer PBM1 bound to the MHC Class I molecule, Kb.10 CBA/CaJ or C57BL/6J mice received abdominal heterotopic cardiac allografts from age-matched C57BL/6J or BALB/cJ donors, respectively, using previously described surgical techniques.11 The mice were examined daily and function of the heart allograft was confirmed by palpation through the abdominal wall. No immunosuppressive agents were used.

Isolation of Intracardiac Lymphocytes Donor hearts were diced into 1-mm pieces and minced between two frosted glass slides. The resulting slurry was placed in 100 U/ml collagenase (Boehringer Mannheim, Mannheim, Germany). After vigorous pipetting, the cell suspension was removed and digestion was stopped by addition of ethylene-diamine tetraacetic acid (EDTA) to 10 mmol/liter. The remaining fragments were digested further in collagenase (400 U/ml) at 37°C for 1 hour. The fragments were minced again, washed and discarded. The cell suspension was then centrifuged through a Nycodenz (Nycomed, Oslo) gradient (1.077 g/ml) for 15 minutes at 1,700g at 4°C. The cells were counted using a hemacytometer and analyzed by flow cytometry.

Extracorporeal Photopheresis Mice were treated with ECP 30 minutes prior to receiving a heterotopic cardiac allograft, as described earlier. ECP was performed by injecting the mice via the tail vein with 1 ⫻ 107 syngeneic splenocytes that had been incubated in complete medium containing 200 ng/ml 8-methoxypsoralen (Uvadex [methoxsalen]; Therakos, Exton, PA) for 2 minutes and then exposed to UVA (350 nm) irradiation at a dose of 2 J/cm2. Control ECP treatments consisted of the aforementioned procedure, but without UVA irradiation. Adoptive Transfer of Lymphoid Cells From ECP-treated Graft Recipients At 3 days post-transplantation, ECP-treated graft recipients were euthanized and the spleens were removed and then disaggregated into single-cell suspensions. Erythrocytes were removed by treatment with ammonium chloride followed by two washes in phosphate-buffered saline (PBS). CD4⫹CD25⫹ T cells were isolated using a magnetic sorting kit for CD4⫹CD25⫹ splenic T cells (Miltenyi Biotec, Auburn, CA) according to the manufacturer’s instructions. Purity of the isolated CD4⫹CD25⫹ cells was consistently ⬎85%. The CD4⫹CD25⫹ T cells were then injected via the tail vein (3 ⫻ 105 cells in 0.2 ml PBS) into syngeneic, untreated cardiac allograft recipients 30 minutes before the transplant surgery. Graft survival in the recipients of the adoptively transferred cells was determined by abdominal palpation. The

Antibodies, Fluorescent Probes and Flow Cytometry Analyses Antibodies specific for CD3 (145-2C11), CD4 (GK1.5), CD11b (M1/70), CD19 (1D3), CD25 (PC61.5) and CD8 (53-6.7) were purchased from Ebioscience (San Diego, CA). Immunofluorescence analysis was performed as previously described.12 FoxP3 (forkhead box P3)-bearing cells were visualized by intracellular staining with phycoerythrin-conjugated anti-FoxP3 antibodies (Clone FJK-16s; Ebioscience). After surface staining, the cells were fixed and permeabilized using fixation/permeabilization buffer (Ebioscience) for 30 minutes, followed by staining with 0.5 ␮g per 1 ⫻ 106 cells of FJK-16s antibodies for 30 minutes. Cells were washed twice in permeabilization buffer (Ebioscience) and analyzed by flow cytometry. CFSE (CFDA-SE; Molecular Probes, Inc., Carlsbad, CA) was also obtained and used to label BM3 splenocytes prior to injection as previously described.12 Statistical Analysis Groups were compared using Kaplan–Meier and logrank statistics. Student’s t-test was used for parametric non–time-related comparisons. p ⬍ 0.05 was considered statistically significant. RESULTS Apoptosis Induction in ECP-treated Splenocytes CBA/Ca splenocytes were prepared by standard means, incubated for 2 minutes in 200 ng/ml 8-MOP, and exposed to UVA irradiation (350 nm) at 2 J/cm2. The cells were washed twice, incubated in culture medium for 20 hours, and then stained with antibodies specific for CD3, CD11b and CD19. Staining with Annexin-V

