A Novel Flow Assay for the Detection of Cytokine Secreting Alloreactive T Cells: Application to Immune Monitoring

A Novel Flow Assay for the Detection of Cytokine Secreting Alloreactive T Cells: Application to Immune Monitoring Yael D. Korin, Clara Lee, David W. Gjertson, Alan H. Wilkinson, Thu-Phoung Pham, Gabriel M. Danovitch, H. Albin Gritsch, and Elaine F. Reed ABSTRACT: The direct and indirect allorecognition pathways play an important role in graft rejection. We hypothesized that the presence of alloreactive memory T cells in the recipient’s circulation increases the risk of rejection after transplantation. The objective of this study was to develop a noninvasive, immune monitoring tool that simultaneously measures donor-specific responses via both the direct and indirect recognition pathways. Our laboratory developed a whole blood flow cytometric cytokine secretion assay to identify interferon (IFN)-␥ secreting memory T cells in whole blood of renal transplant patients. The assay readily detected IFN-␥ producing CD3⫹ T cells in response to recall antigens tetanus toxoid, purified protein derivative, and alloantigens in whole blood from healthy controls. Analysis of sequential posttransplant blood samples from 19 renal allograft reABBREVIATIONS Abs antibodies APC antigen-presenting cell AR acute rejection FCCS flow cytometry cytokine secretion LDA limiting dilution analysis

INTRODUCTION Despite improvements in immunosuppression and patient management, renal allograft rejection still remains a major obstacle to long-term transplant survival. Acute rejection occurs in up to 20% of transplants and can be From the UCLA Immunogenetics Center, Departments of Pathology and Laboratory Medicine (Y.D.K., C.L., D.W.G., E.F.R.), Urology (H.A.G.), and Division of Nephrology (A.H.W., T.-P.P.), David Geffen School of Medicine, University of California, Los Angeles, CA. Address reprint requests to: Elaine F. Reed, Ph.D., Department of Pathology and Laboratory Medicine, David Geffen School of Medicine at UCLA, 1000 Veteran Avenue, Los Angeles, CA 90095. E-mail: [email protected]. Received August 20, 2005; accepted October 13, 2005. Human Immunology 66, 1110 –1124 (2005) © American Society for Histocompatibility and Immunogenetics, 2006 Published by Elsevier Inc.

cipients showed that alloimmune responses were higher in transplant recipients who had undergone acute rejection than in those without acute rejection episodes. In addition, patients showing increased creatinine levels 3 months after transplantation were more likely to exhibit alloimmune responses than recipients with stable graft function. The flow cytokine secretion assay provides a reliable and simple method for identification of patients at risk of acute rejection and early graft dysfunction. Human Immunology 66, 1110 –1124 (2005). © American Society for Histocompatibility and Immunogenetics, 2006. Published by Elsevier Inc. KEYWORDS: Alloreactivity; immune monitoring; IFN-␥; cytokine secretion assay

MLR PPD SEB TT

mixed lymphocytes reaction purified protein derivative staphylococcal enterotoxin B tetanus toxoid

mediated by cellular or humoral immune responses directed against alloantigens on the graft. The current approach to diagnose renal allograft rejection is monitoring changes in serum creatinine and examination of graft histology. Although monitoring the creatinine levels is a sensitive marker for graft rejection and dysfunction, it is at best a late one, because graft rejection must have already preceded deterioration in renal function. Furthermore, although examination of graft histology is critically important to diagnose rejection, it is not routinely used as a monitoring tactic because the biopsy procedure is invasive and carries risk of complications. 0198-8859/05/$–see front matter doi:10.1016/j.humimm.2005.10.010

Cytokine Secreting Alloreactive T Cells

Identification of early surrogate markers of graft rejection and graft dysfunction that have the potential to diagnose rejection before the advent of irreversible renal injury could provide the clinician with an opportunity for early intervention and to monitor effectiveness of immunotherapies. A recipient’s T cells that recognize foreign donor major histocompatibility complex (MHC) antigens displayed on the graft mediate allograft rejection. Recognition occurs by two distinct pathways: direct allorecognition of intact donor MHC molecules expressed on the membrane of graft cells and indirect allorecognition of donor MHC peptides bound to the MHC molecules on host antigen-presenting cells (APCs) [1]. Although T cells activated through the direct allorecognition play a primary role in the process of acute rejection during the early posttransplant period, they are unlikely to contribute to later rejection events because passenger donor dendritic cells migrate out of the graft early after transplantation [2–5]. Several reports implicate the indirect recognition pathway as the major contributor to late allograft rejection [6 –10]. Thus, T cells recognizing allopeptides associated with host restriction elements were detected in the graft and in the periphery of transplant recipients undergoing chronic rejection of kidney, lung, liver, and heart allografts [9, 11–15]. Reliable in vitro assays that permit the quantitation and characterization of the alloimmune response hold promise for identifying patients at risk of acute and chronic rejection, optimizing drug regimens, and evaluating the effectiveness of new immunotherapies. Several methods to determine the frequency of alloreactive T cells have been reported (for review, see [16, 17]). Serial posttransplant monitoring studies by limiting dilution analysis (LDA) showed that T-cell proliferation to donor allopeptides was strongly associated with episodes of acute and chronic allograft rejection [2, 5, 7, 14]. Although LDA has become a standard approach for estimating T-cell frequencies, the test is limited by the long culture time of 7–10 days that may result in clonal expansion [18]. Newer approaches to determine T-cell frequencies are based on the detection of a cytokine produced by single cells after antigen stimulation. The enzyme linked immunospot (ELISPOT) assay can identify antigen-specific memory T cells producing cytokines in short-term assays [19 –21]. Several studies have shown that estimation of frequencies of alloantigen specific T cells by ELISPOT correlates with graft outcome [19, 22]. More recently, flow cytometry based methods for the detection of cytokine producing alloreactive T cells have emerged [18, 23]. Although the ELISPOT technique detects single cytokine-producing cells with great sensitivity and specificity, the assay does not identify the phenotype of the

