1 Departments of Pathology and 2 Medicine, College of Physicians and Surgeons of Columbia University, 630 West 168 Street, New York, NY 10032, USA
Correspondence to: N. Suciu-Foca; E-mail: ns20{at}columbia.edu
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Abstract |
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Keywords: inhibitory receptors, direct allorecognition, regulatory T cells, tolerance, transplantation
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Introduction |
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Infiltrates of allospecific CD4+ T cells, activated via the direct and indirect allorecognition pathways, as well as CD8+ T cells are found in human heart allografts during episodes of acute rejection (13). However, such infiltrates do not always expand and/or cause injury to the myocardium (4). Frequently, T cell infiltrates regress spontaneously without rejection therapy, suggesting that some regulatory mechanisms are at play (2,4).
Immunoregulatory mechanisms such as those mediated by naturally occurring or antigen-induced regulatory T cells have been shown to inhibit the growth and functional maturation of alloreactive T effector cells (511). There is evidence that naturally occurring CD4+CD25+ regulatory T cells (natural TR) prevent autoimmunity and mediate transplantation tolerance in rodents (58) and humans (911). These cells are characterized by the constitutive expression of CD25, the interleukin (IL) 2 receptor alpha, and forkhead transcription factor FOXP3 (1214). Natural TR inhibit conventional CD4+CD25 T cell activation in response to antigens or mitogens in an antigen-nonspecific, MHC non-restricted and cytokine-independent manner via T cellT cell interaction (514). However, their mechanism of action is still unclear.
A distinct population of antigen-primed T cells, characterized by their CD8+CD28phenotype and lack of cytotoxic activity has also been shown to display regulatory functions in human transplant recipients and in a murine autoimmune disease model (15,16). As opposed to natural TR cells, human CD8+CD28 T suppressor (TS) cells are antigen specific, MHC class I-restricted, and interact directly with antigen-presenting cells (APC) (1720). TS render APC tolerogenic, inducing the downregulation of costimulatory molecules and upregulation of the inhibitory receptors, immunoglobulin-like transcripts (ILT)3 and ILT4 (15). ILT3 and ILT4 display long cytoplasmic tails containing immunoreceptor tyrosine-based inhibitory motifs (ITIM), which mediate inhibition of cell activation by recruiting tyrosine phoshatase SHP-1 (21,22). Both of these molecules were invariably co-expressed on dendritic cells (DC) rendered tolergenic by exposure to TS or to certain cytokines such as IL-10 and interferon (IFN) alpha (23).
Studies of rejection-free recipients of heart transplants have demonstrated the presence of circulating CD8+CD28 T cells which inhibit CD40L-induced upregulation of costimulatory molecules and enhance ILT3 and ILT4 expression on donor APC (15,24). Since TS require direct contact with professional APC of donor origin in order to inhibit T cell allorecognition via the direct pathway (15,1719), it is unclear how they maintain quiescence once donor APC have migrated out of the graft. However, it was recently demonstrated that in mice non-professional APC, such as graft endothelial cells (EC), directly activate cytotoxic CD8+ T cells (Tc), triggering acute rejection in the absence of indirect allorecognition (25). We have explored the hypothesis that the interaction of TS with graft EC results in suppression of the direct pathway of allorecognition and maintenance of quiescence in human heart allograft recipients.
We provide evidence that allospecific CD8+CD28 human TS express FOXP3 and induce upregulation of ILT3 and ILT4 in EC, converting them to a tolerogenic state. The interaction between allospecific CD8+CD28 human TS and EC is bi-directional since tolerogenic ILT3+ILT4+ EC induce the in vitro differentiation of CD8+CD28 TS in unprimed populations of human CD8+ T cells. We found that allospecific CD8+CD28 FOXP3+ T cells which induce the upregulation of inhibitory receptors in EC carrying donor HLA class I antigens are present in the circulation of rejection-free patients.
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Methods |
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Endomyocardial biopsies
Endomyocardial biopsies were performed by standard transjugular approach weekly for the first month and then at progressively longer intervals to a baseline schedule of every 6 months. A minimum of four biopsy fragments were fixed in 4% buffered formalin, paraffin embedded, and multiple hematoxylin and eosin stained sections from three levels in the block were examined. An additional fragment was frozen for RTPCR of ILT3 and ILT4 and/or immunohistochemical studies. Histologic grades were assigned according to the criteria of the International Society for Heart Transplantation (ISHLT) (4). Acute allograft rejection was defined as ISHLT grades 3A or higher.