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Figure 1. Induction of apoptosis in vitro in CBA/CaJ splenocytes 20 hours after treatment with 8-MOP and UVA irradiation. Plots are representative results of four independent experiments. Hatched bars: no 8-MOP or UVA irradiation; open bars: treatment with UVA irradiation only; solid bars: treatment with both 8-MOP and UVA irradiation. Each plot shows the proportion of cells positive for both 7-AAD and Annexin-V (7AAD⫹/AnnV⫹) or positive for Annexin-V only (7AAD⫺/AnnV⫹) as measured by flow cytometry. Cells in each plot are gated for the antigen indicated at the top. In all cases, maximal induction of apoptosis occurred in cells treated with both 8-MOP and UVA.

and 7-AAD was used to determine the proportion of cells that were late apoptotic or dead (Annexin-V⫹/7AAD⫹) and early apoptotic (Annexin-V⫹/7-AAD⫺). Figure 1 shows that cells undergoing the early stages of apoptosis (7-AAD⫹/Annexin-V⫺) were 2- to 3-fold more abundant 20 hours after treatment with UVA and 8-MOP in comparison to no treatment or to treatment with UVA alone. Similar proportions were obtained for cells that were dead or undergoing later stages of apoptosis (7-AAD⫹/Annexin-V⫹). All subsequent ECP treatments were performed using cells treated with UVA and 8-MOP as noted previously, with the exception that they were infused immediately after irradiation, as is performed clinically. ECP Prolongs Survival of Murine Heterotopic Cardiac Allografts ECP-treated CBA/Ca recipients of C57BL/6 cardiac allografts rejected cardiac allografts more slowly than graft recipients infused with 1 ⫻ 107 splenocytes that were treated with 8-MOP alone (Figure 2a; p ⫽ 0.044). Similar results were obtained using a different strain combination. C57BL/6J recipients of BALB/c cardiac allografts rejected the grafts more slowly when treated with ECP in comparison to the controls (Figure 2b; p ⫽ 0.005). Therefore, ECP modulates the immune response to the graft strongly enough to delay graft rejection, even in the absence of immunosuppression. In C57BL/6J mice, the infusion of 1 ⫻ 107 ECP-treated BALB/cJ splenocytes was the approximate minimum number that conferred maximal protection. Half that number resulted in graft survival that was significantly decreased relative to animals receiving 1 ⫻ 107 ECP-treated cells. Infusion of up to 4 ⫻ 107 ECPtreated splenocytes did not significantly increase graft survival (Table 1). Effect of ECP on Graft-specific Alloreactive T Cells In Vivo One day prior to transplantation with C57BL/6J hearts and ECP treatment, CBA/CaJ graft recipients were in-

jected intravenously with 1 ⫻ 107 CFSE-labeled syngeneic splenocytes from BM3 mice, which contain T cells of a known anti-donor MHC specificity. As seen in Figure 3c and e, no decrease in the frequency of

Figure 2. Kaplan–Meier depictions of cardiac allograft survival in mice that received fully allogeneic heterotopic cardiac allografts 30 minutes after treatment with 1 ⫻ 107 syngeneic splenocytes treated with 8-MOP and UVA irradiation (see Methods). (a) Survival of C57BL/6J hearts in CBA/CaJ recipients infused with ECP-treated splenocytes from normal, non-transplanted CBA/CaJ mice. Each group consisted of 6 animals (p ⫽ 0.044, log-rank test). (b) Survival of BALB/cJ hearts in C57BL/6J recipients infused with ECP-treated splenocytes from normal, non-transplanted C57BL/6J mice. In each experiment, control groups were infused with 1 ⫻ 107 syngeneic splenocytes that were treated with 8-MOP, but no UVA irradiation. Each group consisted of 6 animals (p ⫽ 0.005, log-rank test).

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Table 1. Effect of Dose of ECP-Treated Splenocytes on Graft Survival Dosea 0.0 0.5 1.0 2.0 4.0 a

Median survival 7.5 10.0 12.5 12.0 10.0

p-valueb 0.005 0.022 — 0.517 0.153

n 6 6 6 6 6

Relative to a dose of 1 ⫻ 107 ECP-treated splenocytes. b Log-rank test in comparison to a dose of 1 ⫻ 107 ECP-treated splenocytes.

