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The Journal of Immunology, 2004, 173: 4828-4837.
Copyright © 2004 by The American Association of Immunologists

A Novel Pathway of Alloantigen Presentation by Dendritic Cells1

Osquel Barroso Herrera, Dela Golshayan, Rebecca Tibbott, Francisco Salcido Ochoa, Martha J. James, Federica M. Marelli-Berg and Robert I. Lechler2

Department of Immunology, Imperial College, Hammersmith Hospital, London, United Kingdom


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
In the context of transplantation, dendritic cells (DCs) can sensitize alloreactive T cells via two pathways. The direct pathway is initiated by donor DCs presenting intact donor MHC molecules. The indirect pathway results from recipient DCs processing and presenting donor MHC as peptide. This simple dichotomy suggests that T cells with direct and indirect allospecificity cannot cross-regulate each other because distinct APCs are involved. In this study we describe a third, semidirect pathway of MHC alloantigen presentation by DCs that challenges this conclusion. Mouse DCs, when cocultured with allogeneic DCs or endothelial cells, acquired substantial levels of class I and class II MHC:peptide complexes in a temperature- and energy-dependent manner. Most importantly, DCs acquired allogeneic MHC in vivo upon migration to regional lymph nodes. The acquired MHC molecules were detected by Ab staining and induced proliferation of Ag-specific T cells in vitro. These data suggest that recipient DCs, due to acquisition of donor MHC molecules, may link T cells with direct and indirect allospecificity.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Alloreactive T cells recognize alloantigens via two distinct, but not mutually exclusive, pathways: direct and indirect. In the direct pathway of allorecognition, responder T cells recognize intact foreign MHC:peptide complexes on the surface of donor cells (1, 2, 3, 4, 5). This pathway is characterized by the high precursor frequency of responder T cells (~100-fold greater than for a response to conventional protein Ags) and is critical in the initiation of the alloresponse and acute graft rejection (3, 6, 7, 8). By the indirect pathway, in contrast, recipient T cells recognize peptides derived from foreign MHC molecules after processing and presentation by self-MHC molecules on recipient APCs (8, 9, 10). The indirect pathway, in other words, is the normal mechanism of self MHC-restricted T cell stimulation. During recent years, increasing experimental evidence has supported the role for this pathway in both acute and chronic graft rejection (11, 12).

Several lines of evidence suggest that T cells with indirect allospecificity can regulate, positively or negatively, direct pathway T cells. Indeed, Lee et al. (13) have shown, using MHC class II-deficient skin grafts, that CD4+ T cells with indirect anti-donor specificity can amplify direct pathway CD8+ T cell responses. Similarly, tolerant indirect pathway T cells can suppress the response of direct pathway T cells in some models (14, 15). It has been shown that the induction of tolerance to minor mismatched skin grafts in mice using nondepleting anti-CD4 and anti-CD8 mAbs involves the reprocessing of minors on host APCs and the induction of regulatory CD4+ T cells with indirect pathway specificity (14). In addition, elimination of CD4+ T cells from tolerant mice resulted in the rejection of long-standing grafts, suggesting that direct pathway CD8+ T cells had been under continuous regulation by tolerant, indirect pathway CD4+ T cells (15). These observations point to a four-cell, unlinked, model for interactions between direct and indirect pathway T cells during the course of graft rejection: helper, or suppressor, CD4+ T cells with indirect specificity are activated by recipient dendritic cells (DCs)3 that reside in secondary lymphoid organs, whereas direct pathway effector CD8+ T cells must recognize determinants expressed on the cells of the donor graft.

We propose in this study that the recently described phenomenon of MHC transfer between cells may provide a mechanism to resolve this conundrum. Several studies have shown that DCs are capable of acquiring intact MHC molecules from other cells (other DCs, macrophages, activated T cells, B cells, and tumor cells) in vitro (16, 17, 18). Harshyne et al. (18) have reported that individual DCs, especially immature DCs, physically extract plasma membrane and, to a lesser extent, intracellular proteins from other DCs in a cell contact-dependent fashion. In addition, DCs are capable of shedding soluble MHC molecules and membrane vesicles, called exosomes, that express MHC class I and class II molecules and costimulatory molecules that can be captured by other DCs (16, 19). As a consequence, there could be a third, semidirect pathway of allorecognition by which direct pathway T cells would recognize allogeneic MHC molecules after being transferred, intact, from the surface of donor cells to the surface of recipient DCs. In this study the acquisition and presentation of MHC:peptide complexes by DCs has been investigated in vitro and in vivo.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Mice

C57BL/6, BALB/c, CBA/Ca, AKR/J, B10.A(2R), and B10.A(4R) mice, 6–10 wk of age, were purchased from Harlan Olac (Bicester, U.K.). MHC class II and {beta}2-microglobulin double-knockout mice (MHC0) (20) were bred at the Biological Services Unit, Hammersmith Hospital, and were donated by Dr. M. Merkenschlager. RAG–/– TCR-transgenic mouse strain F5 (CD8+ T cells specific for influenza strain A/HK/8/68 nucleoprotein-derived peptide NT68/H-2Db, sequence ASNENMDAM), were obtained from Dr. D. Kioussis (National Institute for Medical Research, London, U.K.). Spleens from RAG–/–, TCR-transgenic OT-II mice (CD4+ T cells specific for OVA323–339/H-2Ab) were obtained from mice kept in a pathogen-free environment at GlaxoSmithKline Research and Development (Stevenage, U.K.). Mice of the same sex were used within experiments.

DC cultures

Mouse bone marrow-derived DCs (BMDCs) were generated as previously described (21), with slight modifications. Briefly, bone marrow was flushed from femurs, passed through a 200-µm pore size mesh to remove fibrous tissue, and RBCs were lysed using ACK buffer. Cells were cultured at 106 cells/ml in RPMI 1640 medium (Invitrogen Life Technologies, Paisley, U.K.) supplemented with 10% FCS, 2 mM glutamine, 50 µM 2-ME, 100 IU/ml penicillin, 100 µg/ml streptomycin, and 6 ng/ml mouse rGM-CSF (produced by Dr. M. Sims, GlaxoSmithKline Research and Development). On day 3 of culture, floating cells were gently removed, and fresh rGM-CSF-containing medium was added. On day 5 of culture, BMDCs were either left untreated or induced to mature by adding 1 µg/ml LPS (Sigma-Aldrich, Gillingham, U.K.) to the cultures. After an overnight incubation, nonadherent cells and loosely adherent proliferating BMDC aggregates were collected, washed, and replated for 1 h at 37°C to remove contaminating macrophages. Subsequently, resulting cell populations were enriched for CD11c-positive BMDCs by positive selection after incubation with anti-CD11c-coated magnetic microbeads (Miltenyi Biotec, Bisley, U.K.) and passing the bead-bound BMDCs through a separation column (MS+ columns; Miltenyi Biotec) placed on the separation unit, according to manufacturer’s instructions.