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cytokine secreting cells present in the population of peripheral blood lymphocytes. The use of the intracellular cytokine staining technique is attractive because it identifies the phenotype of the cytokine-producing cell; however, this assay is based on the assumption that intracellular production of cytokines always results in its secretion. Furthermore, the cells are rendered nonviable during the staining procedure, preventing subsequent studies. The aim of this study was to develop a simple and quantitative measurement of direct and indirect allorecognition in the peripheral blood of the transplant recipient. We designed a cytokine secretion flow cytometry– based assay for the simultaneous evaluation of direct and indirect allorecognition directly in whole blood. We report that this assay detects cytokine production by antigen specific T memory cells at the single cell level, providing an indication of their in vivo function. Investigation of the direct and indirect pathways in sequential samples of blood obtained from 19 renal transplant recipients shows that the assay can distinguish patients with higher risk of graft rejection and early graft dysfunction from those with stable function. MATERIALS AND METHODS Study Population Fifteen healthy volunteers were asked to participate based on their human leukocyte antigen (HLA) typing and previous history of tetanus toxoid (TT) and purified protein derivative (PPD) immunizations, and were used as controls for assay development and calibration. The study population consisted of 19 adult deceased donor renal allograft recipients (14 males, 5 females) transplanted between May 2003 and March 2004. Patients were enrolled in this institutional review board– approved immune monitoring study when there was an increased risk of renal allograft rejection due to anticipated delayed graft function, prior exposure to transplantation antigens, or poor HLA matching. Sequential samples of blood were scheduled to be collected pretransplantation and at 1, 2, 3, 4, 6, 8,12, 24,36, 52, 62, 80, 92, and 104 weeks after transplantation. The average number of time points studied per patient was seven and most samples (80%) were collected during the first year after transplantation. Clinical data were obtained by chart review and from the clinical database, including patient demographics, donor demographics, and posttransplant outcome variables and creatinine levels. All recipients had negative pretransplant T- and B-lymphocyte cytotoxicity and flow cytometry crossmatches with the donor. All patients received either Thymoglobulin or Simulect induction. Three of the 19 patients were treated with Thymoglobulin and 16 of 19

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received Simulect induction therapy. Maintenance therapy consisted of either tacrolimus or cyclosporine, mycophenolate mofetil, and prednisone. All patients received an initial prednisolone bolus of 1000 mg and tapering doses to 20 mg of prednisone over 7 days. The prednisone dose was reduced to 5 mg per day over the first 3 months. Immunosuppression was adjusted for episodes of rejection, side effects, or infections. Cell Isolation Sodium heparinized peripheral blood was obtained from healthy volunteers and adult renal allograft recipients. Peripheral blood mononuclear cells (PBMC) were isolated by Ficoll-Hypaque gradient centrifugation. Singlecell suspensions of spleen cells in RPMI medium were obtained by mechanical disruption of the tissue and mononuclear cells were isolated from this cell suspension by Ficoll-Hypaque gradient centrifugation. Stimulator and donor APCs were purified by depleting cell suspensions of T cells using the Rosette Sep (StemCell Technologies, Vancouver, Canada) as recommended by the manufacturer. T-cell depletion of spleen cell suspensions was performed by negative selection using anti-CD3 antibodies (Abs) (Miltenyi Biotec, Auburn, CA), and MACS® magnetic columns according to the manufacturers directions (Miltenyi Biotec, Auburn, CA). Antigens and Peptides Staphylococcal enterotoxin B (SEB) was obtained from Sigma Chemical Co. (St. Louis, MO) and was used at a final concentration of 1 ␮g/ml. Purified protein derivative (PPD; Mycos Research, Loveland, CO), was used at a final concentration of 10 ␮g/ml, and TT (Calbiochem, San Diego, CA), was used at 1 ␮g/ml. Peptides (20- to 25-mers) corresponding to the hypervariable regions of HLA-A*0201 residues 60-84 (WDGETRKVKAHSQTHRVDLGTLRGY) and DRB1*0401 RESIDUES 69-88 (EQKRAAVDTYCRHNYGVGES), were synthesized by Biopeptide Co. (San Diego, CA) and were used at a concentration of 1–10 ␮g/ml. Membrane-Bound Cell-Free Donor Antigen Preparations Cytoplasmic membrane protein preparations from stimulator and donor PBMC were obtained as previously described [15]. A total of 5–10 ⫻ 107 PBMCs or spleen cells were lysed by three cycles of freezing and thawing in a Tris-EDTA-based buffer containing 1/5000 Nonidet P-40 0.1 mM PMSF, 1/200 protease inhibitor mixture, and 5 ng/ml soy-bean trypsin inhibitor (Sigma Chemical Co.), centrifuged at 1000g for 2 minutes, and the supernatant was collected and further centrifuged for 45 minutes at 14,000g. The pellet was collected and resuspended in assay medium and tested for protein concentration. The equivalent of 1 ⫻ 106 intact cells was