CFSE⫹CD8⫹ T cells was observed at Day 3 post-transplantation in the spleen and lymph nodes relative to animals that were not treated with ECP. There was little evidence of strong proliferative activity in the peripheral lymphoid organs as indicated by CFSE dye dilution (Figure 3d and f). A decrease in the frequency of CD8⫹CFSE⫹ cells was observed in the donor hearts of ECP-treated animals relative to controls, which was statistically significant by Day 3 post-transplantation (Figure 3a and b; p ⫽ 0.03). In the control animals, we observed a trend toward a linear increase in the proportion of CD8⫹CFSE⫹ graft-infiltrating cells on Days 1 to 3 post-transplantation. No such increase was noted in the ECP-treated animals, suggesting that graft infiltration by T lymphocytes was reduced (Figure 3a). Comparison of the absolute numbers of CD3⫹ T cells recovered from the donor hearts at the time of killing supports this conclusion. By Day 3 post-transplantation, the absolute numbers of recovered T cells was reduced by at least 3-fold in the ECP-treated mice compared with the control mice (Figure 4; p ⬍ 0.01). Therefore, ECP treatment of cardiac allograft recipients does not cause peripheral deletion of graft-reactive T cells. CD4ⴙCD25ⴙ T-cell Frequency and Activity in ECP-treated Animals Among CD4⫹CD25⫹ T cells, the proportion of FoxP3⫹ cells was significantly increased in the spleens of CBA/ CaJ ECP-treated animals at Day 3 post-transplantation (from 44.1 ⫾ 4.5% to 82.6 ⫾ 5.2%, p ⬍ 0.0001, Student’s t-test), a time-point that corresponds with the observations of reduced T-cell frequency in the transplanted heart. Observations of the frequency of CD4⫹CD25⫹ cells (Tregs) in transplant recipients at 60 days post-transplantation, long after all but one graft in an ECP-treated animal had been rejected, showed that the increase in the frequency of Tregs in ECP-treated animals had disappeared (68.1 ⫾ 17.6 in treated animals vs 75.4 ⫾ 13.9 in the controls, p ⫽ 0.49). These proportions are similar to those found in normal, untransplanted mice (65.2 ⫾ 5.11% of CD4⫹CD25⫹ cells). The correlation of the change in FoxP3⫹ T-cell frequency with ECP treatment led us to test the prediction that adoptive transfer of CD4⫹CD25⫹ cells from ECP-

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treated animals into untreated recipients would also convey the effects of ECP to the untreated animal. CBA/CaJ recipients of C57BL/6J cardiac allografts were treated with ECP as described earlier. At 3 days post-transplantation, these animals were killed and CD4⫹CD25⫹ cells were isolated from the spleen by magnetic sorting. Then 3 ⫻ 105 CD4⫹CD25⫹ cells were injected via the tail vein into untreated cardiac allograft recipients of the same strain combination 30 minutes prior to transplantation. Kaplan– Meier plots of cardiac allograft survival in these animals showed that the transfer of CD4⫹CD25⫹ cells from ECPtreated animals to untreated allograft recipients resulted in extended cardiac allograft survival in comparison to animals that received purified CD4⫹CD25⫹ cells from control animals (Figure 5; p ⫽ 0.001, log-rank test). Therefore, the effects of ECP, as defined by the extension of graft survival, can be conferred by adoptive transfer of CD4⫹CD25⫹ T cells. DISCUSSION To approach the issue of the mechanisms of ECPmediated immunomodulation, we established an animal model of ECP treatment in which the histocompatibility differences were known, and the disposition of antigenspecific T cells could readily be analyzed within the graft and peripheral lymphoid organs. Although not a direct model of clinical ECP, the mouse system, as described in this report, provides information about the effects of intravenous infusion of UVA-irradiated cells on responses to allogeneic tissues in vivo under defined conditions. We showed that the infusion of ECP-treated cells extends cardiac allograft survival in two strains of mice, indicating that ECP can inhibit allograft rejection even under the highly unfavorable condition of no concomitant immunosuppression. In these experiments, the cells used for ECP treatment were obtained from a mouse that was syngeneic with respect to the transplant recipient. Therefore, there was no possibility of the presence of donor-antigen– bearing cells within the ECP-treated infusate prior to its introduction into the recipient, suggesting that the presence of donor antigen in the ECP infusate is not necessary for immunomodulation. It is now well established that ECP treatment induces programmed cell death of most leukocytes,5,13–15 an effect that was confirmed in our hands. Therefore, infusion of the ECP-treated cells results in a large bolus of apoptotic cells or cells destined to undergo apoptosis. There is extensive evidence that the presence of apoptotic cells within a microenvironment in vivo or in vitro is not immunologically neutral, and can profoundly affect immune function.16 –20 Studies in many different model systems have shown that phagocytosis of apoptotic cells can cause phagocytes to actively suppress inflammatory responses. Monocytes co-cul-