Endothelial cell (EC) cultures

ECs derived from hearts of C57BL/6 and CBA/Ca mice were cultured as previously described (22) in DMEM (Invitrogen Life Technologies, Gaithersburg, MD) supplemented with 20% heat-inactivated FCS, 2 mM glutamine, 100 IU/ml penicillin, 100 µg/ml streptomycin, 1 mM sodium pyruvate, 20 mM HEPES, 1% nonessential amino acids, 50 µM 2-ME, 150 µg/ml EC growth supplement (Sigma-Aldrich), and 12 U/ml heparin in 2% gelatin-coated tissue culture flasks. At confluence, the ECs were detached from the culture flasks using a solution of 0.125% trypsin in 0.2% EDTA and passaged. For phenotypic analysis, the ECs were used between passages 4 and 10.

CFSE staining

Cells were resuspended at 5–10 x 106 cells/ml in a 2-µM solution of CFSE (Molecular Probes, Leiden, The Netherlands) in PBS and incubated at 37°C for 10 min in the dark. At the end of the incubation period, the cells were immediately washed once in cold PBS/8% FCS and twice more in cold PBS/2% FCS. In the case of DCs to be injected, cells were labeled with 5 µM CFSE and finally resuspended in PBS before injection.

Transfer cultures

CFSE-labeled C57BL/6 or CBA/Ca DCs were pulsed for 4 h with NT68 (10 µM) and OVA323–339 (20 µM) peptides, washed twice, and mixed at equal numbers (5 x 105) with BALB/c DCs. Cells were cocultured at 4 or 37°C in 24-well plates in 1 ml of complete RPMI 1640 medium for 20 h and harvested for Ab staining. In some experiments DCs were cocultured in the presence or the absence of titrated concentrations of DNP (1–100 µg/ml) or sodium azide (1–10 µM).

Confluent ECs were detached from the culture flasks by trypsin/EDTA treatment and added at 106 cells/flask (25 cm2; Nunc, Roskilde, Denmark) in 3 ml of complete EC culture medium supplemented, or not, with 1000 U of mouse rIFN-{gamma} (PeproTech, London, U.K.) for 96 h. Cells were recovered by trypsin/EDTA treatment, washed twice, stained or not, with CFSE, and replated at 2 x 105 cells/well in 24-well plates overnight to allow the formation of a monolayer. In some experiments the cells were pulsed for 4 h with NT68 and OVA323–339 peptides and washed in the wells three times with PBS before the addition of 3 x 105 BALB/c DCs/well. Cells were cocultured in 1 ml of complete RPMI 1640 medium for 20 h and collected by pipetting off nonadherent cells, followed by trypsinization of adherent ECs. For trans-well studies, peptide-pulsed labeled DCs or ECs were added to 0.4-µm pore size trans-well chambers (Costar, Cambridge, U.K.) inserted into wells containing unlabeled DCs. After 20 h of culture, cells in the lower wells were collected and analyzed by flow cytometric staining or were used as stimulators of T cells.

Flow cytometry

All the mAbs used, unless otherwise stated, were purchased from BD Pharmingen (Cowley, U.K.). For analysis of DC purity and phenotype, cells were washed in cold PBS supplemented with 2.5% FCS and 0.05% sodium azide. BMDCs were first incubated with an anti-CD16/CD32 (anti-FcR{gamma} III/FcR{gamma} II, clone 2.4G2) mAb for 10 min and subsequently double-stained for 30 min with a PE-conjugated anti-CD11c mAb (clone HL3) in conjunction with FITC-conjugated mAbs to MHC class II (anti-H-2Ab, clone AF6-120.1; anti-H-2Ad, clone AMS-32.1; anti-H-2Ak, clone 11-5.2), MHC class I (anti-H-2Kb, clone AF6-88.5; anti-H-2Kd, clone SF1-1.1), anti-H-2Kk (Caltag Medsystems, Silverstone, U.K.), CD80 (clone 16-10A1), CD86 (clone GL1), or CD40 (clone 3/23). In each case, an FITC-conjugated mAb of the same isotype as the marker-specific mAb was used in conjunction with the anti-CD11c mAb as a negative control. The purity of DCs was consistently between 80 and 90% CD11c-positive cells.

The purity of responder T cells was assessed by staining with FITC- or PE-conjugated mAbs to CD3 (clone 145-2C11), CD4 (clone RM4-5), CD8 (clone 53-6.7), and H-2Ad or H-2Ab. Adherent ECs were detached from the culture flasks with trypsin/EDTA and resuspended in the same FCS-containing buffer before staining with PE- or FITC-conjugated mAbs specific for H-2Ab, H-2Kb, H-2Db (clone KH95), H-2Ak, H-2Kk, CD80, CD86, CD40, or CD106 (clone 429). For analysis of MHC transfer in cultures of mixed cells, cells were incubated with the anti-FcR{gamma} mAb and stained with the specific PE-conjugated mAbs as indicated. Analyses were performed on a FACSCalibur flow cytometer (BD Biosciences, Mountain View, CA) using CellQuest acquisition and analysis software on cells gated for homogeneous forward scatter (FSC) and side scatter characteristics.

FACS

DCs in the mixed cultures were purified to no less than 99% purity from CFSE-labeled DCs or ECs on a FACStar cell sorter (BD Biosciences).