Immunohistochemistry
For detection of ILT4 expression, endomyocardium sections from frozen biopsies were fixed for 10 min at room temperature in PBS with 4% paraformaldehyde. Sections were then washed with 0.1% saponin in PBS and incubated with rat anti-human ILT4 IgG for 1 h. Sections were next incubated with goat anti-rat IgG conjugated with biotin followed by streptavidinhorseradish peroxidase (HRP). Endogenous peroxidase activity was blocked with 0.3% H2O2 prior to incubation of HRP. Tissue biotin was blocked by treatment with avidin and then biotin (Dako, Carpinteria, CA). The color was developed with DAB (diaminobenzidine; Dako). Samples stained only with goat anti-rat IgG were used as controls.
Molecular typing of HLA class I and class II antigens
HLA genotypes of transplant recipient/donor pairs and healthy blood or EC donors were determined by PCR with sequence specific primers (SSP) using commercially available kits (One Lambda, Los Angeles, CA).
Generation of alloreactive T cell lines (TCLs) and T cell clones (TCCs)
Peripheral blood mononuclear cells (PBMCs) from healthy volunteers were separated by FicollHypaque centrifugation. Responding PBMCs (1 x 106/ml) were stimulated in 24-well plates with irradiated (1600 rad) monocyte-derived DC (1 x 105/ml) obtained from allogeneic PBMCs. Cells were cultured for 7 days in complete medium [RPMI 1640 supplemented with 10% fetal bovine serum (Atlas Biologicals, Fort Collins, CO), 2 mM 1-glutamine and 50 µg/ml of gentamycin (Gibco-BRL, Grand Island, NY)]. After 7 days, responding cells were collected, washed and rechallenged with the original stimulating cells. Three days later, recombinant human (rh) IL-2 (Boehringer Mannheim, Indianapolis, IN) was added (10 U/ml) and the cultures were expanded for an additional 4 days.
On day 14, CD8+ T cells were separated using CD8 isolation kit II (Miltenyi Biotech, Auburn, CA). CD8+CD28 T cells were obtained from the CD8+ T cell suspension by depletion of CD28+ cells using goat anti-mouse IgG beads (Dynal, Lake Success, NY) coupled with mAb to CD28 (Becton Dickinson). The purity of the CD8+CD28 T cell suspension was >98%.
CD8+CD28 T cells from TCL were cloned by limiting dilution at 0.5 T cells per well in 96-well U-bottom plates (Costar, Corning Inc., Corning, NY). Irradiated (3000 rads) PBMC (1 x 105) and lymphoblastoid JY cells (2 x 104) (from ATCC, Manassas, VA) were added to each well. Cultures were grown in RPMI 1640 medium containing 5% pooled human serum (Sigma Chemical, St Louis, MO), and rh IL-2 50 U/ml (Boehringer Mannheim) and 1 µg/ml phytohemagglutinin-L (PHA-L; Sigma). Medium without PHA-L was replenished at 5-day intervals. Proliferating T cell clones were expanded and tested for suppressor activity.
Generation of monocyte-derived DC
Monocytes were obtained from PBMCs using a Monocyte Negative Selection Kit (Dynal, Lake Success, NY). Immature DC were generated by culturing monocytes in 6-well plates at a concentration of 2 x 106 cells per well for 7 days with granulocyte macrophage colony stimulating factor (GM-CSF) (R&D Systems, Minneapolis, MN) and interleukin (IL)-4 (R&D Systems), as previously described (15).
Endothelial cell culture
Human EC (HUVEC and HAEC) from Cambrex (Walkersvillle, MD) were cultured in EC growth medium (EGM BulletKit, Cambrex, Walkersville, MD) containing 2% bovine calf serum, 10 µg/ml of recombinant human epithelial growth factor (rh EGF), 1.0 mg/ml hydrococortisone, 3 mg/ml bovine brain extract (BBE), 100 U/mL penicillin/streptomycin, and 50 µg/mL gentamicin. For functional assays the EC were used between the second and fourth passages. To modulate ECs, recombinant human interferon (IFN)- (100 ng/ml) or tumor necrosis factor (TNF)-
(50 ng/ml) or a mixture of IL-10 (10 ng/ml) and IFN-
(1000 U/ml) (R&D systems, Minneapolis, MN) were added to the cultures 48 h prior to use.