Y. Korin et al.

added per each 1 ⫻ 106responder cells. The absence of whole intact cells was confirmed by microscopy. The presence of HLA class I and class II molecules in the pellet was confirmed using the GTI solid phase ELISA kit according to the manufacturer’s recommendations (GTI, Inc., Waukesha, WI). Generation of Short-Term T-Cell Line Alloreactive T cells were generated in mixed lymphocyte reaction (MLR) by stimulating 106/ml responder PBMCs with 106/ml irradiated HLA mismatched stimulator APCs in RPMI 1640 supplemented with 10% pooled human Ab serum, 2 mM L-glutamine, 100 ug/ml each penicillin and streptomycin. After 10 days, the MLR culture was restimulated via the direct recognition pathway using irradiated donor APCs (106) or via the indirect recognition pathway, using membrane-bound cell-free donor antigens (4 ␮g/ml, the equivalent of 106 cells/ml) or synthetic peptides (1–10 ␮g/ml) corresponding to donor-mismatched HLA-A*0201 and DRB1*0401 antigens in the presence of 106 autologous APCs. Shortterm alloreactive T-cell lines were produced by restimulation and expansion in recombinant interleukin (IL)-2 (RDI Inc., Flanders, NJ). Fourteen days after priming, T cells were restimulated at 106/ml with an equal number of irradiated stimulator APC or 4 ␮g/ml (the equivalent of 106 cells/ml) membrane-bound cell-free donor antigens, in the presence of 106 autologous APCs, and the cells were expanded with 10 ng/ml IL-2 at weekly intervals for an additional 2 weeks. Whole Blood MLR The whole blood MLR was performed as described by Pauly et al. [24] and Bromelow et al. [25] with the following modifications: freshly drawn sodium heparinized blood (0.5 ml) was stimulated with 1–2 ⫻ 106irradiated stimulator APCs or T-cell– depleted spleen cells, resuspended in 0.5 ml 10% human Abs RPMI, supplemented with 2 mM L-glutamine, 100 ␮g/ml each penicillin and streptomycin. Cells were cultured for 15 h, at 37 °C in a 5% CO2 incubator in 48-well plates, and cytokine production was measured using cell surface affinity matrix technology (Miltenyi Biotech, CA), as specified below. Flow Cytometry Cytokine Secretion (ECCS) Assay Freshly drawn sodium heparinized whole blood (0.5 ml) or 0.5–1 ⫻ 106 PBMCs were cultured at 37 °C in a 5% CO2 incubator in 48-well plates in the presence or absence of irradiated stimulator APCs (2 ⫻ 106), or 4 ␮g (the equivalent of 1 ⫻ 106 cells) membrane-bound cellfree donor antigens, 1 ␮g/ml SEB, 10 ␮g/ml PPD, or 1 ␮g/ml TT. Anti-CD28 (0.1 ␮g/ml) was added to cultures stimulated with PPD, TT, and cell-free donor

Cytokine Secreting Alloreactive T Cells

antigens. After a 15-h incubation, cytokine production was measured using cell surface affinity matrix technology (Miltenyi Biotech), according to the manufacturer’s instructions with some modifications and stained with a phycoerythrin-labeled interferon (IFN)-␥ detection antibody (Myltenyi Biotech) and FITC-conjugated anti-CD3 mAb. Exclusion of dead cells was accomplished by staining cells with 7-amino-actinomycin D (Sigma Aldrich). For whole blood, red blood cells were lysed using 1.55 M NH4CL, 100 mM KHCO3,1 mM EDTA (adjusted pH to 7.3) for 20 minutes at room temperature in the dark, centrifuged for 5 minutes at 1500 rpm and the cells were fixed with 0.8% paraformaldehyde containing 0.5 ␮g/ml Actinomycin D (Sigma Aldrich). Cell fluorescence was analyzed on a FACSCalibur flow cytometer (BD Biosciences, San Jose, CA) and analyzed with CellQuest Pro software (BD Biosciences). The percent of cytokine producing CD3⫹ T cells in control wells was subtracted from the total percent of CD3⫹ responder T cells. The flow cytometric cytokine secretion (FCCS) method permits the detection of as few as 0.01% of the gated positive cells. Because frequencies of antigen-specific cytokine-producing cells are low, a minimum of 1 ⫻ 105 live PBMCs were acquired and up to 20,000 CD3⫹ cells were collected. FCCS assays with less than 1000 detectable CD3⫹ T cells were excluded from the study. The cutoff value for a positive CD3⫹ T cell IFN-␥ response was calculated based on a 95% confidence interval of the negative and positive controls and was determined to be 0.07%, equivalent to a frequency of 14 IFN-␥⫹ cells, adjusted per 20,000 CD3 T cells. Spiking Experiments The accuracy and sensitivity of the FCCS method were assessed in “spiking” experiments via recovery tests. Serial dilutions of known numbers of SEB-stimulated, IFN-␥⫹ cells were added to non-stimulated cells and the numbers of recovered IFN-␥⫹ cells were measured. The correlation coefficient between the measured and the expected numbers of IFN-␥⫹ cells was calculated. HLA Typing All study subjects were HLA typed using DNA-based typing methods. Low-resolution typing of HLA class I and class II antigens was accomplished using polymerase chain reaction and hybridization with sequence-specific oligonucleotide probes using luminex reagents purchased from One Lambda (One Lambda Inc., Canoga Park, CA). Allele level HLA-A and HLA-B locus typing was performed by sequenced-based typing using reagents (Atria Genetics, San Francisco, CA). DRB1 and DQB1 typing was performed using sequenced based typing as we have previously described [26].