Preparation of responder T cells

Responder T cells were purified from splenocytes of normal BALB/c or TCR-transgenic F5 or OT-II mice. Briefly, cell suspensions were prepared by mashing spleens through a cell strainer, and RBC were lysed using RBC lysis buffer. Cell preparations were passed through a nylon wool column to remove most contaminating macrophages and B cells. For further purification of CD4+ or CD8+ T cells, remaining B cells, macrophages, NK cells, and CD8+ or CD4+ T cells were removed after incubation with a mixture of rat mAbs to B220 (clone RA3-6B2; BD Pharmingen), MHC class II (culture supernatant from MS/114.15.2 hybridoma), CD16/CD32 (clone 2.4G2; BD Pharmingen), and anti-CD8 (culture supernatant from 53-6.72 hybridoma) or anti-CD4 mAbs (clone YTS191; purified at GlaxoSmithKline Research and Development), respectively, followed by an incubation period with sheep anti-rat IgG-coated Dynabeads (Dynal Biotech, Oslo, Norway). The bead/mAb-bound cells were selected using a magnet, and the purified T cell populations were recovered from the fluid phase. The purity of the T cell preparations obtained was consistently >85%, as determined by flow cytometric analysis of cell phenotype as described above.

T cell proliferation assays

MLR. Purified irradiated (30 Gy) DCs (104) were used as stimulators of 105 purified CD4+ or CD8+ BALB/c T cells in triplicate wells of 96-well, flat-bottom plates. T cell proliferation was measured by [3H]thymidine incorporation after 5 days in culture (1 µCi/well for the last 18 h of culture; Amersham Biosciences, Little Chalfont, U.K.). Results are presented as the mean cpm of triplicate determinations ± SD.

Peptide-specific T cell responses. Purified TCR-transgenic OT-II CD4+ or F5 CD8+ T cells (2.5 x 104) were stimulated with titrated numbers of irradiated DCs in triplicate wells of 96-well plates. Alternatively, 105 responder R2.2 T cells (23) were stimulated with 2 x 104 irradiated DCs in the presence of increasing amounts (0.01–1 µM) of H-Y peptide. T cell proliferation was measured by [3H]thymidine incorporation after 3 days in culture. Results are presented as the mean cpm of triplicate determinations ± SD.

In vivo mouse model

To investigate the acquisition of MHC molecules in vivo, different experimental approaches were used (Fig. 1). First, 6- to 8-wk-old C57BL/6 mice were injected i.p. with 200 µl of PBS or 1000 U of rIFN-{gamma} in PBS to induce local inflammation. Forty-eight hours later, mice within each group received an i.p. injection of 2.5 x 106 CFSE-labeled immature or mature MHC0 DCs. Twenty hours after DC inoculation, mesenteric lymph nodes (MLNs) were collected for immunofluorescence staining (Fig. 1A).



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  		FIGURE 1. Experimental designs to investigate the acquisition of allogeneic MHC molecules in vivo. A, C57BL/6 mice were injected i.p. with 200 µl of PBS or 1000 U of rIFN-{gamma} in PBS to induce local inflammation. Forty-eight hours later, mice were injected with 2.5 x 106 CFSE-labeled immature or mature MHC0 DCs. Twenty hours after DC inoculation, MLNs were collected for immunofluorescent staining. B, B10.A(2R) or AKR/J mice were injected i.p. with PBS or CFSE-labeled mature DCs derived from B10.A(2R) or B10.A(4R) mice. Twenty hours later, cell suspensions were prepared from MLNs, depleted of T and B cells by negative selection, and double-stained with a PE-conjugated anti-H-2E{alpha} and a Cy5-conjugated anti-CD8{alpha} mAbs and analyzed by flow cytometry. C, Recipient B10.A(2R) or B10.A(4R) mice were injected i.p. with 1000 U of rIFN-{gamma}. Forty-eight hours later, 107 CFSE-labeled mature DCs were transferred i.p. into i) matched (negative controls, B10.A(4R) DCs into 4R hosts), ii) H-2Ek-positive into H-2Ek-negative mismatched recipients (positive controls, B10.A(2R) DCs into 4R hosts), and iii) H-2Ek-negative into H-2Ek-positive mismatched recipients (experimental group, B10.A(4R) DCs into 2R hosts). Thirty hours after the transfer, the recipients were killed, and cell suspensions were prepared from the spleen and pooled abdominal lymph nodes. CFSE-positive cells were selected by FACS and used in an in vitro proliferation assay as stimulators of the HY-specific and H-2Ek-restricted R2.2 T cell clone.

 
In the second method, naive B10.A(2R) or AKR/J mice were injected i.p. with PBS or CFSE-labeled mature DCs derived from B10.A(2R) or B10.A(4R) mice. Twenty hours later, cell suspensions were prepared from MLNs and depleted of T and B cells by negative selection using rat mAbs specific for B220 and CD3 and sheep anti-rat IgG-coated Dynabeads as described above. Cells were double-stained with a PE-conjugated anti-H-2E{alpha}k (clone 14-4-4S; BD Pharmingen) and a Cy5-conjugated anti-CD8{alpha} (clone 53-6.7; BD Pharmingen) mAb and were analyzed by flow cytometry (Fig. 1B).

Finally, mature DCs were prepared from B10.A congenic strains B10.A(2R) and B10.A(4R), labeled with CFSE, and transferred i.p. into matched (negative controls, B10.A(4R) DCs into 4R hosts) and mismatched recipients (positive controls, B10.A(2R) DCs into 4R hosts; experimental group, B10.A(4R) DCs into 2R hosts). Thirty hours after the transfer, the recipients were killed, and cell suspensions were prepared from the spleen and pooled abdominal lymph nodes. CFSE-positive cells were selected by FACS and used in an in vitro proliferation assay as stimulators of the HY-specific and H-2Ek-restricted R2.2 T cell clone (Fig. 1C).