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Figure 3. Quantitative and qualitative analysis of T-cell infiltrate within the transplanted heart (a, b), and cells recovered from the lymph nodes (c, d) and spleen (e, f) at 1 to 3 days post-transplantation in animals injected at Day 0 with 1 ⫻ 107 CBA/CaJ splenocytes treated with 8-MOP ⫹ UVA (ECP Treated) or 8-MOP only (Control). Histograms represent the mean proportion of CFSE⫹CD8⫹ T cells ⫾ standard error of the mean calculated from 5 to 10 animals. Statistical comparisons were made using Student’s t-test. Flow cytometry histograms are gated on viable lymphoid cells as determined by forward and side scatter as well as staining with 7-AAD. The circles in (b) indicate CFSE⫹/CD8⫹ lymphocytes. Percentages shown in the two-parameter histograms (b, d, and f) represent the percentage of viable lymphoid cells.

tured with apoptotic lymphocytes19,20 or neutrophils21 secrete less tumor necrosis factor (TNF) and more transforming growth factor (TGF)-␤1 and interleukin (IL)10. Exposure of dendritic cells (DCs) to apoptotic cells causes them to become tolerogenic.22–25 Immature DCs are much more efficient than mature DCs in phagocytosing apoptotic cells.22,26 –28 Sauter et al showed that addi-

tion of apoptotic cells to immature dendritic cells prevented maturation.29 Furthermore, phagocytosis of apoptotic cells by DCs can also result in cross-tolerization of CD8⫹ T cells even with antigens that are normally presented via the exogenous pathway.30 Infusion of apoptotic cells via the peripheral blood results in their accumulation in the spleen,15,31 partic-

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Figure 4. Absolute numbers of CD3⫹ cells recovered from the donor hearts of ECP-treated and control animals at Day 3 post-transplantation. Results are expressed as the mean number of CD3⫹ cells recovered per heart. Each mean value was calculated from 3 or 4 mice, except for Day 3 in which measurements were collected from 8 to 10 mice. Error bars are the standard error of the mean. Statistical comparisons were made using Student’s t-test. N.S., no statistical difference in the mean values.

ularly in the marginal zones31 and in the liver.15 Most DCs in the peripheral lymphoid organs are functionally immature.32 Therefore, it is reasonable to speculate that infusion of apoptotic cells by ECP treatment results in their phagocytosis by macrophages and immature DCs in the marginal zones of the spleen, which, in turn, are inhibited from maturation in the presence of antigen. The data show that ECP treatment can downregulate alloreactive responses to vascularized allografts and that this downregulation occurs via a non-deletional mechanism. Although we observed a reduction in T-cell infiltration of cardiac allografts in ECP-treated animals, there was no decrease in the numbers of adoptively transferred CFSE-labeled H-2Kb–specific transgenic T cells in the peripheral lymphoid organs, indicating that the observed delay of graft rejection was effected by downregulation of allospecific T-cell function. ECP treatment decreased the rate at which these cells infiltrated cardiac allografts without affecting their frequency in the lymph nodes and spleen. The key finding in this report relative to the question of the means by which ECP can potentially modulate alloreactive responses is that this modulation has a cellular basis. Following ECP, we observed a short-term

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increase in the frequency of CD4⫹CD25⫹FoxP3⫹ T cells. More importantly, the effects of ECP, as measured by increased time to rejection, can be reproduced by adoptive transfer of purified CD4⫹CD25⫹ splenic T cells to untreated recipients. Notably, transfer of only 3 ⫻ 105 CD4⫹CD25⫹ T cells conferred extended graft survival. This quantity of cells was chosen because it corresponds to the number of CD4⫹CD25⫹ cells found in 1 ⫻ 107 splenocytes, which, when adoptively transferred from ECP-treated animals, will confer increased graft survival to some untreated graft recipients (data not shown). These data do not exclude the possibility that the transferred CD4⫹CD25⫹ cells undergo antigendriven expansion in vivo. It is also important to note that the antigenic specificity of the CD4⫹CD25⫹ cells in this system remains unknown. These findings are consistent with previous studies in a murine contact hypersensitivity model. Maeda et al showed that the capacity of ECP to reduce ear swelling in response to the hapten 2,4-dinitrofluorobenzene could be transferred with whole splenocytes, but not splenocytes in which CD4⫹ cells or CD25⫹ cells were depleted. There are differences in this model relative to the transplant system. One of the most important differences is that inhibition of contact hypersensitivity was observed only when the ECP infusate was obtained from sensitized animals, a requirement that we did not observe in our system. One possibility for this observation could be in the nature of the antigen-presentingcell (APC)/T-cell interactions of a hapten system in comparison to allografts. Allografts are characterized by a very high frequency of reactive T cells in the naive