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Statistical Analysis Demographic characteristics in recipients with and without acute rejection episodes were compared using Fisher’s exact test for discrete variables (reported as counts) and Wilcoxon’s rank sum test for continuous variables (shown as means ⫾ standard deviations). Pearson’s squared correlation coefficient and simple linear regression analysis were used to assess the accuracy of the FCCS assay to estimate numbers of IFN-␥⫹ cells in standard solutions of IFN-␥⫹ cell concentrates. Concerning outcome data, contingency tables were analyzed for differences in proportions of all response-positive recipients with and without rejection and with and without stable graft function using Fisher’s exact test. Statistical significance was defined as a p value ⬍ 0.025 via Bonferroni’s procedure, allowing for two comparisons of main variables. All p values were two-sided, and all estimates were done via the STATA statistical software package (StataCorp 2003 Stata Statistical Software, release 8; College Station, TX).

RESULTS Detection of cytokine-secreting alloreactive memory T cells using the FCCS assay. We tested short-term allogeneic T-cell lines in recall FCCS assays to determine the feasibility of detecting alloreactive cytokine secreting T cells. Peripheral blood lymphocytes (PBLs) from healthy donor A (HLAA*3303, A*3402, B*3501, B*5801, DRB1*1501, DRB1*1102, DRB3*02, DRB5*01) were stimulated with irradiated PBLs from healthy donor B (HLAA*0201, A*0301, B*1501, B*3501, DRB1*0401, DRB1*1101, DRB3*02, DRB4*01) for 10 days followed by restimulation with purified irradiated stimulator donor B’s APCs or membrane-bound cell free donor antigens preparations derived from donor B in the presence of autologous APCs. In addition, cultures were rechallenged with a mixture of two allopeptides corresponding to the first domain of the mismatched stimulator DRB1*0401 and A*0201 molecules in the presence of autologous APCs. As shown in Figure 1, rechallenge of A␣B CD3⫹T cells with APCs from donor B elicited IFN-␥ production in 3.8% of CD3⫹ T cells (Panel B), compared with 0.03% in cells cultured in medium alone (Figure 1A). Only a small percentage of A␣B T cells produced IFN-␥ (0.02%) when stimulated with third-party cells from a donor that did not share HLA antigens with stimulator B (Figure 1C). In four independent experiments using different responder/stimulator combinations, IFN-␥ producing alloreactive memory T cells were readily detectable (range 1– 4.5%) after primary rechallenge of MLR cultures with donor

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Y. Korin et al.

A

B

Unstimulated

C

Allostimulation

0.03%

0.02%

IFN-γ

IFN-γ

3.8%

IFN-γ

101

100

102

104

103

CD3

D

101

100

102

104

103

E

0.14%

IFN-γ

IFN-γ 100

10 4

10 3

101

CD3

G

102

H

104

102

103

10 2

104

CD3

10 3

10 4

CD3

I

Stimulation with cell free alloantigens and restimulaiton with allo-APCs 0.77%

IFN-γ

IFN-γ 101

10 1

3.83%

IFN-γ 100

10 0

Expansion and restimulation with allopeptides

5.49%

J

103

CD3

Expansion and restimulation with cell free alloantigens

10 4

Expansion with allo-APC and restimulation with cell free allopeptides

F

0.16%

IFN-γ

10 2

10 3

10 2

CD3

Expansion with allo-APC and restimulation with cell free alloantigens

26%

10 1

10 1

10 0

CD3

Expansion and restimulation with allo-APC

10 0

3P-allostimulation

10 0

10 1

10 2

CD3

10 3

10 4

100

101

102 FL1 H

103

104

CD3

Stimulation with cell free alloantigens and restimulation with 3P cell free alloantigens

IFN-γ

0.15%

100

101

102

103

104

CD3

FIGURE 1 Interferon (IFN)-␥ secretion by in vitro directly and indirectly primed T cells. Short-term alloreactive T cells (A␣B), were produced in vitro by mixed lymphocytes reaction (MLR) and assayed for IFN-␥ production after restimulation on day 10 via the direct recognition pathway. (A) Unstimulated; (B) irradiated antigen-presenting cells (APCs) of donor B; (C) irradiated third-party mismatched APCs. (D) Cells were further expanded in interleukin-2 and restimulated twice with irradiated APCs from donor B and assayed for IFN-␥ production. Short-term alloreactive T cells (A␣B), were produced in vitro by MLR and assayed for IFN-␥ production after restimulation on day 10 via the indirect recognition pathway. (E) Autologous APCs and cell-free antigens from donor B; (F) autologous APCs and synthetic HLA-A*0201 and DRB1*0401 peptides. Cells were further expanded in interleukin-2 and restimulated twice and assayed for IFN-␥ production after restimulation with: (G) autologous APCs and donor cell–free antigens; (H) autologous APCs and synthetic HLA-A*0201 and DRB1*0401 peptides; (I) irradiated APCs of donor B; (J) autologous APCs and third-party cell-free alloantigens. Percent of CD3⫹ IFN-␥⫹ T cells is shown in the upper right quadrant of each panel, respectively.

