Impact of Psoralen/UVA-Treatment on Survival, Activation, and Immunostimulatory Capacity of Monocyte-Derived Dendritic Cells : Transplantation

Journal Logo

Original Articles: Immunobiology and Genomics

Impact of Psoralen/UVA-Treatment on Survival, Activation, and Immunostimulatory Capacity of Monocyte-Derived Dendritic Cells

Holtick, Udo1; Marshall, Scott R.1; Wang, Xiao-Nong1; Hilkens, Catharien M.U.2,3; Dickinson, Anne M.1

Author Information
doi: 10.1097/TP.0b013e31816650f6

Abstract

ArticlePlus

Click on the links below to access all the ArticlePlus for this article.

Please note that ArticlePlus files may launch a viewer application outside of your web browser.

Extracorporeal Photopheresis (ECP) has been developed in the 1980s for the treatment of cutaneous T-cell lymphoma (CTCL) (1). Its therapeutic potential has also been shown in a number of T-cell-mediated diseases and conditions not responsive to conventional immunosuppressive therapies (2–4). Clinical benefit of ECP therapy has been demonstrated in patients with acute and chronic graft-versus-host disease (GvHD) after allogeneic hematopoietic stem cell transplantation (aHSCT) (5–8).

In the setting of organ allografting, ECP has shown considerable success in treating cardiac allograft rejection (9–11), and randomized trials have demonstrated efficacy for both the prevention and treatment of cardiac rejection (12, 13). ECP efficacy has also been reported for the treatment of renal and lung rejection postallografting (14–16).

ECP is an apheresis-based therapy in which mononuclear cells are separated from whole blood, exposed to 8-methoxypsoralen (8-MOP), photoactivated with UVA light, and returned back to the patient. Although only 5% to 10% of circulating mononuclear cells are treated during one ECP procedure, long-lasting immunomodulatory effects have been demonstrated.

To date, there have been no reports that ECP therapy increases the rate of infection (17, 18). In fact, T-cell or B-cell responses to novel or recall antigens remain unaltered in patients after ECP therapy (19). In the setting of aHSCT, ECP treatment does not lead to raised relapse rates, suggesting preservation of the graft versus leukemia (GvL) effect, which is crucial for long-term survival and typically diminished by conventional immunosuppression (18).

Possible mechanisms of ECP include induction of lymphocyte apoptosis as well as modulation of antigen-presenting cells (APCs) (20). Changes in cytokine production such as Th2 skewing (21) have been observed after ECP therapy, but the data are very contentious depending on the cell source and disease condition under investigation (see recent review, Ref. 22). Maeda et al. (23) showed that the infusion of PUVA-treated apoptotic cells induces regulatory T cells in mice in a process dependent on CD11c positive cells. Furthermore, a recent report proposed that ingestion of apoptotic bodies by immature dendritic cells (DCs) led to tolerogenic DCs capable of inducing regulatory T cells (24). Although the mechanisms underlying these observations have yet to be determined, DCs are widely considered to be important in the mechanism of action of ECP (25, 26).

The current literature is controversial with regard to pro-apoptotic effects of ECP on APC. It has been suggested that monocytes are resistant to ECP-induced apoptosis (27). Edelson's group reported that monocytes are stimulated by ECP and rapidly turn into immature DCs (20). These DCs would then engulf and present tumor-associated antigens after maturation and evoke an antitumor response (Vaccination Theory). A recent study, however, observed induction of apoptosis in human monocytes in response to PUVA treatment (28). In mice, induction of apoptosis in all CD11c+ cells has been demonstrated after PUVA treatment (23), but a small study described long-term survival of immature DCs in three patients treated with ECP for chronic GvHD (29).

DCs are the most potent APC and ECP effects on DCs have to date not been studied in detail. Our work specifically investigates modulation of monocyte-derived DC (mo-DC) function and viability by ECP using a human in vitro ECP model (in vitro PUVA). Our findings show that DCs become apoptotic but that mo-DC phenotypic and functional properties are modulated by in vitro PUVA before undergoing apoptosis. These data demonstrate for the first time in vitro, that PUVA modulation of mo-DCs resulted in a shift in the pattern of stimulated T cells toward a Th2 profile.

METHODS

Cell Culture Conditions

Human cells were cultured in RPMI 1640 (Gibco; www.invitrogen.com) containing 100 IU/mL penicillin, 100 μg/mL streptomycin (Gibco) and 2 mM L-glutamin (Gibco) supplemented with 10% heat inactivated FCS (Sera Lab; www.harlanseralab.co.uk). All cultures were incubated at 37°C in a humidified 5% CO2 in air incubator.

Generation of Monocyte-Derived Dendritic Cells

Healthy donor buffy coats were obtained from the National Blood Service. Informed consent was obtained and the project was approved by the local ethics committee. Peripheral blood mononuclear cells (PBMCs) were prepared by density-gradient centrifugation on lymphoprep (Axis-Shields; www.axis-shield-density-gradient-media.com). Monocyte-derived DCs (mo-DCs) were generated as previously described (30). Magnetically isolated CD14-positive monocytes were cultured for 6 days with 50 ng/mL GM-CSF and interleukin (IL)-4 (Immunotools; www.immunotools.com); media and cytokines were refreshed on day 3. For DC maturation, immature DCs were treated for 48 hr with 10 ng/mL IL-1β and 10 ng/mL tumor necrosis factor (TNF)-α (Immunotools).

Both immature and mature mo-DC populations were tested in all subsequent experiments. Mo-DC refers therefore to both populations unless otherwise stated.