Immunofluorescent staining of tissue sections

MLNs were removed, placed in compound-embedding medium (OCT; BDH, Dorset, U.K.), snap-frozen in liquid nitrogen, and stored at –80°C for no more than 96 h. Ten-micron cryostat sections were cut, collected on poly-L-lysine-coated slides (VWR International, Lutterworth, U.K.), and allowed to air-dry. Slides were stored at 4°C for 24 h before staining. Sections were fixed with cold acetone for 5 min, air-dried, and incubated with the appropriate dilutions in 5% FCS/PBS of rabbit anti-FITC (1/1000; DakoCytomation, Ely, U.K.) and biotinylated anti-H-2Ab (5µg/ml; clone AF6-120.1; BD Pharmingen) or anti-H-2Kb (5 µg/ml; clone AF6-88.5; BD Pharmingen) mAbs for 1 h. Isotype-matched mIgG2a (BD Pharmingen) and rabbit Ig (DakoCytomation) served as controls. The slides were washed three times (once for 1 min, once for 2 min, and once for 3 min) in PBS under stirring and then incubated with an FITC-conjugated swine anti-rabbit mAb (1/40; DakoCytomation) in conjunction with ExtrAvidin-tetramethylrhodamine isothiocyanate (ExtrAvidin-TRITC; 5 µg/ml; Sigma-Aldrich) in 5% FCS/PBS for 1 h. Slides were washed again four times (once for 1 min, once for 2 min, once for 3 min, and once for 4 min) and mounted in mounting medium (Citifluor, Kent, U.K.). Slides were visualized with a Coolview 12-cooled CCD camera (Photonic Science, Newbury, U.K.) mounted over an Axiovert S100 microscope equipped with Metamorph software (Zeiss, Welwyn Garden City, U.K.). Photomicrographs were taken under equal exposure conditions to obtain a permanent record. Final image processing was performed using Photoshop 5.0 (Adobe Systems, San Jose, CA).


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Transfer in vitro of surface MHC molecules between live allogeneic DCs

Based on the assumption that DCs are the most efficient donors of MHC molecules due to exosome release, we first examined the transfer of allogeneic MHC molecules in cocultures of mouse BMDCs. Both immature and LPS-treated, mature DCs were generated from three different mouse strains: BALB/c, C57BL/6, and CBA/Ca (see Table I). The populations obtained were then enriched (up to 90%) for CD11c-expressing DCs by positive selection using anti-CD11c-coated magnetic beads before being used in the cocultures. The phenotypic characterization of DCs by flow cytometric staining showed a marked up-regulation of MHC class I and class II molecules and the costimulatory ligands CD80, CD86, and CD40 in LPS-treated, as compared with nontreated, immature DCs (not shown).


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  		Table I. Mouse strains used

 
We devised a system to single out DC populations by labeling donor cells with an intracellular dye, CFSE, before coculture with allogeneic DCs. Labeling of the donor cells and subsequent analysis of CFSE-negative recipient cells excludes the possibility of mistakenly analyzing doublets comprising cells from the two input populations. DCs obtained from BALB/c mice (H-2d) were cocultured with CFSE-labeled C57BL/6 (H-2b) or third-party, control CBA/Ca DCs (H-2k). After coculture for 0 and 20 h, CFSE-negative BALB/c DCs were analyzed by flow cytometry using PE-conjugated mAbs specific for H-2Ab (Fig. 2, B–E, left panels) and H-2Kb (Fig. 2, B–E, right panels). For comparison, the expression of H-2Ab and H-2Kb on C57BL/6 cells, acting as donors of MHC molecules, is shown in Fig. 2A. The results showed acquisition of allogeneic H-2b MHC molecules by a significant number of BALB/c DCs. The efficiency of transfer was higher for MHC class II molecules than for MHC class I molecules. Mature DCs acquired higher levels of allogeneic MHC class II and class I molecules (Fig. 2E) compared with immature cells (Fig. 2C). Mature DCs were also much better donors of surface MHC molecules than immature DCs. For example, ~42% of immature acceptor BALB/c DCs became H-2Ab-positive when cocultured with mature C57BL/6 DCs compared with 10% when the donor cells were immature (Fig. 2C, left panel). This may reflect the higher levels of MHC expression on mature DCs. Similar results were obtained when the acceptor and donor DC strain combinations were inverted (data not shown). Labeled DCs were viable during the entire coculture period as measured by lack of staining using a PE-conjugated annexin V-specific mAb to measure early apoptosis in conjunction with phosphatidylinositol incorporation to assess induction of necrosis (not shown), suggesting that transfer of MHC molecules was not the result of uptake of dead or dying cells.



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  		FIGURE 2. Acquisition of intact allogeneic MHC molecules by DCs. A, The expression of self MHC class II (H-2Ab; left panel) and class I (H-2Kb; right panel) molecules on immature (thin lines) and mature (heavy lines) CFSE-labeled donor C57BL/6 DCs was assessed after staining with PE-conjugated specific mAbs. As a negative control, cells were stained with a PE-conjugated, isotype-matched, irrelevant mAb (filled profiles). B–E, Equal numbers of immature (B and C) or mature (D and E) BALB/c DCs (H-2d) were mixed with immature (thin lines) or mature (heavy lines) CFSE-labeled C57BL/6 DCs. After 0 h (B and D) or 20 h (C and E) of coculture, cells were stained with either a PE-conjugated anti-H-2Ab (left panels) or anti-H-2Kb mAb (right panels). As controls for the staining, BALB/c DCs were cocultured with third-party CFSE-labeled CBA/Ca DCs (H-2k; filled profiles) and stained with the same PE-conjugated mAb. All analyses shown on B–E were on populations with a homogeneous, single peak on FSC characteristics, which were CFSE-negative (F; region R1xR4; i.e., BALB/c DCs acting as acceptors of allogeneic H-2b MHC molecules). Numbers within figures correspond to percentages of BALB/c DCs expressing the H-2b MHC molecules after coculture with CFSE-labeled mature CBA/Ca or immature or mature C57BL/6 DCs, respectively.

 
Parallel experiments were performed in which the donor and acceptor DCs were physically separated by a 0.4-µm pore size semipermeable membrane that allows the traffic of soluble molecules and exosomes, but not of intact cells. After 20 h of culture, a considerable percentage of CFSE-negative DCs in the lower wells expressed the MHC of DCs in the upper wells. The transfer, however, was ~3-fold less efficient than when the cells were cocultured (data not shown), suggesting that direct cell-cell contact is not essential for MHC transfer between DCs. This correlates with the findings reported by Emerson and Cone (24) describing the shedding of mouse MHC molecules within membrane-derived lipid vesicles (exosomes).

Transfer efficiency is affected by temperature and levels of available ATP

To test whether the observed transfer of MHC molecules is a temperature- and/or an energy-dependent phenomenon, allogeneic DCs were cocultured at either 4 or 37°C in the presence or the absence of titrated concentrations of sodium azide (1 or 10 µM) or DNP (1, 10, and 100 µg/ml). Azide ions inhibit the mitochondrial respiratory chain, whereas DNP is an uncoupler of oxidative phosphorylation. In both cases, the synthesis of ATP is inhibited, thus decreasing the energy supply of the cell.