Cytokine Secreting Alloreactive T Cells

APCs. Subsequent restimulation (two times) of the primary A␣B culture with irradiated APCs from stimulator B resulted in the enrichment of alloreactive CD3⫹ IFN-␥ producing T cells responding via the direct allorecognition pathway (Figure 1D). The frequency of T cells producing IFN-␥ was 7-fold higher (26%) than the primary MLR cultures, consistent with the enrichment of alloreactive memory T cells. The process of repeated stimulation of A␣B T cells did not contribute to expansion of memory T cells responding via the indirect allorecognition pathway as rechallenge with stimulatorderived cell-free donor antigens or synthetic peptides corresponding to mismatched donor HLA-A*0201 and DRB*0401 molecules resulted in only 0.16% (Figure 1E) and 0.14% (Figure 1F) of IFN-␥ producing CD3⫹ T cells, respectively. These results demonstrate the ability to detect T cells activated through the direct pathway using the FCCS assay. We next determined whether we could identify selfrestricted alloreactive T cells activated via the indirect allorecognition pathway. The A␣B T cells were restimulated on Day 10 with membrane-bound cell-free donor antigens in the presence of autologous APCs and subsequently expanded in IL-2 for 2 additional weeks. Cells were then tested for reactivity against either irradiated donor’s APCs (direct pathway) or via the indirect allorecognition pathway using cell-free donor antigens or synthetic peptides corresponding to mismatched donor HLA-A*0201 and DRB1*0401 molecules. A␣B CD3⫹ T cells primed through the indirect pathway produced IFN-␥ in response to stimulation with cell-free donor antigens (5.49%, Figure 1G) or allopeptides (3.83%, Figure 1H). In contrast, cells treated with donor cells or third-party cell free donor antigens failed to show significant IFN-␥ responsiveness (Figures 1I and 1J, respectively). Analysis of results from two additional independent experiments, using different healthy responder/ stimulator combinations, showed that self-restricted alloreactive T cells were detectable after rechallenge of MLR culture with cell-free donor antigens (range 1.5– 3.8%) or peptides (range 0.8 –2.5%) corresponding to mismatched stimulator HLA class I and class II antigens. These data show that the FCCS assay can detect alloreactive T-cell memory responses initiated via the indirect recognition pathway. Furthermore, these findings demonstrate that the FCCS assay accurately detects IFN-␥ production only by antigen specific primed T memory cells. Detection of Cytokine-Producing T Memory Cells in Whole Blood Using the FCCS Assay Because our goal was to develop an assay to detect antigen primed T cells in the peripheral blood of transplant recipients, we enumerated cytokine-producing T

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memory cells in whole blood obtained from healthy donors immunized to tetanus toxoid or PPD. Blood samples from healthy donors were stimulated with 1 ␮g/ml TT or 10 ␮g/ml PPD and 0.1 ␮g/ml anti-CD28 mAb and IFN-␥ secretion was measured 15 h later using the FCCS assay. As a positive control, blood was stimulated in parallel with 1 ␮g/ml SEB. Table 1 summarizes the percentage and frequency of IFN-␥⫹CD3⫹ antigenspecific T cells in six healthy controls. Stimulation with SEB resulted in CD3⫹ T-cell activation and production of IFN-␥ in all individuals tested (range 2.91–7.17%, frequency 582–1434 IFN-␥⫹, 20,000 CD3⫹ T lymphocytes). Whole blood incubated in medium alone showed minimal production of IFN-␥ (range 0.00 – 0.02%, frequency 0 – 4 IFN-␥⫹E20,000 CD3⫹ T lymphocytes). CD3⫹ T cells from individuals with a history of vaccination to Bacillus Calmette-Guérin produced a response to PPD (range 0.45– 4.51%, frequency 90 –916 IFN␥⫹20,000 CD3⫹ T lymphocytes). Similarly, CD3⫹ T cells from individuals with a recent history of immunization to TT (past 3–10 years) produced in response to stimulation with TT (range 0.37– 0.48%, frequency 74 –96 IFN-␥⫹20,000 CD3⫹ T lymphocytes). Figure 2 illustrates data from two healthy individuals. The upper panels represent data from an individual that has a history of vaccination to TT but not PPD. Analysis of memory T cells in the peripheral blood showed that 0.56% and 0.06% of CD3⫹ T cells secreted IFN-␥ in response to TT and PPD, respectively. The lower panels represent data from an individual that has a history of vaccination to Bacillus Calmette-Guérin but has not been vaccinated against TT during the past 10 years. A vigorous IFN-␥ response (1.59%) was observed when blood cells were challenged with PPD, but there was little spontaneous production of IFN-␥ in unstimulated blood and no significant response to TT (0.03%). These results indicate the ability of the FCCS assay to enumerate antigen specific cytokine producing memory T cells directly in whole blood. We next investigated the feasibility of the FCCS assay to enumerate alloreactive T cells in whole blood. As presented in Table 2, IFN-␥ producing alloreactive CD3⫹ memory T cells were observed in the whole blood of 13 of 34 (38%) healthy donors after a 15-h incubation with irradiated stimulator APCs. The percentage of IFN-␥ producing memory CD3⫹ T cells ranged from 0.12– 0.78%, equivalent to 24 –156 IFN-␥ producing cells/20,000 total CD3⫹. These results demonstrate the ability of the FCCS assay to measure functional T-cell alloimmune responses directly in whole blood without purification of the responding T-cell populations.