Keyhole limpet hemocyanin (KLH, Sigma, www.sigmaaldrich.com) was added to the DC culture at a concentration of 200 μg/mL 2 hr before PUVA treatment. IL-12p70 (Immunotools) was added to a separate DC culture at a concentration of 50 ng/mL 2 hr before PUVA treatment as described (31). In DC-naive T-cell co-culture experiments IL-12p70 was used at a concentration of 1 ng/mL.

Isolation of Myeloid Blood DC/In Vivo Extracorporeal Photopheresis

PBMCs from patients treated with ECP for chronic GvHD were isolated from the ECP device (UVAR XTS, Therakos, www.therakos.com) before and after UVA irradiation. Informed consent was obtained and the project was approved by the local ethics committee. Myeloid blood DCs were isolated from PBMCs using the CD1c+ (BDCA-1) Dendritic Cell Isolation kit (Miltenyi, www.miltenyi.com; purity >80%). DCs were cultured in RPMI 10%FCS containing IL-4/GM-CSF (50 ng/mL) for 24 and 48 hr and the rate of apoptosis was detected by flow cytometry using Annexin-V FITC and propidium iodide (PI) (description below).

In Vitro Psoralen/UVA Treatment

DC suspensions were irradiated with a UVA light box (Hospital Light Laboratories, UK). UVA emission was measured by a radiometer using a low-pass filter (400 nm) (International Light Inc., USA) and the irradiated power was 45 W/m2. In each well up to 5×105 mo-DCs were incubated in 50 μL of 8-MOP solution (300 ng/mL in PBS, UVADEX, Therakos) for 15 min and then exposed to UVA light (2 J/cm2). This dose is comparable to the dose used in clinical ECP procedures. After UVA-irradiation the cells were pelleted and resuspended in culture medium.

CD40-Ligand Stimulation

Untreated or PUVA-treated mo-DCs (4×104) were stimulated with 4×104 CD40-Ligand (CD40L)-transfected cells (J558L mouse cell line; kindly provided by Peter Lane, Birmingham University, UK) in 96-well flat-bottomed plates for 24 hr. Supernatants were collected and stored at −80°C for cytokine analysis.

Analysis of Cell Proliferation

CD4+ CD45RA+ naive T cells were isolated from PBMCs using the RoboSep naive T-cell isolation kit (Stem cell technology; www.stemcell.com; purity >90%). Naive T cells (105) were added to 104 mo-DCs in flat-bottomed 96-well plates in 200 μL medium. Day 5 responses were assessed after a 16-hr pulse of 0.185 MBq/mL 3H-Thymidine (TRA310, Amersham, www.amersham.com) per well and cpm determined by scintillation counting with a direct Beta-Counter (Matrix 9600, Packard Instrument, Meridian, USA).

In re-stimulation assays, mo-DCs and allogeneic naive T cells (1:10) were co-cultured for 6 days and then rested for 4 days in 0.1 ng/mL IL-2 (Immunotools) before re-stimulation with anti-CD3/CD28 coated beads (1 bead: 1 T cell, Dynabeads, Invitrogen).

Flow Cytometry

DCs were processed through a FACScan flow cytometer (Becton Dickinson; www.bd.com). DCs were identified by forward scatter (FSC) versus side scatter (SSC) plots and by the respective surface molecule expression. Data for FITC binding (FL1 channel) and PE binding (FL2 channel) were accumulated on a log fluorescence histogram plot (MFI, mean fluorescence intensity). For apoptosis detection the TACS Annexin-V FITC detection kit (R&D systems) was used according to the manufacturer's instructions. Annexin-V single positive cells and Annexin-V/PI double positive cells were considered to be early and late apoptotic respectively. A minimum of 10,000 events was acquired for each test.

For T-cell analysis, 50,000 events were acquired for each test on an LSRII cytometer (Becton Dickinson). CD4 T cells were identified using FSC/SSC, CD3, and CD4 expression, before analyzing them for CCR5/CXCR3 and CCR4/CCR10 expression.

Antibodies were obtained from Becton Dickinson unless stated otherwise. Antigen (clone): CD11c (B-ly6); HLA-DR (L243); CD86 (2331:FUN-1); CD83 (HB15e); CD80 (L307.4); CD14 (rmC5–3); CD3 (SK7); CD4 (RPA-T4); CCR4 (1G1); CCR5 (3A9); CXCR3 (1C6); CCR10 (314305; R&D systems; www.rndsystems.com); CD1a (NA1/34; DAKO; www.dako.com). Data were analyzed using FlowJo software (www.flowjo.com).

Assessment of Endocytosis

To assess DC endocytosis capacity, 5×105 mo-DCs were PUVA-treated. Untreated mo-DCs were used as control cells. After 4 hr the cells were incubated with 1 mg/mL FITC-Dextran (Sigma) for 1 hr. The cells were then fixed using 2% formalin and analyzed for intracellular FITC-Dextran by flow cytometry. Data were analyzed using FlowJo software (www.flowjo.com).

Analysis of Cell Migration

Mo-DCs or PUVA-treated mo-DCs were resuspended in RPMI 1640 medium with 0.25% FCS. Cells (5×105) were added on top of a transwell culture insert with 6.5 mm diameter and 5 μm pore size (Corning Costar; www.corning.com). The chemokines CCL-21 (200 ng/mL), CCL-19 (200 ng/mL), and CXCL-12 (100 ng/mL; all from PeproTech; www.peprotech.com) were added to the lower compartments of a 24-well plate. The number of cells that had migrated to the lower chamber within 4 hr was determined by counting in a hemocytometer.