Figure 5. A Kaplan–Meier depiction of cardiac allograft survival in C57BL/6J recipients of heterotopic abdominal vascularized cardiac allografts from BALB/cJ mice. “ECP” indicates cardiac allograft recipients received (on Day 0) 3 ⫻ 105 CD4⫹CD25⫹ cells magnetically sorted from splenocytes recovered on Day 3 post-transplant from cardiac allograft recipients that had been previously treated with ECP. “Control” designates the group that received CD4⫹CD25⫹ cells from cardiac allograft recipients that were infused with cells treated with 8-MOP only. Statistical significance was determined using the log-rank test.

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recipients because of the availability of both the direct and indirect pathways of presentation.33 Overall, these results support the hypothesis that the beneficial effects of ECP occur through induction of CD4⫹CD25⫹ T cells. Therefore, it is important to determine whether increases in the frequency and activity of these cells can be observed in human allograft recipients treated with ECP. Observations from 4 patients, 2 lung transplants and 2 heart transplants, have given a preliminary indication of increased regulatory T-cell frequency in the peripheral blood,5 suggesting that significant differences could be found in larger groups of patients. If this proves to be the case, then the efficacy of ECP could be potentially enhanced by optimization of ECP protocols for the generation of CD4⫹CD25⫹ regulatory T cells, and the frequency of the cells could possibly be further used as an indicator of treatment efficacy. The authors thank Dr Anupam Agarwal and Dr David Peritt for many helpful discussions.

REFERENCES 1. Komanduri KV, Couriel D, Champlin RE. Graft-versus-host disease after allogeneic stem cell transplantation: evolving concepts and novel therapies including photopheresis. Biol Blood Marrow Transplant 2006;12:1– 6. 2. McCuin JB, Hanlon T, Mutasim DF. Autoimmune bullous diseases: diagnosis and management. Dermatol Nurs 2006;18:20 –5. 3. Comi G. Extracorporeal phototherapy in multiple sclerosis. Neur Sci 2006;27:3– 4. 4. Kirklin JK, Brown RN, Huang ST, et al. Rejection with hemodynamic compromise: objective evidence for efficacy of photopheresis. J Heart Lung Transplant 2006;25:283– 8. 5. Lamioni A, Parisi F, Isacchi G, et al. The immunological effects of extracorporeal photopheresis unraveled: induction of tolerogenic dendritic cells in vitro and regulatory T cells in vivo. Transplantation 2005;79:846 –50. 6. Barr ML, Meiser BM, Eisen HJ, et al. Photopheresis for the prevention of rejection in cardiac transplantation. N Engl J Med 1998;339:1744 –51. 7. Yoo EK, Rook AH, Elenitsas R, Gasparro FP, Vowels BR. Apoptosis induction of ultraviolet light A and photochemotherapy in cutaneous T-cell lymphoma: relevance to mechanism of therapeutic action. J Invest Dermatol 1996;107:235– 42. 8. Johnson R, Staiano-Coico L, Austin L, et al. PUVA treatment selectively induces a cell cycle block and subsequent apoptosis in human T-lymphocytes. Photochem Photobiol 1996;63:566 –71. 9. Auphan N, Curnow J, Guimezanes A, et al. The degree of CD8 dependence of cytolytic T cell precursors is determined by the nature of the T cell receptor (TCR) and influences negative selection in TCR-transgenic mice. Eur J Immunol 1994;24:1572–7. 10. Kranz DM. Incompatible differences: view of an allogeneic pMHC–TCR complex. Nat Immunol 2000;1:277– 8. 11. Corry RJ, Winn HJ, Russell PS. Heart transplantation in congenic strains of mice. Transplant Proc 1973;5:733–5. 12. George JF, Lu A, Thomas JM, Kirklin JK, Pinderski LJ. Allogeneic bone marrow inhibits T-cell activation and clonal expansion in vitro. Transplantation 2003;76:237– 43.