The results in Fig. 3A showed that the coculture of DCs at 4°C resulted in a notable decrease in molecule transfer. At 20 h, a mere 7.6% of DCs had acquired the H-2Ab molecule compared with 27.6% of DCs cocultured at 37°C (not shown). A reduction in the number of molecules acquired per cell was also observed, as demonstrated by the drop in mean fluorescence intensities (MFIs) at 4°C to over half that seen at 37°C (Fig. 3A). A similar pattern was observed after 40 h of coculture. These results confirm the originally reported observations showing that MHC shedding and absorption between splenic mouse cells were temperature-dependent (25, 26).



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  		FIGURE 3. Transfer of MHC molecules in vitro is temperature- and energy-dependent. Equal numbers of recipient BALB/c DCs (H-2d) were mixed with CFSE-labeled C57BL/6 DCs (H-2b) and cocultured at 4 or 37°C for different periods of time (0, 20, or 40 h; A). Alternatively, cells were cocultured at 37°C for 20 h in the presence or the absence of titrated concentrations of DNP (1, 10, and 100 µg/ml; B) or sodium azide (1 and 10 µM; C). After coculture, cells were harvested and stained using a PE-conjugated anti-H-2Ab mAb. Cells cocultured under normal conditions (37°C in medium alone) and stained with a PE-conjugated, isotype-matched, irrelevant mAb served as negative controls.

 
The presence of either azide or DNP in the cocultures also caused a significant decrease in the efficiency of allogeneic MHC capture by DCs (Fig. 3, B and C). Interestingly, the azide appeared to exert a greater inhibitory effect on transfer, with different concentrations seemingly irrelevant. Only 3.6–3.8% of recipient DCs acquired the molecules in the presence of azide compared with 27.6% under normal conditions (not shown). MFIs were also 4- to 5-fold lower, indicating that far fewer molecules per cell had been transferred (Fig. 3C). Although not as marked as with azide, addition of DNP also had some effect (Fig. 3B). Overall, transfer was between 7.4 and 13.6% of DCs, with a 2- to 3-fold drop in MFIs.

Acquired foreign MHC-peptide complexes allow recognition by T cells

To confirm that the acquired allogeneic MHC molecules were fully functional, acceptor mature BALB/c DCs, displaying both endogenous H-2d and acquired H-2b molecules, were used as allostimulators of polyclonal BALB/c CD4+ and CD8+ T cells and H-2b-restricted peptide-specific TCR transgenic T cells from F5 and OT-II mice. The donor C57BL/6 DCs had previously been pulsed with the influenza nucleoprotein peptide NT68 and the OVA323–339 peptide presented by H2-Db and H-2Ab, respectively.

CFSE-negative DCs were purified from the cocultures by cell sorting to no less than 99% of purity (Fig. 4A). Cells that had acquired foreign MHC molecules in the trans-well system were also recovered and used as stimulators of T cells. BALB/c DCs, cultured in medium alone, were used as negative controls. The CFSE-labeled C57BL/6 DCs, selected by cell sorting from the mixed cultures, served as positive controls. As shown in Fig. 4, both polyclonal BALB/c (Fig. 4B) and peptide-specific H-2b-restricted (Fig. 4C) CD8+ (left panels) and CD4+ T cells (right panels) proliferated in response to the BALB/c DCs derived from the cocultures, indicating that they acquired fully functional MHC-peptide complexes from C57BL/6 DCs. Although the polyclonal and transgenic CD8+ T cells proliferated less in response to the BALB/c DCs that had acquired the H-2b molecules than to control C57BL/6 DCs (Fig. 4, B and C, left panels), the H-2b-expressing BALB/c cells induced proliferation comparable to that seen with C57BL/6 DCs by the CD4+ T cell populations (Fig. 4, B and C, right panels). In accordance with the levels of MHC transfer observed by flow cytometry, the BALB/c DCs harvested from the trans-well cultures showed a lower capacity to stimulate T cells, especially CD8+ T cells. As expected, stimulation of T cells with BALB/c DCs cultured in medium alone resulted in negligible T cell proliferation.



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  		FIGURE 4. T cell recognition of foreign MHC-peptide complexes acquired by DCs. Equal numbers of LPS-treated, mature BALB/c DCs (H-2d) were mixed with mature CFSE-labeled C57BL/6 DCs (H-2b) that had previously been pulsed for 4 h with NT68 peptide and OVA323–339 peptide. Alternatively, these allogeneic DCs were physically separated using a 0.4-µm trans-well chamber. As controls, BALB/c DCs were also pulsed with the NT68 and OVA323–339 peptides and cultured in medium alone. After 20 h of culture, CFSE-negative BALB/c DCs in the mixed cultures were separated from CFSE-expressing C57BL/6 DCs by FACS (A), and BALB/c DCs were recovered from the lower wells of the trans-well cultures or the control single cultures. DCs (104) were used as stimulators of 105 purified BALB/c CD8+ (B, left panel) and CD4+ T cells (B, right panel). Alternatively, various numbers of DCs were used as stimulators of 2.5 x 104 TCR-transgenic, RAG–/– NT68-specific F5 CD8+ T cells (C, left panel) and OVA323–339-specific OT-II CD4+ T cells (C, right panel). T cell proliferation was measured by [3H]thymidine incorporation after 5 days (B) or 3 days (C) in culture. Results are presented as the mean cpm of triplicate determinations ± SD.

 
DCs acquire intact MHC molecules from allogeneic ECs

Given the importance of the endothelium in the regulation of leukocyte migration into grafted tissues (27), we examined the capture of foreign MHC molecules by DCs upon interaction with ECs. ECs were isolated from hearts of C57BL/6 and CBA/Ca mice and cultured as described in Materials and Methods. The cells were treated, or not, with 330 U/ml mouse rIFN-{gamma} for 96 h, and their phenotype was analyzed by flow cytometry. The rIFN-{gamma} treatment significantly increased the expression of MHC class I molecules, which was very low in nontreated ECs (Fig. 5A). As previously reported (22), constitutive low expression of CD80 molecules was observed, whereas CD86 and CD40 were negative. The expression levels of these molecules were not altered by rIFN-{gamma} treatment (data not shown).