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TABLE 1 Detection of antigen-specific T memory cells in the whole blood of healthy individuals Donor A

B

C

D

E

F

Recall Ag

%IFN-␥ ⫹ CD3 ⫹ cellsa

# IFN-␥ ⫹ CD3 ⫹ cellsb

Frequency IFN-␥ ⫹ CD3⫹c

None SEB PPD TT None SEB PPD TT None SEB PPD TT None SEB PPD TT None SEB PPD TT None SEB PPD TT

0.01 5.39 0.45 0.03 0.00 2.91 0.05 0.37 0.00 3.22 0.06 0.45 0.02 7.17 4.51 0.04 0 3.69 0.06 0.48 0.02 4.69 3.22 0.01

3 1078 90 6 0 582 11 74 0 665 13 90 4 1434 916 7 0 738 12 96 4 938 644 2

1:79,050 1:146 1:1972 1:17,243 NA 1:120 1:7048 1:750 NA 1:153 1:7521 1:1185 1:10,152 1:25 1:91 1:8682 NA 1:153 1:8500 1:804 1:30,000 1:90 1:118 1:87,000

Description

PPD skin test positive No TT immunization within 3–10 years PPD skin test negative Immunized for TT within 3–10 years PPD skin test negative Immunized for TT within 3–10 years PPD skin test positive No TT immunization within 3–10 years PPD skin test negative Immunized for TT within 3–10 years PPD skin test positive No TT immunization within 3–10 years

Abbreviations: Ag ⫽ antigen; IFN ⫽ interferon; PPD ⫽ purified protein derivative; SEB ⫽ staphylococal enterotoxin B; TT ⫽ tetanus toxoid. a Percentage of IFN-␥ ⫹ cells of the total CD3⫹ T lymphocytes. b Number of IFN-␥ ⫹ cells per 20,000 CD3⫹ T lymphocytes. c Frequency of IFN-␥ ⫹ cells per total acquired PBMCs.

Assay Reproducibility and Sensitivity The precision of the FCCS assay was established by determining the intra- and interassay variability, accuracy, and sensitivity. To determine intra-assay variability, fresh whole blood was obtained from five healthy donors and tested in triplicate for reactivity to SEB, TT, and PPD. As shown in Figure 3A, the intra-assay variability is low with a coefficient of variation of 5–15%. To determine the interassay variability, healthy individuals were tested on different days over a 6-month period, for reactivity to SEB (Figure 3B), TT, PPD, or MHC disparate irradiated APCs (Figure 3C). As shown in Figures 3B and 3C, interassay variability was acceptable with a coefficient of variation (CV) of 12–32%. Note that the CV is higher when the frequencies of antigen-specific T cells are low (Figure 3C). Furthermore, the interassay CV is influenced not only by variability in sample handling on the different days, but also biologic changes that might have occurred over the 6-month period during which the individuals were tested. To determine the accuracy and sensitivity of the FCCS assay, we tested cytokine production in “spiking” experiments. SEB-stimulated T cells were added at increasing concentrations to a constant number of lymphocytes.

Thus, the numbers of IFN-␥-producing T cells in each assay tube was known and the frequency of expected IFN-␥⫹ cells was determined and plotted against the recovered number of IFN-␥⫹ cells. Figure 4 represents two independent “spiking” experiments. The number of IFN-␥ producing T cell titrates linearly against the number of IFN-␥ producing cells added in increasing concentration to 3 ⫻ 105 PBMCs or to 1 ⫻ 10 6 PBMCs in experiment A and B, respectively. Moreover, almost 100% of IFN-␥⫹cells were detected by the FCCS technology, producing an almost perfect correlation between measured and expected data. Experiment B also demonstrates that as few as 50 IFN-␥⫹ cells can be detected out of 1 ⫻ 105 CD3⫹ gated cells, and a total of 5 ⫻ 105 acquired cells. The lower limit of detection of the FCCS is therefore 0.01%, or 1/10,000 cells. Study of Alloreactive T-Cell Memory Responses in Renal Transplant Recipients To assess the frequency of donor specific alloreactive T cells in the circulation of transplant recipients we tested serial post transplant blood samples from 19 recipients of deceased donor renal allografts. A summary of the patient demographics is presented in Table 3. Nine of the

Cytokine Secreting Alloreactive T Cells

A Unstimulated

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C TT stimulation

B SEB stimulation

3.83%

103

102

101

CD3

102

103

104

104

104

101

102

103

104

100

CD3

CD3

101

102

103

104

CD3

H PPD stimulation 1.59%

IFN-γ 100

100

0.03%

IFN-γ 103

103

G TT stimulation

5.11%

IFN-γ

102

102

101

CD3

F SEB stimulation

0.02%

101

100

CD3

E Unstimulated

100

IFN-γ

IFN-γ 100

104

IFN-γ

101

100

0.06%

0.56%

IFN-γ

IFN-γ

0.01%

D PPD stimulation

at e ts/8/06/03 00

101

102

103

104

100

CD3

101

102

103

104

CD3

FIGURE 2 Detection of antigen-specific interferon (IFN)-␥⫹ CD3⫹ memory cells in whole blood using the flow cytometric cytokine secretion assay. Whole blood from two healthy donors were unstimulated (A, E), or stimulated overnight with Staphylococcal enterotoxin B (B, F), 1 ␮g/ml TT (C, G), or 1 ␮g/ml PPD (D, H). Percent of CD3⫹IFN-␥⫹ cells are shown in the upper right quadrant of each panel.