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13. Heng AE, Sauvezie B, Genestier L, et al. PUVA apoptotic response in activated and resting human lymphocytes. Transfus Apheresis Sci 2003;28:43–50. 14. Legitimo A, Consolini R, Di SR, Bencivelli W, Mosca F. Psoralen and UVA light: an in vitro investigation of multiple immunological mechanisms underlying the immunosuppression induction in allograft rejection. Blood Cells Mol Dis 2002;29:24 –34. 15. Maeda A, Schwarz A, Kernebeck K, et al. Intravenous infusion of syngeneic apoptotic cells by photopheresis induces antigenspecific regulatory T cells. J Immunol 2005;174:5968 –76. 16. Mahnke K, Knop J, Enk AH. Induction of tolerogenic DCs: ’you are what you eat.’ Trends Immunol 2003;24:646 –51. 17. Savill J, Dransfield I, Gregory C, Haslett C. A blast from the past: clearance of apoptotic cells regulates immune responses. Nat Rev Immunol 2002;2:965–75. 18. Bittencourt MC, Perruche S, Contassot E, et al. Intravenous injection of apoptotic leukocytes enhances bone marrow engraftment across major histocompatibility barriers. Blood 2001;98:224 –30. 19. Voll RE, Herrmann M, Roth EA, Stach C, Kalden JR, Girkontaite I. Immunosuppressive effects of apoptotic cells. Nature 1997;390: 350 –1. 20. Fadok VA, Bratton DL, Konowal A, et al. Macrophages that have ingested apoptotic cells in vitro inhibit proinflammatory cytokine production through autocrine/paracrine mechanisms involving TGF-beta, PGE2, and PAF. J Clin Invest 1998;101:890 – 8. 21. Byrne A, Reen DJ. Lipopolysaccharide induces rapid production of IL-10 by monocytes in the presence of apoptotic neutrophils. J Immunol 2002;168:1968 –77. 22. Nouri-Shirazi M, Guinet E. Direct and indirect cross-tolerance of alloreactive T cells by dendritic cells retained in the immature stage. Transplantation 2002;74:1035– 44. 23. Skoberne M, Beignon AS, Larsson M, Bhardwaj N. Apoptotic cells at the crossroads of tolerance and immunity. Curr Top Microbiol Immunol 2005;289:259 –92. 24. Ferguson TA, Kazama H. Signals from dying cells: tolerance induction by the dendritic cell. Immunol Res 2005;32:99 –108. 25. Lutz MB, Schuler G. Immature, semi-mature and fully mature dendritic cells: which signals induce tolerance or immunity? Trends Immunol 2002;23:445–9. 26. Steinman RM, Hawiger D, Liu K, et al. Dendritic cell function in vivo during the steady state: a role in peripheral tolerance. Ann NY Acad Sci 2003;987:15–25. 27. Steinman RM. The control of immunity and tolerance by dendritic cells. Pathol Biol (Paris) 2003;51:59 – 60. 28. Mellman I, Steinman RM. Dendritic cells: specialized and regulated antigen processing machines. Cell 2001;106:255– 8. 29. Sauter B, Albert ML, Francisco L, et al. Consequences of cell death: exposure to necrotic tumor cells, but not primary tissue cells or apoptotic cells, induces the maturation of immunostimulatory dendritic cells. J Exp Med 2000;191:423–34. 30. Albert ML, Jegathesan M, Darnell RB. Dendritic cell maturation is required for the cross-tolerization of CD8⫹ T cells. Nat Immunol 2001;2:1010 –7. 31. Morelli AE, Larregina AT, Shufesky WJ, et al. Internalization of circulating apoptotic cells by splenic marginal zone dendritic cells: dependence on complement receptors and effect on cytokine production. Blood 2003;101:611–20. 32. Wilson NS, El-Sukkari D, Belz GT, et al. Most lymphoid organ dendritic cell types are phenotypically and functionally immature. Blood 2003;102:2187–94. 33. Harris PE, Cortesini R, Suciu-Foca N. Indirect allorecognition in solid organ transplantation. Rev Immunogenet 1999;1:297–308.