Analysis of Cytokine Secretion

Cytokines (IL-4, IL-10, IL-13, TNF-α, IFN-γ) were detected in supernatants collected from proliferation assays using the BD cytometric bead array and analyzed using the BD-FCAP Array software (Becton Dickinson). Mo-DC cytokines (IL-12, TNF-α, IL-10) were detected by specific sandwich ELISAs obtained from BD Pharmingen.

Statistical Analysis

Statistical significance was assessed with the two-tailed Student t test using Graphpad Prism software (Graphpad; www.graphpad.com) with P less than 0.05 considered significant.

RESULTS

PUVA Induces Partial Maturation and Apoptosis of mo-DCs

We first analyzed the rate of apoptosis in immature and mature mo-DCs 24 and 48 hr after in vitro PUVA treatment by flow cytometry. Annexin-V and PI were used as markers for early and late apoptosis. In parallel, we analyzed the expression of mo-DC cell surface markers involved in APC function (HLA-DR, CD80, CD83, and CD86).

In vitro PUVA induced apoptosis in both immature and mature mo-DCs. The percentage of viable nonapoptotic mo-DCs (i.e., double negative for Annexin-V and PI) was reduced from 81.5% to 29.4% for immature mo-DCs and from 87.8% to 40% for mature mo-DCs after 24 hr of PUVA treatment (Fig. 1A,C). After 48 hr of PUVA treatment, the percentage of viable mo-DCs declined to 13.5% (immature) and 23.2% (mature) (Fig. 1B,D). UVA irradiation or 8-MOP alone did not induce apoptosis (data not shown) as has been reported before (32).

F1-17
FIGURE 1.:
Rate of apoptosis at 24 hr (A,C) and 48 hr (B,D) after PUVA treatment. Data shown are percentages of one representative experiment of four. (E) PBMCs of patients treated with ECP showing increased apoptosis at comparable levels after 24 and 48 hr. Data shown are representative of 3 experiments. Background levels of apoptosis were attributed to the processing of cells by the ECP device and DC isolation.

In vivo ECP also induced apoptosis in CD1c+ myeloid blood DCs at comparable levels after 24 and 48 hr (Fig. 1E). Analysis of apoptosis was extended to Langerhans cells and dermal DCs and similar rates of apoptosis were observed after in vitro PUVA treatment (Supplemental Fig. 1; available for viewing online only).

The induction of apoptosis was preceded by a transient and partial maturation of immature mo-DCs. In vitro PUVA treatment significantly up-regulated HLA-DR in immature mo-DCs at 4 and 8 hr and CD86 at 4 hr (P<0.05; Table 1 and Supplemental Fig. 2A, available for viewing online only). CD83 was up-regulated 1.8–fold, but this was not significant (P=0.07). CD80 remained unchanged 4 to 8 hr after PUVA treatment (Table 1 and Supplemental Fig. 2, available for viewing online only). Although PUVA enhanced expression of HLA-DR, CD83, and CD86 on immature mo-DCs, the levels of these surface molecules were below levels observed on mature mo-DCs (Supplemental Fig. 2, available for viewing online only), indicating that PUVA treatment led to partial maturation of mo-DCs. Up-regulation of surface molecules however was only transient and could not be observed 24 hr after PUVA treatment (Table 1).

T1-17
TABLE 1:
Fold induction of CD80/83/86 and HLA-DR expression on immature and mature mo-DC 4 hr, 8 hr, and 24 hr after in vitro PUVA treatment as analyzed by flow cytometry

In mature mo-DCs, PUVA treatment also led to up-regulation of HLA-DR at 4 and 8 hr, but again this effect was transient and could not be observed after 24 hr. In contrast to immature mo-DCs, expression of CD80, CD86, and CD83 was not transiently up-regulated in mature mo-DCs on PUVA treatment. Instead, levels of these surface markers decreased continuously after treatment and after 24 hr were between 61% and 82% of control (no PUVA) levels (Table 1 and Supplemental Fig. 2B, available for viewing online only). Treatment with UVA irradiation or 8-MOP alone did not affect surface molecule expression (data not shown).

These data show that in vitro PUVA induces apoptosis in DCs, but in immature mo-DCs, this is preceded by transient and partial maturation as defined by up-regulation of HLA-DR, CD83, and CD86.

PUVA-Induced mo-DC Apoptosis Cannot Be Rescued by Antigen or IL-12

We hypothesized that stimulation of DCs by either antigen or IL-12 could induce survival factors protecting from apoptosis. IL-12 has been shown to protect cells from UVB irradiation-induced DNA damage (31). Prestimulation with antigen (keyhole limpet hemocyanin, KLH) slightly enhanced viability of PUVA-treated immature but not mature mo-DCs (Fig. 1). However, antigen stimulation did not restore the reduced cell viability of PUVA-treated mo-DCs to the same levels as nontreated mo-DCs. Prestimulation with IL-12 also enhanced the viability of immature mo-DCs 24 hr, but not 48 hr after PUVA treatment (Fig. 1A,B). In contrast, the viability of mature mo-DCs was not affected by IL-12 prestimulation at either time point (Fig. 1C,D).

PUVA Decreases Endocytosis and Increases Migratory Capacity

The observation that PUVA treatment led to transient and partial mo-DC maturation raised the question of whether typical DC functions associated with maturation were modulated by PUVA. First of all, we examined the effect of in vitro PUVA on the endocytosis capacity of mo-DCs, which is known to decline during maturation. Four hours after PUVA treatment, immature and mature mo-DCs were cultured with FITC-coupled dextran for 1 hr. Cells were assessed by flow cytometry for FITC-Dextran uptake. As anticipated, the majority (94%) of immature mo-DCs had taken up dextran when compared with 28% of mature mo-DCs (Fig. 2A). On PUVA treatment, the endocytosis capacity of mo-DCs was decreased and only 54% of immature and 23% of mature mo-DCs were FITC-dextran positive (Fig. 2A).