When used for T cell priming, cytokine treated or untreated EC were plated at 104 cells/well in 96-well flat bottom culture plates (Costar). Purified CD8+ T cells in complete medium were added at 105 cells/well. On day 3, rhIL-2 (Boehringer Mannheim) was added to the cultures. On day 7, T cells were collected and transferred to 96-well plates containing fresh cytokine or untreated EC. T cells were stimulated as above for an additional 7 days, then harvested and tested.
EC transfection and reporter gene luciferase activity assay
The 766 bp of the 5'-flanking region of the ILT4 gene was cloned into the SacI and BglII sites, upstream of the pGL3 basic luciferase reporter gene (pGL3/766). For ILT3, 1034 bp of the 5' flanking region was cloned into pGL3 basic reporter gene (pGL3/1034). Transient transfection of human umbilical cord vein EC (HUVEC) was performed by electroporation according to the manufacturer's recommendation (HUVEC Nucleofector, Program U1, Amaxa, Germany). HUVEC were co-transfected with 2 µg of pGL3/766 or pGL3/1034 and 1 µg of pGL3 TK-renilin luciferase reporter DNA. Purified CD8+CD28 TS cells were added 24 h later and incubation was allowed to proceed for an additional 15 h. After removal of T cells, HUVEC were harvested and cell lysates subjected to luciferase assay using a Promega dual-luciferase reporter assay kit (Promega, Madison, WI) and TD 20/20 Luminometer (Turner Biosystems, Sunnyvale, CA) according to the manufacturer's instructions. Experimental luciferase reporter gene activity was normalized with renilin luciferase activity.
Semi-quantitative RTPCR
Total RNA was extracted from endomyocarditis biopsies and T cells (obtained from TCL, TCC or PBMC) using the RNAqueous4PCR kit (Ambion, Inc., Austin, TX) following the manufacturer's recommendation. First-strand cDNA was synthesized with a cDNA synthesis kit (Roche Diagnostic, Indianapolis, IN). The following primers were used in PCR reactions. ILT3: 5' primer ACGTATGCCAAGGTGAAACACT; 3' primer CATTGTGAATTGAGAGGTCTGC (expected size 493 bp). ILT4: 5' primer GCATCTTGGTGGCCGTCGTCCTAC; 3' primer CCCAAAGTTCCCAGCATCTCCTCA (expected size 551 bp). CD8: 5' primer AGGTGCTTGAGTCTCCAACGG; 3' primer TGCCTCATCCCTGTATCTGCTAGT (expected size 504 bp). CD40: 5' primer TTTCTGATACCATCTGCGAGCC; 3' primer TCTCCTGCACTGAGATGCGAC (expected size 420 bp). CD83: 5' primer ATGTCGCGCGGCCTCCAG; 3' primer TCATACCAGTTCTGTCTTGTGAGGAG (expected size 950 bp). CD54: 5' primer AAAACACTAGGCCACGCATCT; 3' primer GGCCTTTGTGTTTTGATGCTAC (expected size 253 bp). CD58: 5' primer AGATGAGCTCTTTTAACTCAAGCGAAA, 3' primer GGGTGGGAAAAAAGCATGTGTA (expected size 230 bp). CD62e: 5' primer AAGGTACACACACCTGGTTGC; 3' primer TTCTCCAGAGGACATACACTG (expected size 562 bp). CD106: 5' primer GAAGAAAAAAGCGGAGACAGGAG; 3' primer GGAGGATGCAAAATAGAGCACGAG (expected size 217 bp). FOXP3: 5' primer TTGGACAAGGACCCGATGCCCAACCCC; 3' primer CCCTGGCAGGCAAGACAGTGGAAACCTC (expected size 1350 or 1450 bp). 5' primer TGTCAGTCCACTTCACCAAGCC; 3' primer CCTTCTCATCCAGAAGATGGTCC (expected size 724 or 619). 5' primer TCCCAGAGTTCCTCCACAAC; 3' primer GCAAGACAGTGGAAACCTCAC (expected size 465 bp). Perforin: 5' primer TACAGCTTCAGCACTGACACGG; 3' primer GAGCTTCACATAGGCATCCGT (expected size 825 bp). IL10: 5' primer; TCATTCTATGTGCTGGAGATGG; 3' primer; GCTCACCATGACCCCTACC (expected size 146 bp). TGFß: 5' primer; AAGATAACCACTCTGGCGAGTCG; 3' primer, CAGAGCTCCGAGAAGCGGTAC (expected size 180 bp). GAPDH: 5' primer CGGAGTCAACGGATTTGGTCGTAT; 3' primer AGCCTTCTCCSTGGTGGTGAAGAC (expected size 362 bp). PCR reactions were done at 30 cycles in a 20 µl volume and PCR products were analyzed in an agarose gel stained with ethidium bromide. Samples were normalized with the use of GAPDH expression.