19 patients were diagnosed with acute rejection, eight with acute cellular rejection and 1 patient was diagnosed with concurrent acute cellular rejection and acute humoral rejection. Six patients with acute rejection (AR) were male and three were female. Four of the 19 patients included in the study were African American (3 in the

AR group and 1 in the non-AR group). The average age of the 9 AR patients was 52.5 years and that of the non-AR patients was 58.8 years. Two of the 9 (22%) patients with AR and 1 of the 10 (10%) patients without AR were recipients of secondary renal allografts. Analysis of patients characteristics showed no association between

TABLE 2 Detection of alloreactive IFN-␥-producing CD3⫹ T memory cells in whole blood Responder-stimulator combinationa 1 2 3 4 5 6 7 8 9 10 11 12 13 Mean, SD Range

%IFN-␥⫹ CD3⫹ cells unstimulatedb

%IFN-␥⫹ CD3⫹ cells Stimulated over nightb

# IFN-␥⫹ CD3⫹ cells Stimulated over nightc

Frequency IFN-␥⫹ CD3⫹d

0.03 0.01 0 0 0 0.02 0.02 0 0.02 0.02 0 0 0.01 0.01 ⫾ 0.01 0–0.03

0.35 0.77 0.39 0.74 0.25 0.17 0.45 0.18 0.78 0.13 0.62 0.12 0.17 0.39 ⫾ 0.25 0.12–0.78

70 154 78 148 50 34 90 36 156 26 124 24 34 79 ⫾ 66 24–156

1:995 1:613 1:1,800 1:850 1:1,445 1:2,450 1:1,770 1:11,000 1:3,700 1:5,400 1:954 1:4,800 1:4,700

See Table 1 for abbreviations. a Data representative of subset of controls positive for alloreactivity. b Percentage of IFN-␥ ⫹ cells of the total CD3⫹ T lymphocytes. c Frequency. d Frequency of IFN-␥ ⫹ cells per total acquired peripheral blood mononuclear cells.

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1200

A # of IFN-γ+CD3+ T cells

1000

600 400 200 0

Average 795 s.d. 44 c.v. 5.5%

% IFN-γ+CD3+ T cells

B

800

794 53 6.6% SEB

957 50 5.2%

193 29 15% TT

133 11 7.6% PPD

7 6 5 4 3 2 1 0

C

% IFN-γ+CD3+ T cells

Average s.d. c.v.

4.5 1.4 31%

3.6 0.5 14%

4.6 0.9 20%

0.9 0.8 0.7 0.6 0.5 0.4 0.3 0.2 0.1 0

Average s.d. c.v.

0.53 0.07 13.4%

0.14 0.04 28.57% TT

0.46 0.10 21.73% PPD

0.47 0.07 14.99%

0.40 0.60 0.20 0.05 12.48% 32.86% Allo

FIGURE 3 Inter- and intra-assay reproducibility. (A) Whole blood obtained from five healthy individuals was tested for reactivity to staphylococcal enterotoxin B (SEB), tetanus toxoid (TT), and purified protein derivative (PPD) in triplicate wells; (B) whole blood obtained from three healthy individuals on 5–7 different days during a 6-month period tested for reactivity to SEB; (C) whole blood obtained from six healthy controls obtained on 3–7 different days over a 6 month-period and tested for reactivity to TT, PPD, or allogeneic major histocompatibly complex mismatched antigen-presenting cells. Each data point represents the percent of interferon-␥ secreting CD3⫹ lymphocytes detected in a single secretion assay. Average, standard deviation (SD) and coefficient of variation (CV) are listed for each data set.

diagnosis of AR and recipient age, gender, race, number of transplants, and number of HLA mismatches between recipient and donor (Table 3). Patient samples were tested in FCCS assays in response to donor APCs (direct pathway), donor cell-free membranes (indirect pathway), and third-party alloantigens. The average number of time points tested per patient was seven. We were unable to measure alloresponses using the FCCS assay in 3 time points in 3 patients treated with Thymoglobulin and in 4 time points in 16 patients treated with Simulect because of an insufficient number of CD3⫹ T cells. Analysis of serial

samples of posttransplant blood showed that 12 of 19 (63%) patients showed increased frequencies of donor alloreactive T cells after transplantation. Ten (52%) patients displayed responses to donor cells (direct pathway), one patient (5%) responded to donor cell-free donor antigens (indirect pathway), and one patient (5%) displayed alloreactivity via both the direct and indirect pathways (Table 4). Analysis of the relationship between the development of posttransplant T-cell alloreactivity and graft outcome indicated that 8 of the 12 patients (67%) demonstrating increased frequencies of IFN-␥ producing T cells in response to donor alloantigens experienced AR compared with only 1 of 7 patients (14%) in the nonalloresponsive group (Figure 5A, p ⫽ 0.06). The mean frequency of IFN-␥ producing donor reactive CD3⫹ T cells was higher in patients with AR compared with patients without AR (Figure 5B, p⫽ 0.04). In contrast, most recipients responded to third-party stimulator cells (no significant difference among groups; Figure 5B). We further categorized the patients into groups based on mean serum creatinine levels measured after the third posttransplant month as a surrogate marker of early graft dysfunction and analyzed the relationship with donor reactive IFN-␥ CD3⫹ T cells. We found that all eight patients with early graft dysfunction (creatinine ⱖ1.5 mg/dl) showed donor alloreactivity. In contrast, only 4 of 11 (36%) patients with stable function (creatinine ⱕ1.5 mg/dL) demonstrated donor alloreactivity (Figure 5C, p⫽ 0.01). Although not statistically significant, there was a positive trend toward association between incidence of acute rejection and early graft dysfunction. Thus, 6 (75%) of 8 patients who had poor graft function experienced an episode of AR, whereas only 3 (27.3%) of 11 patients with stable graft function had an episode of AR (Figure 5D, p⫽ 0.07). Together, these data indicate that development of donor-specific T-cell alloreactivity is found more often in recipients with graft rejection and early graft dysfunction. Serial monitoring studies permitted us to examine the temporal relationship between donor-specific alloimmune responses and onset of AR. One of the nine patients who experienced AR failed to show donor alloreactivity and was not included in this analysis. Six of the eight remaining patients (67%) with acute rejection exhibited donor alloreactivity before diagnosis, whereas two patients showed donor alloreactivity following the episode of AR. Four patients experienced AR during the first 3 posttransplant months (early AR) and four patients were diagnosed with AR after the third posttransplant month (late AR). Increased T-cell alloreactivity was observed in two of four patients before the onset of early AR and in four of four patients with late AR (Figure 5D and Table 4).