F2-17
FIGURE 2.:
(A) Endocytosis capacity of immature and mature mo-DCs before and after in vitro PUVA treatment. FITC staining showing % positive cells, representative experiment out of three. (B) Migratory capacity of immature and mature mo-DCs before and 4 hr after in vitro PUVA treatment. Data shown are mean±SD of four experiments. *P<0.05, treated vs. the control group.

Second, the migratory capacity of mo-DCs after in vitro PUVA was analyzed using a transwell system. Maturation of DCs is known to enhance the migration of DCs in response to specific chemokines. As expected, the migratory capacity of untreated mature mo-DCs toward CCL-19, CCL-21, and CXCL-12 was 20-fold higher than migration of untreated immature mo-DCs. However, migration of immature mo-DCs toward CCL-21 and CCL-19 was significantly enhanced after in vitro PUVA treatment (P<0.05; Fig. 2B), whereas migration of mature mo-DCs was decreased after PUVA treatment (Fig. 2B). The results from experiments on surface molecule expression, endocytosis, and migratory capacity suggested that the effect of in vitro PUVA led to partial maturation of immature mo-DCs, whereas it had only marginal effects on the functional mature mo-DCs.

PUVA Enhances Immunostimulatory Capacity of mo-DCs

To determine whether the observed partial maturation of mo-DCs by in vitro PUVA enhanced their immunostimulatory capacity, treated or untreated immature and mature mo-DCs were used as APC for allogeneic naive T cells. In vitro PUVA effects were compared with γ-irradiation and UVA. When immature mo-DCs were used as APC immediately after PUVA treatment, proliferation of naive T cells was significantly enhanced to levels observed when mature mo-DCs were used as APC (Fig. 3A). This effect was specific for PUVA and could not be observed when mo-DCs were treated with γ-irradiation or UVA. None of the treatments affected the T-cell stimulatory capacity of mature mo-DCs (Fig. 3A). In contrast, when mo-DCs were used as APC 24 hr after in vitro PUVA treatment, the immunostimulatory capacity of both immature and mature mo-DCs was significantly diminished, whereas it was found unchanged for γ-irradiation or UVA treatment (Fig. 3B). These results reflect the short-lived changes in mo-DC phenotype and the induction of apoptosis as described before.

F3-17
FIGURE 3.:
Proliferation of allogeneic naive T cells after stimulation with immature or mature mo-DCs treated with in vitro PUVA, 20 Gy, or UVA. (A) Immediately after and (B) 24 hr after the respective treatment. Data shown are mean ± SD of three separate experiments in triplicates. *P<0.05, treated vs. the control group.

PUVA Treatment Abrogates CD40L-Induced mo-DC Activation and IL-12 Production

CD40-Ligand (CD40L) stimulates the expression of DC surface molecules and cytokine production by signaling through CD40 on DCs, thereby mimicking DC-T-cell interaction. It was tested if in vitro PUVA treatment interfered with CD40-mediated DC activation. Stimulation with CD40L strongly up-regulated the expression of CD80, CD83, CD86, and HLA-DR on immature (Fig. 4A) and mature mo-DCs (Fig. 4 B). In vitro PUVA treatment abrogated these CD40L effects (Fig. 4A,B). The effects of PUVA treatment on CD40L-induced cytokine production (IL-12p70, TNF-a, IL-10) was also assessed. CD40L induced IL-12, IL-10, and TNF-α secretion in immature and mature mo-DCs (Fig. 4C–H). In vitro PUVA inhibited the effect of CD40L-induced cytokine secretion (P<0.05, Fig. 4C–H), which was especially evident for IL-12 and IL-10.

F4-17
FIGURE 4.:
In vitro PUVA affects mo-DC phenotype and cytokine secretion on CD40L-activation. Expression of CD80/83/86 and HLA-DR on (A) immature or (B) mature mo-DCs as analyzed by flow cytometry. Cells were gated for CD11c to identify DC. Data shown are representative of four experiments. IL-12 secretion by (C) immature or (D) mature mo-DCs; TNF-α secretion by (E) immature or (F) mature mo-DCs; IL-10 secretion by (G) immature or (H) mature mo-DCs; data shown are mean±SD of four experiments. *P<0.05, treated vs. the control group.

PUVA-Treated mo-DCs Polarize Naive T Cells Toward a Th2 Phenotype

We next addressed the question of whether naive T cells primed by PUVA-treated mo-DCs would develop a distinct cytokine profile. Untreated or PUVA-treated mo-DCs were used as APC for allogeneic naive T cells. After 10 days of culture, the T cells were re-stimulated, and production of IL-4, IL-10, IL-13, TNF-α, and interferon (IFN)-γ secretion was measured. Naive T cells that had been primed with PUVA-treated immature mo-DCs secreted significantly more IL-4, IL-10, IL-13, and less IFN-γ and TNF-α (P<0.05, Fig. 5A) than T cells primed with untreated immature mo-DCs. T cells primed by PUVA-treated mature mo-DCs showed a similar pattern, but TNF-α levels were slightly higher than TNF-α levels of T cells primed with untreated mature mo-DCs (Fig. 5B). The proliferative response was not significantly altered in any of the groups (data not shown).

F5-17
FIGURE 5.:
PUVA-treated or untreated immature (A,C,D) or mature (B,E,F) mo-DCs as primary stimulators for allogeneic naive T cells. (A,B) Data shown are mean±SD of four separate experiments. *P<0.05 treated vs. control group. (C–F) Data shown are representative of four experiments.