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  		FIGURE 5. Transfer of MHC molecules between DCs and ECs is unidirectional. A, Immature (left panels) or LPS-treated, mature (right panels) BALB/c DCs (H-2d; 3 x 105) were cultivated on a monolayer of nonirradiated, CFSE-labeled allogeneic C57BL/6 ECs (H-2b). ECs had previously been cultured in medium alone (Aa) or in medium containing 330 U/ml rIFN-{gamma} (Ab) for 96 h. As controls for the staining, BALB/c DCs were cultured on a monolayer of third-party, CFSE-labeled, rIFN-{gamma}-treated CBA/Ca ECs (H-2k; A-c). After 20 h of coculture, cells were recovered from the culture wells and stained with a PE-conjugated anti-H-2Db mAb. B, CFSE-labeled mature C57BL/6 DCs (H-2b; 3 x 105) were cultured on a monolayer of nonirradiated allogeneic CBA/Ca ECs (H-2k) that had previously been cultured in medium alone (left panels) or in medium containing 330 U/ml mouse rIFN-{gamma} (right panels) for 96 h. After 20 h of coculture, cells were recovered from the culture wells and analyzed by flow cytometry using PE-conjugated mAbs specific for H-2Kb (Ba), H-2Ab (Bb), or CD40 (Bc).

 
ECs were labeled with CFSE and pulsed with the NT68 peptide before being cocultured with allogeneic immature or mature BALB/c DCs. A fraction of both immature and mature DCs became H-2Db-positive after coculture (Fig. 5Ab). No H-2Db expression was detected on the DCs cultured with untreated C57BL/6 ECs (Fig. 5Aa) or third-party CBA/Ca ECs (Fig. 5Ac). No H-2Db acquisition was observed if the ECs and DCs were separated in a trans-well system (not shown).

To evaluate whether ECs were also able to acquire foreign MHC molecules and DC-specific markers after interaction with the DCs, a similar coculture experiment was performed using CFSE-labeled C57BL/6 DCs as donors and CBA/Ca ECs as acceptors of MHC class I (Fig. 5Ba), MHC class II (Fig. 5Bb), or CD40 molecules (Fig. 5Bc). As shown in Fig. 5B, ECs did not acquire any of these molecules from mature DCs regardless of the activation state of the ECs. These results show that the transfer of MHC molecules between ECs and DCs is unidirectional.

DCs acquire foreign MHC molecules in vivo

To determine whether this phenomenon could be observed in vivo, we designed an experimental system by which we could visualize in vivo the acquisition of recipient MHC molecules by transferred DCs. We injected C57BL/6 mice with PBS or 1000 U of mouse rIFN-{gamma} i.p. to induce local inflammation in the peritoneal cavity. Injection of rIFN-{gamma} at this low dose does not cause systemic inflammation (M. J. James and F. M. Marelli-Berg, unpublished observations). Forty-eight hours later, immature or mature BMDCs, prepared from MHC0 mice of the same genetic background as the recipient mice were labeled with CFSE and injected i.p. (Fig. 1A).

Twenty hours later, the acquisition of MHC molecules by migrated DCs was assessed by immunostaining of acetone-fixed MLN sections (Fig. 6). Because acetone fixing significantly affects CFSE fluorescence, injected DCs were visualized with a FITC-specific primary mAb, followed by a FITC-conjugated secondary mAb (Fig. 6, left panels). The strongest CFSE staining was seen in MLNs of mice pretreated with rIFN-{gamma} (Fig. 6, Ad and Bd) and in mice injected with mature DCs (Fig. 6, Ag and Bg). This suggests that already mature DCs or immature DCs that encountered an inflammatory environment migrated out of the peritoneal cavity into secondary lymphoid organs. Recipient cells, expressing either H-2Ab (Fig. 6A, middle panels) or H-2Kb (Fig. 6B, middle panels) molecules were visualized with specific biotinylated mAbs, followed by ExtrAvidin-TRITC. Optically merged fluorescence images (Fig. 6, right panels) showed the colocalization of CFSE and recipient MHC molecules on migrated injected DCs, indicating that DCs can effectively acquire foreign MHC molecules in vivo.



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  		FIGURE 6. Trafficking DCs acquire intact foreign MHC molecules in vivo. CFSE-labeled immature (rows 1, 2, 4, and 5) or mature MHC0 DCs (rows 3 and 6) were injected into PBS- (rows 1, 3, 4, and 6) or rIFN-{gamma}-injected syngeneic C57BL/6 mice (rows 2 and 5). Tissue sections were prepared from MLNs 20 h later, fixed with acetone, incubated with mAbs specific for FITC and either H-2Kb or H-2Ab molecules, and analyzed by fluorescence microscopy. CFSE-labeled injected DCs were visualized by FITC-specific primary mAb, followed by a FITC-conjugated second mAb and displayed as green staining (left panels). H-2Ab (A, middle panels) and H-2Kb molecules (B, middle panels) were visualized using specific biotinylated mAbs plus ExtrAvidin-TRITC as a secondary reagent and displayed as red staining (middle panels). Double-staining analysis (right panels) showed the coexpression, displayed as yellow staining, of host MHC molecules and CFSE on injected DCs.

 
It could be argued, however, that some of the MLN-resident recipient cells might have become CFSE-positive after the phagocytosis of injected CFSE-labeled DCs that died after arriving in the lymph nodes, as suggested by Inaba et al. (28), or were killed by NK cells, as has been recently shown for allogeneic B cells (29). To address this possibility, we transferred CFSE-labeled, purified mature B10.A(4R) BMDCs i.p. into either MHC class I-matched, class II-mismatched B10.A(2R) recipient mice (see Table I) or MHC class I- and MHC class II-mismatched AKR/J recipients to control for host NK cell-mediated killing of injected DCs (Fig. 1B). As controls, B10.A(2R) mice were injected with PBS or transferred with CFSE-labeled syngeneic mature DCs. Twenty hours after DC transfer, we analyzed the acquisition of host MHC class II molecules (H-2E{alpha}) by injected B10.A(4R) BMDCs that had trafficked to the MLNs by flow cytometry.

For analysis, we took into consideration the observation made by several different groups (29, 30, 31) that in the mouse only the CD8{alpha}+ DCs are capable of phagocytosing dying cells. Because the injected BMDCs were CD8{alpha}– (not shown), the expression of H-2E{alpha} by CD8{alpha}–CFSE+ DCs must be the result of acquisition of host MHC class II molecules by transferred H-2E{alpha}-deficient BMDCs. If, in contrast, MLN-resident host CD8{alpha}+ DCs became CFSE-positive, this would indicate the uptake of dead injected BMDCs.