Flow cytometry analysis
Flow cytometry studies were done with a FACScan (Becton Dickinson). CaliBRITE beads from Becton Dickinson were run under the FACSComp program to calibrate the instrument. Human CD8+ T cell subsets were defined by staining with phycoerythrin (PE)-conjugated mAbs to CD28 and Cy-Chrome (CYC) conjugated mAbs to CD8. For EC staining the following mAb were used: FITCCD31, PECD40, PECD54, PECD62E, PECD83, PECD106, CYCCD58, CYCHLA ABC and CYCHLA DR (all from Becton Dickinson) and ILT3PC5 (Coulter, Miami, FL).
For ILT4 staining EC were incubated with 25 µl of rat anti-human ILT4 mAb (clone 42D1, a generous gift from Dr Marco Colonna), then washed and stained with R-PE conjugated goat anti-rat IgG (Caltag, Burlingame, CA). Rat IgG was added as a blocking agent prior to staining with FITCCD31 (Becton Dickinson). For each cell surface or intracellular marker, a corresponding isotype-matched control antibody conjugated with the same fluorescent dye was used. Six parameter analyses (forward scatter, side scatter and four fluorescence channels) were used for list mode data analysis.
Proliferation assay
EC monolayers (at 1 x 104 cells per well) were seeded in 96-well flat bottom plates (Costar) and rh IFN- (R&D Systems) was added to each well at 400 ng/ml. After 48 h, supernatant was removed and plates were washed twice with complete medium. Mitomycin C (Sigma) was added to each well at 10 µg/ml. After 2 h of incubation, plates were washed four times. Primed CD4+ T cells, obtained from each TCL using CD4 isolation kit II (Miltenyi Biotech, Auburn, CA) were added to each well at 1 x 105 cells/well. In parallel control cultures CD4+ T cells (1 x 105) were added to mitomycin C treated allogeneic monocyte-derived DC (1 x 104) in 96-well U bottom plates (Costar).
CD8+CD28 T cells from allospecific TCL were first tested for FOXP3 expression. FOXP3+ T cells were added (at 1 x 105) to cultures containing primed CD4+ T cells and either EC or DC. Monoclonal antibodies to ILT3 (clone ZM3.8, a generous gift from Dr Marco Colonna) or mixtures of mAb to ILT4 and mAb to HLA class I (W6/32) (ATCC, Manassas, VA) or rh lL-2 (10 U/ml) were added to the cultures where indicated. Triplicate cultures were set up for each condition. After 2 days of incubation, cultures were pulsed with tritiated thymidine and harvested 18 h later. [3H]thymidine incorporation was measured by scintillation spectrometry.
Statistical analysis
Fisher exact test for two-tail analysis was used for determining the significance of differences between two groups.
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Results |
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To determine whether FOXP3 is also expressed in CD8+CD28 TS we generated allospecific TCL. CD8+CD28 T cells from these TCL were separated, tested for suppressor activity and then cloned by limiting dilution. FOXP3 expression was assessed by semi-quantitative RTPCR. CD8+CD28 TS cells from TCL and TCC expressed FOXP3. No expression was detected in primed CD8+CD28+ T cells from these TCL. CD8+CD28 and CD8+CD28+ T cells from fresh peripheral blood did not express FOXP3. Unfractionated CD4+ T cells and sorted CD4+CD25+ T cells from the same samples of blood were FOXP3+ while the CD4+CD25 fraction was FOXP3 (Fig. 1A).