It is known that IL-12 is a crucial cytokine for the development of a Th1 response (33). As secretion of IL-12 by DCs is strongly suppressed after PUVA treatment (Fig. 4C,D), we hypothesized that lack of IL-12 might contribute to the observed Th2 shift. To test this hypothesis, IL-12 was added to the primary DC-naive T-cell co-culture and the cytokine profile was again assessed after re-stimulation of T cells. Addition of IL-12 restored IFN-γ levels, elevated TNF-α levels, and partially reversed the higher levels of IL-4, IL-10, and IL-13 (Fig. 5A,B).

This Th2 shift was confirmed by analysis of Th1/Th2 associated chemokine receptor expression after re-stimulation using flow cytometry (34–36). Priming with PUVA-treated mo-DCs decreased the percentage of CCR5/CXCR3 expressing T cells (Th1 associated) and increased the percentage of CCR4/CCR10 positive T cells (Th2 associated). This finding was partially reversed when IL-12 was added to the primary DC-naive T-cell co-culture (Fig. 5C–F). Together these data suggest that PUVA treatment renders both mature and immature mo-DCs to shift T-cell responses in favor of Th2. Down-regulation of IL-12 secretion could be a mechanism to explain this observation.

DISCUSSION

Our study addresses the controversy in the literature regarding ECP effects on APCs. Using in vitro PUVA treatment of mo-DCs as a model for ECP we provide evidence that mo-DCs are directly modulated by in vitro PUVA and that this modulation impacts on the resulting T-cell response. In contrast to previous findings, we also demonstrate that both in vitro PUVA and in vivo ECP induced apoptosis in DCs.

Spisek et al. (29) reported that DCs isolated from three chronic GvHD patients during the ECP process survived for a number of days and produced IL-10. Our study did not reproduce these results. In contrast, we demonstrate a profound effect of in vitro PUVA and in vivo ECP on the viability of DCs. Spisek et al. may have underestimated apoptosis as only Annexin-V/PI double positive cells were classified as apoptotic, whereas we included Annexin-V single positive cells as early apoptotic. More importantly, our in vitro system was titrated to resemble the closed-circuit UVAR XTS ECP device (Therakos), which was also used for the in vivo experiment, whereas the study by Spisek et al. used the Cobe spectra apheresis system (Gambro BCT) and a separate UVA source. To our knowledge, a comparison of these two systems with regard to apoptosis induction has not been done.

The absence of generalized immunosuppression after ECP treatment suggests rather specific effects against pathogenic T cells. T-cell receptors (TCRs) of clonal T cells can become targets of TCR-specific anti-idiotype immune responses (37, 38). In CTCL, circulating clonally amplified malignant T cells are reduced by ECP treatment (39). Oligoclonal T-cell populations have also been described in autoimmune disease and chronic GvHD (40). Data from animal autoimmune and transplantation models imply that the process of photopheresis induces specific effects against clonal amplified T-cell populations (41, 42). It has been reported that ECP induces the generation of activated DCs from monocytes and instigation of antitumor responses in CTCL by these DCs loaded with ECP-induced apoptotic lymphoma cells (20, 43). Our own unpublished results and a recent publication (44) do not support ECP/PUVA-induced DCs generation from monocytes. However, we observed activation of immature mo-DCs after PUVA treatment. These activated DCs could work similar to DCs described by the Edelson group and take part in the stimulation of beneficial immune responses, but the role of anti-idiotype responses in ECP treatment of GvHD needs to be further investigated.

The rapid kinetics of DC activation, trafficking and priming of immune responses have been demonstrated before (45). Watanabe et al. (46) reported DC migration toward secondary lymphoid tissues as soon as 3 hr after activation through CD40. These DCs were capable of priming T cells to provide antitumor effects after subcutaneous injection. A different study showed superiority of early activated DCs over 24 hr activated DCs in their ability to induce specific T-cell responses (47). Work in a murine GvHD model showed that preterminal DCs derived from lethally irradiated mice were immediately activated after irradiation and aggregated in T-cell areas where they activated alloreactive T cells (48). Similarly, PUVA-modulated immature mo-DCs acquired enhanced immunostimulatory capacity and could efficiently prime T cells shortly after PUVA treatment. However, 24 hr after PUVA treatment, the immunostimulatory capacity of PUVA-treated immature and mature mo-DCs was reduced in keeping with the decreased expression of co-stimulatory molecules and decreased production of inflammatory cytokines.

Although ECP can be successfully used for the treatment of GvHD or transplant rejection, the response rates vary significantly (17, 18, 49). Mediators provided by the inflammatory condition might alter cellular responses to ECP. IL-12 is an interesting cytokine in the context of ECP, as it is involved in DNA protection from UV-irradiation damage (31). Conversely, IL-12 gene expression is suppressed in macrophages after phagocytosis of apoptotic cells (50). IL-12 has been shown to protect cells from UVB irradiation-induced DNA damage by up-regulation of DNA repair enzymes (51), and DC activation has been reported to induce survival factors (52). However, we observed only a small effect of IL-12 or KLH DC prestimulation on survival of PUVA-treated immature mo-DCs. Apoptosis of PUVA-treated mo-DCs could not be prevented, indicating that these two stimuli are not sufficient to alter DC responses to ECP in vitro.

In our study, in vitro PUVA treatment abrogated activation of mo-DCs through CD40 ligation. Blocking CD40 signaling has been used as one strategy to induce tolerance in both organ and hematopoietic cell transplant models (53–55). Hence, blocking DC susceptibility to CD40 activation by PUVA treatment could contribute to tolerance induction by ECP.