We double-stained the cells prepared from MLNs with mAbs specific for H-2E{alpha} and CD8{alpha}. For analysis, we gated on H-2E{alpha}-positive cells with homogeneous FSC characteristics and analyzed the expression of CD8{alpha} and CFSE (Fig. 7). The results showed that injected CFSE-labeled BMDCs migrated to MLNs (Fig. 7, A–C). These migrated BMDCs remained CD8{alpha}–, as assessed by analysis of BMDCs that had been injected into a syngeneic recipient (Fig. 7B). Furthermore, transferred H-2E{alpha}-deficient BMDCs were able to capture MHC class II molecules expressed by recipient cells (Fig. 7, C and D, left panels). When BMDCs were transferred into MHC class I-matched recipients, most injected DCs became H-2E{alpha}-positive and remained CD8{alpha}– (Fig. 7C, right panel), showing that phagocytic CD8{alpha}+ recipient DCs did not uptake CFSE-labeled BMDCs in detectable numbers. However, when the B10.A(4R) BMDCs were injected into MHC class I-mismatched AKR/J mice, a significant fraction of MLN-resident CD8{alpha}+ DCs expressed CFSE (Fig. 7D, right panel), suggesting that a proportion of transferred CFSE-labeled BMDCs had been killed by host NK cells and phagocytosed by CD8{alpha}+ DCs. Nevertheless, the majority of CFSE-positive, H-2E{alpha}-positive cells were CD8{alpha}–, indicating H-2E{alpha} acquisition by the injected BMDCs.



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  		FIGURE 7. DCs acquire MHC molecules from allogeneic cells in vivo. B10.A(2R) mice were injected i.p. with PBS (A) or 2.5 x 106 CFSE-labeled mature BMDCs prepared from syngeneic B10.A(2R) (B) or MHC class II-mismatched B10.A(4R) mice (C). To control for host NK cell activity, AKR/J mice served as recipients of B10.A(4R) BMDCs (D). Twenty hours after BMDC transfer, MLN cells from injected mice were depleted of T and B cells using lineage-specific rat mAbs (to CD3 and B220, respectively) and sheep anti-rat Dynabeads. Cells were then double-stained with a PE-conjugated anti-H-2E{alpha} mAb and a Cy5-conjugated anti-CD8{alpha} mAb and analyzed by flow cytometry. All analyses shown are of cell populations with a homogeneous, single peak on FSC characteristics. Cells positive for H-2E{alpha} were gated (region R in left panels, see A) and analyzed for the expression of CFSE and CD8{alpha} (right panels).

 
Finally, to test for the functionality of in vivo acquired MHC molecules, injected CFSE-labeled B10.A(4R) DCs, which had acquired allogeneic H-2Ek molecules upon transfer into B10.A(2R) recipients in vivo (Fig. 8A), were purified from spleens and regional lymph nodes by cell sorting (Fig. 1C). The sorted, CFSE-positive cells were pulsed with an H-Y peptide and used as stimulators of the H-Y-specific, H-2Ek-restricted, R2.2 T cell clone. The results of the proliferation assay in Fig. 8B show that these DCs induced a proliferative response of R2.2 cells comparable to that seen after their stimulation with H-2Ek-positive B10.A(2R) DCs. In contrast, no proliferation was observed if injected B10.A(4R) DCs were purified from syngeneic H-2Ek-negative B10.A(4R) mice.



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  		FIGURE 8. In vivo acquired allogeneic MHC molecules are fully functional. A, B10.A(2R) and B10.A(4R) DCs were labeled with 5 µM CFSE and injected i.p (107 cells/mouse) 48 h after local inflammation was induced in the recipients by i.p. IFN-{gamma} (1000 U/mouse). The injected B10.A(2R) DCs were 60% H-2Ek positive after in vitro maturation. Three groups of mice were prepared: B10.A(4R) DCs into 4R hosts, B10.A(2R) DCs into 4R hosts, and B10.A(4R) DCs into 2R hosts. Thirty hours after the transfer, the recipients were killed, and spleen and pooled MLN cell suspensions were prepared for flow cytometric analysis. B, CFSE-positive cells were isolated by cell sorting and used as stimulators of the R2.2 T cell clone. Responder T cells (1 x 105) were stimulated for 3 days with 2 x 104 irradiated stimulators in the presence of increasing amounts of H-Y peptide. T cell proliferation was measured by [3H]thymidine incorporation after 3 days in culture. Results are presented as the mean cpm of triplicate determinations ± SD.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
The data presented in this study illustrate the capacity of DCs to acquire intact MHC:peptide complexes from other cells and present them to T cells both in vitro and in vivo. These findings have major implications for the evolution of alloimmune responses after transplantation and create a mechanism for physically linking the direct and the indirect pathways of allorecognition, thereby offering an explanation for what has previously appeared to be unlinked help and unlinked suppression.

DCs were able to acquire functional foreign MHC:peptide complexes not only from other DCs, but also from activated ECs, as assessed by flow cytometry and the ability of DCs harvested from cocultures to stimulate allo- and peptide-specific CD4+ and CD8+ T cells. Importantly, both immature and mature DCs were capable of acquiring allogeneic MHC molecules with similar ability, indicating that this phenomenon could take place equally in peripheral and lymphoid tissues, where DCs of different maturational states reside. The mechanism by which the transfer between DCs occurs seems to involve close cell contact between the interacting DCs, a unique attribute of this kind of APC. Nevertheless, we and others (16, 19, 24) have shown that this transfer is also possible, although with less efficiency, by the shedding by DCs of MHC molecules either as soluble proteins or incorporated into exosomes.

We have also shown in this study that an important source of allogeneic MHC:peptide complexes could be ECs, which become activated during the process of inflammation that usually follows the transplantation of vascularized grafts or during infections. It is well known that the trafficking of DCs through the endothelium plays an important role in DC maturation (32). It might well be that during this process DCs acquire MHC molecules from ECs, thus augmenting their antigenicity. Interestingly, DC-EC contact was essential for the transfer of MHC molecules from ECs to DCs, even though the ectocytosis of plasma membrane constituents by ECs has been reported (33). Conversely, ECs were not able to acquire MHC or costimulatory molecules from DCs, confirming previous observations that this is an exclusive characteristic of DCs (16, 17, 18) and activated T cells (34, 35, 36). In the case of DCs, this may reflect their shape and motility, with large membrane processes that facilitate cell clustering and intercellular interactions.