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To establish whether this novel FOXP3 isoform, which we refer to as FOXP3, is common to CD8+CD28 TS and CD4+CD25+ natural TR cells, we generated a primer pair which allows size discrimination of FOXP3
from FOXP3 (Fig. 1C). CD4+CD25+ T cells, but not CD4+CD25 and CD8+ T cells from fresh peripheral blood of 20 donors expressed both FOXP3 and FOXP3
. CD8+CD28 T cells from allospecific TCL generated from these donors were FOXP3 and FOXP3
positive (Fig. 1D). However, when TCC derived from one of these TCL were tested, exclusive expression of canonic FOXP3 was seen in 20 clones, while FOXP3
was expressed in 30 clones (Fig. 1D). Further characterization of TCC by RTPCR using specific primers showed that they were PRF1 negative (Fig. 1E) as well as IL-10 and TGF-ß negative (data not shown). The functional relevance of the differential expression of canonic FOXP3 and FOXP3
is currently being investigated.
EC activation is inhibited by allospecific TS cells
Serially passaged HUVEC and aortic EC (HAEC) do not display HLADR, CD80, CD86 (27,28) or detectable amounts of ILT3 or ILT4. They stimulate peripheral blood CD8+ T cells in vitro inducing the generation of alloreactive TC which express PRF1 and IFN- intracellularly (29,30). IFN-
treated EC express HLA class II as well as high levels of HLA class I and CD58 and activate both alloreactive CD4+ and CD8+ T cells from allogeneic blood donors (3133).
To study the effect of allospecific CD8+CD28 FOXP3+ TS on serially passaged human EC we selected HUVEC, which shared at least one HLA-A and HLA-B allele with the DC used for TS priming. CD8+CD28 T cells from eight different TCLs were tested by RTPCR for expression of FOXP3 and PRF1 prior to use. HUVEC were co-incubated for 18 h with TS at a 1:10 ratio and then analyzed by flow cytometry for expression of cell surface markers. The expression of costimulatory molecules CD40 and CD58 was downregulated while the inhibitory receptor ILT4 was induced. CD83 was not expressed on untreated HUVEC neither before nor after incubation with CD8+CD28 FOXP3+ TS (Fig. 2A and B).
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In previous studies we and others have shown that treatment of immature, monocyte-derived DC with IFN- and IL-10 or vitamin D3 results in the induction of a tolerogenic phenotype characterized by the high expression of ILT3 and ILT4 (23) and decreased expression of costimulatory molecules (2338). Under the same treatment conditions with IL-10 plus IFN-
, serially passaged HUVEC also expressed high cell surface levels of ILT4 (Fig. 5A) and ILT3 (Fig. 5B) as well as decreased levels of CD40 and CD58 (data not shown). Induction of ILT3 and ILT4 occurred together in IFN-
and IL-10 treated EC as previously shown to be the case for DC (23). We, therefore used the mixtures of IFN-
and IL-10 as proxies (for CD8+CD28 TS) to induce the expression of ILT3 and ILT4 on EC and determine whether ILT3+ILT4+ EC elicit the generation of CD8+CD28 FOXP3+ TS cells. In four independent experiments, allostimulation of CD8+ T cells with ILT3+ ILT4+ EC (treated with the cytokine mixture and then extensively washed) resulted in an increase in the size of the population of CD8+CD28 T cells from 30 ± 6% (Fig. 5C) before priming to 70 ± 5% after two rounds of stimulation (Fig. 5D). RTPCR studies demonstrated that these cells expressed FOXP3 (Fig. 5F) but not PRF (data not shown). In contrast, CD8+ T cells stimulated repeatedly with ILT3 ILT4 EC showed no increase in the frequency of CD8+CD28 T cells. Instead 80 ± 5% of the cells were CD8+CD28+ (Fig. 5E). T cells from these cultures were FOXP3 (Fig. 5F) yet PRF1+ by RTPCR (data not shown). Hence, ILT3+ILT4+ EC induce the differentiation of CD8+CD28FOXP3+ TS cells.