The characteristics of PUVA-treated mo-DCs after CD40 ligation are reminiscent of DC exhaustion, which usually occurs late after maturation (45, 56, 57). Similar to our findings, it has been described that in addition to low IL-12 secretion these exhausted DCs become unresponsive to CD40L stimulation (45). Moreover, with the loss of IL-12 secretion a switch from a Th1 to a Th2 inducing mode has been reported (33, 45, 58). We demonstrated decreased secretion of the Th1 cytokine IFN-γ, but increased secretion of the Th2 cytokines IL-4, IL-10, and IL-13 by naive T cells primed with PUVA treated immature or mature mo-DCs. In solid organ graft rejection, beneficial effects of IL-4, IL-10, and IL-13 have been reported (59–62), whereas reduced IFN-γ levels were correlated with prolonged graft survival in a liver transplant model (63). In the setting of aHSCT, Gorgun et al. (21) could demonstrate a shift of the cytokine balance toward Th2 in nine of ten patients treated with ECP for chronic GvHD. Studies on PUVA-treated PBMCs also indicated increased IL-10 and IL-4 levels (64, 65). The mechanism of this shift has not been elucidated yet, but our data suggest that PUVA-modulated DCs could be involved in polarizing these Th2 responses. Interestingly, the observed Th2 shift was partially reversed by the addition of IL-12. Lack of IL-12 secretion by DCs is one of the factors contributing to induction of Th2 cells. Another factor favoring a Th2 response is IL-10 (58, 66). PUVA treatment abrogated IL-12 secretion by mo-DCa, but IL-10 levels were also markedly reduced. These results suggest that lack of IL-12 facilitates the Th2 shift.

In conclusion, our study is the first to analyze in detail the direct effects of PUVA on mo-DCs. We describe the induction of apoptosis and DC modulation as two separate mechanisms of action of ECP and provide a possible mechanism for the Th2 shift reported in ECP-treated chronic GvHD patients. Future work will have to test in vivo the implications for clinical efficacy of ECP.