Most importantly, we have shown that the acquisition of intact foreign MHC molecules by DCs occurs in vivo. The encounter by DCs of an inflammatory environment induced the migration of immature DCs out of the site of injection, whereas already mature DCs showed the capacity to do so without inflammation. Immunofluorescent staining of injected MHC double-knockout DCs showed that CFSE-positive cells expressed the MHC class I and class II molecules of recipient cells. It could not be ruled out, however, that migrated DCs were short-lived and might have been endocytosed by recipient DCs resident in T cell areas of MLNs. To minimize this possibility, we performed the analysis 20 h after transfer, when live migrated DCs can still be detected in the nodes (28). Alternatively, injected MHC0 DCs might have been killed by host NK cells.

The observation that the phagocytosis of dead cells in vivo by mouse lymphoid, organ-resident DCs is restricted to DCs that express the CD8{alpha} dimmer (29, 30, 31) allowed us to discriminate between the acquisition of foreign MHC class II molecules by injected MHC class II-, CD8{alpha}-deficient DCs and the uptake of dead or dying injected cells by recipient CD8{alpha}+ DCs. The results showed that MLN-resident CD8{alpha}+ DCs can indeed phagocytose cells that are killed by NK cells (Fig. 7D). However, even in an MHC class I-mismatched donor/recipient combination, most injected BMDCs remained CD8{alpha}-negative and showed acquisition of host MHC class II molecules. In this context, it is possible that the interaction with ECs that mediates the traffic of DCs into lymphoid tissues played a significant role in this acquisition. Finally, we were able to show that the MHC molecules acquired in vivo were fully functional, in that the injected DCs, purified by sorting for CFSE-positive cells from spleens and lymph nodes of recipient animals, induced proliferation of Ag-specific T cells whose Ag specificity is restricted by the acquired MHC allele.

These findings could be of substantial importance in transplantation immunology. The existence of both the direct and indirect pathways of alloantigen recognition during transplantation is well established. However, controversy still exists regarding the relative contributions of these two pathways to graft rejection. Historically, the direct pathway has been predominantly associated with the early onset of the alloresponse and acute rejection. The anti-donor alloreactive T cells, being naive or resting memory T cells, must be activated in lymphoid tissue (37). Consequently, the activation of direct pathway T cells has been associated with the presence of migrating donor-derived DCs, which are available only during the first few weeks after transplantation. Indeed, we, and others, have reported that the frequency of T cells with direct anti-donor allospecificity declines with time in many patients (38, 39, 40). One explanation for this is that alloreactive T cells are rendered unresponsive by encounter with costimulation-deficient parenchymal cells of the graft. However, the data described in this study raise an alternative possibility, namely, that the decrease in anti-donor frequency is due to activation-induced cell death as a result of continuing direct pathway presentation by trafficking recipient DCs. These two possibilities are not mutually exclusive. One prediction of the semidirect pathway proposed in this study is that there should be an equal decline in direct pathway anti-donor T cells in the naive and the memory subset of peripheral T cells. This prediction is currently being tested.

Animal models support a role for the indirect pathway in both acute and chronic rejection. The indirect pathway alone has been shown to be sufficient to elicit allograft destruction in the absence of direct allorecognition (12). The same donor DCs that migrate from the graft to lymphoid organs may constitute a source of alloantigens for priming indirect pathway T cells in the early period of the response. Once the donor APCs are replaced by recipient APCs over time, indirect effector mechanisms are likely to be most effective, and therefore, they are especially important in chronic rejection. Elevated frequencies of T cells with indirect anti-donor specificity have been detected in patients with chronic heart, kidney, and lung transplant rejection (41, 42, 43, 44).

The semidirect pathway could explain the apparent violation of the immunological principle of linked help observed by Auchincloss and colleagues (13) after the transplantation of MHC class II-negative skin grafts. This generally accepted concept postulates that CD4+ and CD8+ T cells must recognize Ag on the same APC for help to be effective and was also described as the three-cell model for the generation of CTLs (45). Auchincloss and colleagues proposed a four-cell model by which CD8+ T cells, activated directly by donor cells, receive help from CD4+ T cells that are primed indirectly by recipient APCs. According to the semidirect hypothesis proposed in this study these events can be accommodated within a three-cell model if the trafficking recipient DCs could acquire the allogeneic MHC class I molecules from donor tissues and simultaneously stimulate indirect pathway CD4+ and direct pathway CD8+ T cells. Similarly, this pathway could explain the observations of apparently unlinked suppression that have been reported in experimental transplant models (14, 15).

Although these results have been discussed in the context of transplantation, the phenomenon of peptide:MHC molecule acquisition by DCs may well have an important role in immunity against pathogens. DCs that traffic through virally infected tissues may well use this pathway to acquire, and subsequently present, viral peptide:self MHC complexes from virus-infected tissue cells in addition to processing and presenting peptides from free or cell-associated viral proteins. This would have the advantage of guaranteeing the display in the draining lymph nodes of the viral peptide:MHC complexes that are most highly expressed by target cells.


   Acknowledgments
 
We thank Prof. Andrew George and Dr. Giovanna Lombardi for their helpful comments.


   Footnotes
 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1 This work was supported by a grant (DMIMM PN0559) from GlaxoSmithKline Research and Development. Back

2 Address correspondence and reprint requests to Dr. Robert I. Lechler, Department of Immunology, Imperial College, Hammersmith Hospital, Du Cane Road, London, U.K. W12 ONN. E-mail address: r.lechler{at}ic.ac.uk Back

3 Abbreviations used in this paper: DC, dendritic cell; BMDC, bone marrow-derived DC; DNP, dinitrophenol; EC, endothelial cell; FSC, forward scatter; MFI, mean fluorescence intensity; MLN, mesenteric lymph node; TRITC, tetramethylrhodamine isothiocyanate. Back

Received for publication December 4, 2003. Accepted for publication August 5, 2004.


   References
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 Introduction
 Materials and Methods
 Results
 Discussion
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