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To determine whether anti-ILT3 mAb abrogate the TS effect, mAb to ILT3 was added to cultures containing primed CD4+ TH, CD8+CD28 TS and IFN--treated EC sharing HLA class I and HLA class II antigens with the APC used for priming. Anti-ILT3 mAb had no effect on TH proliferation in cultures without TS, yet restored at least in part TH reactivity in the presence of TS (Fig. 7C). This effect was optimal at a concentration of 1 µg/well (Supplementary fig. 2).
Similarly, to establish whether ILT4 contributes to the reduced stimulatory capacity of EC exposed to TS we added mAb to ILT4 and HLA class I to cultures containing primed CD4+ TH cells, CD8+CD28 TS and IFN--treated EC. The inhibitory effect of TS on CD4+ TH proliferation in response to EC was reversed partially by the mixture of mAbs to ILT4 and HLA class I but not by either of the mAbs when used alone. This mixture of mAb had no effect on CD4+ T cell proliferation in cultures without TS (Fig. 7C). These data indicate that the effect of TS on TH proliferation in response to EC is mediated at least in part by ILT4. The combinations of mAbs to ILT3, ILT4 and HLA class I showed a synergistic effect, increasing TH proliferation in the presence of TS above the level seen when either mAb to ILT3 or mixtures of mAb to ILT4 and HLA class I were used alone (Fig. 7C and Supplementary fig. 2).
In vivo expression of ILT4 on donor EC
To understand the in vivo relevance of ILT3 and ILT4 expression on EC we studied serial endomyocardial biopsies, obtained within the first 12 months post-transplantation, from 30 heart allograft recipients. Nine of these 30 patients had at least one episode of acute rejection (histologic grade 3A or higher).
ILT4 was not expressed on EC from biopsies obtained 1 and 2 weeks after transplantation. However, ILT4 was expressed on EC from at least two consecutive biopsies obtained from 16 out of the 21 rejection-free patients (Fig. 8A). Of the nine patients with acute rejection episodes one showed ILT4+ EC on a single biopsy. Hence, the absence of acute rejection episodes during the first year post-transplantation is associated with ILT4 expression on graft EC (P < 0.0009, Fisher exact test).
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Out of 15 rejection-free patients, 12 had CD8+CD28 FOXP3+ T cells which triggered the transcription of ILT3 and ILT4 in EC sharing HLA class I antigens with the donor but not in control EC mismatched from the donor. None of the five patients with a history of acute rejection had CD8+CD28 T cells that induced ILT3 or ILT4 transcription in EC, as illustrated in Fig. 8(BD).
To determine whether direct allorecognition of EC by TS still plays a role late after transplantation when chronic rejection may occur, we next studied the presence of TS in blood samples obtained 3 years following heart transplantation from a cohort of 14 different patients. This group included four recipients with coronary artery vasculopathy (CAV) indicative of chronic rejection. Seven of the 10 rejection-free patients had CD8+CD28 FOXP3+ T cells which triggered the transcription of ILT3 and ILT4 in donor-matched EC as illustrated in Fig. 8(EG). None of the four patients with chronic rejection had circulating CD8+CD28 T cells that triggered either ILT3 or ILT4 in EC (P < 0.01).
These data suggest that the presence in the circulation of donor specific CD8+CD28 FOXP3+ T cells, which trigger ILT3 and ILT4 expression in EC, is associated with quiescence.
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Discussion |
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However, alloantigen specificity is an attribute of in vitro or in vivo generated CD8+CD28 TS which inhibit the direct T cell recognition pathway by tolerizing the APC and blocking their allostimulatory capacity (15,1720,23,24). Upregulation of inhibitory receptors ILT3 and ILT4 and down-regulation of costimulatory molecules is a critical feature of tolerogenic DC (15,1720,23,24,37,38).
Our finding that TS tolerize allogeneic APC, which in turn convert effector T cells into suppressor/regulatory T cells, fulfills the requirements of infectious tolerance and linked suppression described by Waldmann and collegues (3941).
Our model predicts that the serial transfer of tolerance to allogeneic heart transplants in Waldmann and collegues (41) was mediated by TR from the primary host which tolerized APC of the graft in secondary hosts. These tolerized APC triggered a new wave of TS and TR perpetuating tolerance through many generations of naive unmanipulated transplant recipients. Furthermore, since recognition of a single alloantigen by TS or TR is sufficient for tolerizing an APC, linked tolerance to any other antigen presented by the same APC is predicted to occur, according to our model (3941). This model does not preclude the possibility that antigen receptors on effector T cells are the targets of TR cells (42).