REFERENCES

1. Edelson R, Berger C, Gasparro F, et al. Treatment of cutaneous T-cell lymphoma by extracorporeal photochemotherapy. Preliminary results. N Engl J Med 1987; 316: 297.
2. Rook AH, Freundlich B, Jegasothy BV, et al. Treatment of systemic sclerosis with extracorporeal photochemotherapy. Results of a multicenter trial[see comment]. Arch Dermatol 1992; 128: 337.
3. Knobler RM, Graninger W, Lindmaier A, Trautinger F, Smolen JS. Extracorporeal photochemotherapy for the treatment of systemic lupus erythematosus. A pilot study. Arthritis Rheum 1992; 35: 319.
4. Reinisch W, Nahavandi H, Santella R, et al. Extracorporeal photochemotherapy in patients with steroid-dependent Crohn's disease: A prospective pilot study. Aliment Pharmacol Ther 2001; 15: 1313.
5. Greinix HT, Volc-Platzer B, Rabitsch W, et al. Successful use of extracorporeal photochemotherapy in the treatment of severe acute and chronic graft-versus-host disease. Blood 1998; 92: 3098.
6. Smith EP, Sniecinski I, Dagis AC, et al. Extracorporeal photochemotherapy for treatment of drug-resistant graft-vs.-host disease. Biol Blood Marrow Transplant 1998; 4: 27.
7. Messina C, Locatelli F, Lanino E, et al. Extracorporeal photochemotherapy for paediatric patients with graft-versus-host disease after haematopoietic stem cell transplantation. Br J Haematol 2003; 122: 118.
8. Foss FM, DiVenuti GM, Chin K, et al. Prospective study of extracorporeal photopheresis in steroid-refractory or steroid-resistant extensive chronic graft-versus-host disease: Analysis of response and survival incorporating prognostic factors. Bone Marrow Transplant 2005; 35: 1187.
9. Costanzo-Nordin MR, Hubbell EA, O'Sullivan EJ, et al. Successful treatment of heart transplant rejection with photopheresis. Transplantation 1992; 53: 808.
10. Dall'Amico R, Montini G, Murer L, et al. Benefits of photopheresis in the treatment of heart transplant patients with multiple/refractory rejection. Transplant Proc 1997; 29: 609.
11. Meiser BM, Kur F, Reichenspurner H, et al. Reduction of the incidence of rejection by adjunct immunosuppression with photochemotherapy after heart transplantation. Transplantation 1994; 57: 563.
12. Barr ML, Meiser BM, Eisen HJ, et al. Photopheresis for the prevention of rejection in cardiac transplantation. Photopheresis Transplantation Study Group. N Engl J Med 1998; 339: 1744.
13. Barr ML, Baker CJ, Schenkel FA, et al. Prophylactic photopheresis and chronic rejection: Effects on graft intimal hyperplasia in cardiac transplantation. Clin Transplant 2000; 14: 162.
14. Sunder-Plassman G, Druml W, Steininger R, Honigsmann H, Knobler R. Renal allograft rejection controlled by photopheresis. Lancet 1995; 346: 506.
15. Salerno CT, Park SJ, Kreykes NS, et al. Adjuvant treatment of refractory lung transplant rejection with extracorporeal photopheresis. J Thorac Cardiovasc Surg 1999; 117: 1063.
16. Slovis BS, Loyd JE, King LE, Jr. Photopheresis for chronic rejection of lung allografts. N Engl J Med 1995; 332: 962.
17. Marques MB, Tuncer HH. Photopheresis in solid organ transplant rejection. J Clin Apher 2006; 21: 72.
18. Marshall SR. Technology insight: ECP for the treatment of GvHD—can we offer selective immune control without generalized immunosuppression? Nat Clin Pract Oncol 2006; 3: 302.
19. Suchin KR, Cassin M, Washko R, et al. Extracorporeal photochemotherapy does not suppress T- or B-cell responses to novel or recall antigens. J Am Acad Dermatol 1999; 41: 980.
20. Berger CL, Hanlon D, Kanada D, Girardi M, Edelson RL. Transimmunization, a novel approach for tumor immunotherapy. Transfus Apher Sci 2002; 26: 205.
21. Gorgun G, Miller KB, Foss FM. Immunologic mechanisms of extracorporeal photochemotherapy in chronic graft-versus-host disease. Blood 2002; 100: 941.
22. Bladon J, Taylor PC. Extracorporeal photopheresis: A focus on apoptosis and cytokines. J Dermatol Sci 2006; 43: 85.
23. Maeda A, Schwarz A, Kernebeck K, et al. Intravenous infusion of syngeneic apoptotic cells by photopheresis induces antigen-specific regulatory T cells. J Immunol 2005; 174: 5968.
24. 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.
25. Greinix HT, Socie G, Bacigalupo A, et al. Assessing the potential role of photopheresis in hematopoietic stem cell transplant. Bone Marrow Transplant 2006; 38: 265.
26. Peritt D. Potential mechanisms of photopheresis in hematopoietic stem cell transplantation. Biol Blood Marrow Transplant 2006; 12 (1 Suppl 2): 7.
27. Tambur AR, Ortegel JW, Morales A, Klingemann H, Gebel HM, Tharp MD. Extracorporeal photopheresis induces lymphocyte but not monocyte apoptosis. Transplant Proc 2000; 32: 747.
28. Legitimo A, Consolini R, Bencivelli W, Crimaldi G, Migliaccio P, Mosca F. Assessment of 8-methoxypsoralen and ultraviolet a light effects on human stroma generation and function. Acta Haematol 2006; 116: 192.
29. Spisek R, Gasova Z, Bartunkova J. Maturation state of dendritic cells during the extracorporeal photopheresis and its relevance for the treatment of chronic graft-versus-host disease. Transfusion 2006; 46: 55.
30. Sallusto F, Lanzavecchia A. Efficient presentation of soluble antigen by cultured human dendritic cells is maintained by granulocyte/macrophage colony-stimulating factor plus interleukin 4 and downregulated by tumor necrosis factor alpha. J Exp Med 1994; 179: 1109.
31. Schwarz A, Stander S, Berneburg M, et al. Interleukin-12 suppresses ultraviolet radiation-induced apoptosis by inducing DNA repair. Nat Cell Biol 2002; 4: 26.
32. Heng AE, Sauvezie B, Genestier L, Demeocq F, Dosgilbert A, Deteix P. PUVA apoptotic response in activated and resting human lymphocytes. Transfus Apher Sci 2003; 28: 43.
33. Hsieh CS, Macatonia SE, Tripp CS, Wolf SF, O'Garra A, Murphy KM. Development of TH1 CD4+ T cells through IL-12 produced by Listeria-induced macrophages. Science 1993; 260: 547.
34. Reiss Y, Proudfoot AE, Power CA, Campbell JJ, Butcher EC. CC chemokine receptor (CCR)4 and the CCR10 ligand cutaneous T cell-attracting chemokine (CTACK) in lymphocyte trafficking to inflamed skin. J Exp Med 2001; 194: 1541.
35. Bonecchi R, Bianchi G, Bordignon PP, et al. Differential expression of chemokine receptors and chemotactic responsiveness of type 1 T helper cells (Th1s) and Th2s. J Exp Med 1998; 187: 129.
36. Langenkamp A, Nagata K, Murphy K, Wu L, Lanzavecchia A, Sallusto F. Kinetics and expression patterns of chemokine receptors in human CD4+ T lymphocytes primed by myeloid or plasmacytoid dendritic cells. Eur J Immunol 2003; 33: 474.