The effect of TS on EC is reminiscent of the changes which they induce in allogeneic DC. In the present study we have demonstrated for the first time that the interaction of allospecific CD8+CD28 FOXP3+ TS with activated EC induces the downregulation of costimulatory and adhesion molecules (such as CD40, CD54, CD58, CD62E, CD83, CD106) which mediate T cell interaction with EC and have been implicated in acute allograft rejection (33,43). Such changes are most likely caused by inhibition of NF-B activation and reduced capacity of EC to transcribe these NF-
B-dependent cell surface molecules (4450) when exposed to TS (15,20). However, TS induce a distinct differentiation pathway in non-activated EC, and reprogram activated EC converting them into tolerogenic cells. Tolerogenic EC express the inhibitory receptor ILT3 and ILT4 which act as negative regulators of T cell responses to allogeneic APC (15,21,22). The ligand of ILT3 is not known yet. However, blocking of ILT3 with anti-ILT3 mAb (ZM3.8) abrogates the TS effect.
The inhibitory receptor ILT4 has been shown to interact with a broad range of HLA class I molecules including HLA-A, B and G (21,22). Our finding that mixtures of mAbs to ILT4 and its ligand HLA class I abrogates the TS effect, demonstrates that ILT4 is involved in suppression.
Compelling evidence that donor EC play an important role in triggering TS, which inhibit the direct recognition pathway, is provided by three important findings. First, CD8+CD28 FOXP3+ T cells that trigger the upregulation of ILT4 in EC are present in the circulation of rejection-free heart allograft recipients. Second, ILT4 is expressed by EC from endomyocardial biopsies obtained from these quiescent patients. Third, the bi-directional interaction between TS and EC perpetuates long-term quiescence as demonstrated by the persistence of TS 3 years following transplantation in patients without chronic rejection.
Our results do not detract from our previous studies indicating the importance of the indirect allorecognition pathway in acute and chronic rejection of heart allografts (13). However, we demonstrated that both in the graft and in the periphery the frequency of allopeptide specific T cells was 10 000-fold lower than that of T cells involved in the direct allorecognition pathway (13,51). The demonstration that EC can elicit effectors or suppressors of the alloimmune response depending on their functional state supports the concept that direct allorecognition is a continuum throughout the lifespan of the transplant and is driven by donor EC.
These findings pertain both to acute and chronic rejection. Donor EC can elicit and become targets of alloreactive TH cells as well as anti-HLA antibodies which induce EC activation, proliferation and obstruction of transplant vasculature (52). Since chronic rejection occurs within 10 years in 40% of solid organ allografts, it is likely that regulatory mechanisms protect the graft in the remaining 60% of the recipients. Study of the capacity of CD8+CD28 FOXP3+ T cells from recipients' circulation to induce the upregulation of inhibitory receptors in EC, in an alloantigen specific manner, may permit the identification of patients who will benefit from partial or complete withdrawal of immunosuppression. This is an important aim in view of the morbidity and mortality associated with the long-term use of immunosuppressive drugs. Furthermore, the development of pharmaceutic agents that can act on DC and/or EC by upregulating inhibitory receptors, such as ILT3 and ILT4, may permit modulation of the immune response in patients with autoimmune diseases or transplants. The recent finding that in vitro-generated tolerogenic APC induce CD8+ T regulatory cells which can suppress ongoing experimental autoimmune encephalomyelitis (53) supports the rationale for developing such new therapeutic strategies.
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Supplementary data |
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Acknowledgements |
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Abbreviations |
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DC | dendritic cells |
EC | endothelial cells |
ILT | immunoglobulin-like transcript |
HAEC | human aortic endothelial cells |
HUVEC | human umbilical vein endothelial cells |
PBMC | peripheral blood mononuclear cells |
PRF | perforin |
Tc | cytotoxic T cells |
TCC | T cell clones |
TCL | T cell lines |
TR | T regulatory cells |
TS | T suppressor cells |
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Notes |
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Received 25 March 2004, accepted 30 April 2004.
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References |
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