37. Offner H, Hashim GA, Vandenbark AA. T cell receptor peptide therapy triggers autoregulation of experimental encephalomyelitis. Science 1991; 251: 430.
38. Zhang J, Medaer R, Stinissen P, Hafler D, Raus J. MHC-restricted depletion of human myelin basic protein-reactive T cells by T cell vaccination. Science 1993; 261: 1451.
39. Rook AH, Prystowsky MB, Cassin M, Boufal M, Lessin SR. Combined therapy for Sezary syndrome with extracorporeal photochemotherapy and low-dose interferon alfa therapy. Clinical, molecular, and immunologic observations. Arch Dermatol 1991; 127: 1535.
40. French LE, Alcindor T, Shapiro M, et al. Identification of amplified clonal T cell populations in the blood of patients with chronic graft-versus-host disease: Positive correlation with response to photopheresis. Bone Marrow Transplant 2002; 30: 509.
41. Berger CL, Perez M, Laroche L, Edelson R. Inhibition of autoimmune disease in a murine model of systemic lupus erythematosus induced by exposure to syngeneic photoinactivated lymphocytes. J Invest Dermatol 1990; 94: 52.
42. Girardi M, Herreid P, Tigelaar RE. Specific suppression of lupus-like graft-versus-host disease using extracorporeal photochemical attenuation of effector lymphocytes. J Invest Dermatol 1995; 104: 177.
43. Berger CL, Xu AL, Hanlon D, et al. Induction of human tumor-loaded dendritic cells. Int J Cancer 2001; 91: 438.
44. Legitimo A, Consolini R, Failli A, et al. In vitro treatment of monocytes with 8-methoxypsolaren and ultraviolet A light induces dendritic cells with a tolerogenic phenotype. Clin Exp Immunol 2007.
45. Langenkamp A, Messi M, Lanzavecchia A, Sallusto F. Kinetics of dendritic cell activation: Impact on priming of TH1, TH2 and nonpolarized T cells. Nat Immunol 2000; 1: 311.
46. Watanabe S, Kagamu H, Yoshizawa H, et al. The duration of signaling through CD40 directs biological ability of dendritic cells to induce antitumor immunity. J Immunol 2003; 171: 5828.
47. Camporeale A, Boni A, Iezzi G, et al. Critical impact of the kinetics of dendritic cells activation on the in vivo induction of tumor-specific T lymphocytes. Cancer Res 2003; 63: 3688.
48. Zhang Y, Louboutin JP, Zhu J, Rivera AJ, Emerson SG. Preterminal host dendritic cells in irradiated mice prime CD8+ T cell-mediated acute graft-versus-host disease. J Clin Invest 2002; 109: 1335.
49. McKenna KE, Whittaker S, Rhodes LE, et al. Evidence-based practice of photopheresis 1987–2001: A report of a workshop of the British Photodermatology Group and the U.K. Skin Lymphoma Group. Br J Dermatol 2006; 154: 7.
50. Kim S, Elkon KB, Ma X. Transcriptional suppression of interleukin-12 gene expression following phagocytosis of apoptotic cells. Immunity 2004; 21: 643.
51. Schwarz A, Maeda A, Kernebeck K, van Steeg H, Beissert S, Schwarz T. Prevention of UV radiation-induced immunosuppression by IL-12 is dependent on DNA repair. J Exp Med 2005; 201: 173.
52. Rescigno M, Martino M, Sutherland CL, Gold MR, Ricciardi-Castagnoli P. Dendritic cell survival and maturation are regulated by different signaling pathways. J Exp Med 1998; 188: 2175.
53. Larsen CP, Elwood ET, Alexander DZ, et al. Long-term acceptance of skin and cardiac allografts after blocking CD40 and CD28 pathways. Nature 1996; 381: 434.
54. Seung E, Iwakoshi N, Woda BA, et al. Allogeneic hematopoietic chimerism in mice treated with sublethal myeloablation and anti-CD154 antibody: Absence of graft-versus-host disease, induction of skin allograft tolerance, and prevention of recurrent autoimmunity in islet-allografted NOD/Lt mice. Blood 2000; 95: 2175.
55. Masunaga T, Yamashita K, Sakihama H, et al. Dimeric but not monomeric soluble CD40 prolongs allograft survival and generates regulatory T cells that inhibit CTL function. Transplantation 2005; 80: 1614.
56. Kalinski P, Schuitemaker JH, Hilkens CM, Wierenga EA, Kapsenberg ML. Final maturation of dendritic cells is associated with impaired responsiveness to IFN-gamma and to bacterial IL-12 inducers: Decreased ability of mature dendritic cells to produce IL-12 during the interaction with Th cells. J Immunol 1999; 162: 3231.
57. Lanzavecchia A, Sallusto F. Regulation of T cell immunity by dendritic cells. Cell 2001; 106: 263.
58. Kalinski P, Hilkens CM, Snijders A, Snijdewint FG, Kapsenberg ML. IL-12-deficient dendritic cells, generated in the presence of prostaglandin E2, promote type 2 cytokine production in maturing human naive T helper cells. J Immunol 1997; 159: 28.
59. Fischbein MP, Yun J, Laks H, et al. Regulated interleukin-10 expression prevents chronic rejection of transplanted hearts. J Thorac Cardiovasc Surg 2003; 126: 216.
60. Zhou X, Schmidtke P, Zepp F, Meyer CU. Boosting interleukin-10 production: Therapeutic effects and mechanisms. Curr Drug Targets Immune Endocr Metabol Disord 2005; 5: 465.
61. Stammberger U, Bilici M, Gugger M, et al. Prolonged amelioration of acute lung allograft rejection by overexpression of human interleukin-10 under control of a long acting ubiquitin C promoter in rats. J Heart Lung Transplant 2006; 25: 1474.
62. Davidson C, Verma ND, Robinson CM, et al. IL-13 prolongs allograft survival: Association with inhibition of macrophage cytokine activation. Transpl Immunol 2007; 17: 178.
63. Obara H, Nagasaki K, Hsieh CL, et al. IFN-gamma, produced by NK cells that infiltrate liver allografts early after transplantation, links the innate and adaptive immune responses. Am J Transplant 2005; 5: 2094.
64. Klosner G, Trautinger F, Knobler R, Neuner P. Treatment of peripheral blood mononuclear cells with 8-methoxypsoralen plus ultraviolet A radiation induces a shift in cytokine expression from a Th1 to a Th2 response. J Invest Dermatol 2001; 116: 459.
65. Craciun LI, Stordeur P, Schandene L, et al. Increased production of interleukin-10 and interleukin-1 receptor antagonist after extracorporeal photochemotherapy in chronic graft-versus-host disease. Transplantation 2002; 74: 995.
66. Liu J, Anderson BE, Robert ME, et al. Selective T-cell subset ablation demonstrates a role for T1 and T2 cells in ongoing acute graft-versus-host disease: A model system for the reversal of disease. Blood 2001; 98: 3367.
Keywords:

Extracorporeal photopheresis; PUVA; Dendritic cells; Graft rejection; Graft-versus-host disease

Supplemental Digital Content

© 2008 Lippincott Williams & Wilkins, Inc.