Cytokine Secreting Alloreactive T Cells

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Experiment A

Experiment B

1400

600

R2

= 0.9999

1200 1000 800 600 400 200 0 0

200

400

600

800

1000

1200

1400

1600

Number expected IFN- γ+ CD3 cells

Number recovered IFN−γ+CD3 cells

Number recovered IFN−γ+CD3 cells

1600

500

R2 = 0.9868

400 300 200 100 0 0

100

200

300

400

Number expected IFN-

500

γ+

600

700

800

CD3 cells

FIGURE 4 Accuracy of the flow cytometric cytokine secretion assay (recovery experiments). Staphylococcal enterotoxin B stimulated interferon (IFN)-␥ secreting PBMCs were added in increasing numbers (in spiking experiments) to a constant number of unstimulated PBMCs (experiment A ⫽ 3 ⫻ 105; experiment B ⫽ 1 ⫻ 106 cells). The expected numbers of IFN-␥⫹ cells are plotted on the x axis against the recovered numbers of IFN-␥⫹cells on the y axis.

DISCUSSION Data presented in this study document the feasibility of detecting antigen primed IFN-␥–producing T cells in the peripheral blood of healthy donors and immunosuppressed transplant recipients in a short-term recall assay. The whole blood FCCS assay accurately detected antigenspecific and alloreactive cytokine-secreting T cells at the single-cell level at frequencies as low as 0.01% of the specific gated population of cells. The sensitivity of the TABLE 3 Patients characteristics Age, years (mean ⫾ SD) 21–30 31–40 41–50 51–60 61–70 71–80 Gender (males/females) Race (AA/non-AA) Retransplants (primary/regrafts) Type of donor CAD HLA-ABDR mismatches (mean ⫾ SD) 0 mm 1–2 mm 2–3 mm 3–4 mm 5–6 mm

Non AR

AR

p valuea

n ⫽ 10 0 2 2 3 2 1 8/2 1/10 9/1

n⫽9 1 0 3 1 2 2 6/3 3/9 7/2

0.67

10 4.7 ⫾ 1.6

9 4.8 ⫾ 0.4

NA 0.45b

0 1 1 1 7

0 0 0 2 7

0.63 0.3 0.58

Abbreviations: AR ⫽ acute rejection; CAD ⫽ cadaveric; HLA ⫽ human leukocyte antigen. a Fisher’s exact test. b Via Wilkinson Ranksum.

FCCS assay increases depending on the total number of acquired cell counts and can reach a sensitivity of 1/100,000 total PBMCs, or 1/20,000 CD3⫹ T cells, when 100,000 or more cells are acquired. Antigenspecific memory responses to TT and PPD were in the range of 0.13–3.22% CD3⫹ T cells and were consistent with the individual’s history of immunization. Thus, the sensitivity and specificity of the FCCS assay was comparable to that reported for other assays, including intracellular cytokine staining and approaching the sensitivity of ELISPOT [18, 21, 22], with the added benefit that the FCCS assay can be performed directly in whole blood, avoiding cell manipulation, and the phenotype of the cytokine producing cells can be readily identified as well [27]. The whole blood assay also has the advantage that it preserves the in vivo milieu, thereby measuring the transplant recipient’s immune response in the context of immunosuppressive drug therapy. Consistent with previous publications [20], we found directly primed alloreactive T cells in the fresh blood of healthy control donors. The frequencies of IFN-␥– producing alloreactive memory T cells present in responders ranged from 0.01% to 0.78% or a frequency of 1/600 –1/40,000 (Table 2) and similar to reports by others using ELISPOT [21]. Interestingly, the frequency of alloreactive T cells was similar to the frequencies of recall responses in healthy individuals immunized with TT and Bacillus Calmette-Guérin (Table 1). This marked increase in the frequency of alloreactive cells in nonallosensitized individuals is consistent with our previous observation that the frequency of T cells engaged in the direct recognition pathway is 100-fold higher than the frequency of T cells responding to nominal antigens

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TABLE 4 Posttransplant alloimmune responses in renal allograft recipients Patient 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

Tx no.

Donor MM A, B, DR

Average serum Cr after 3 mo

Rejection type

Alloimmune response typea

2 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 2 1 2

A31, B60, B51, DR8, DR11, DQ4 A2, A31, B60, B51, DR8 DR11, DQ4, DQ7 -, -, DR2 A2, B62, B42, DR4, DQ8 A2, A24, B7, B44, DR15, DQ1 A24, A26, B35, B45, DR7, DQ2 A2, B7, B61, DR11, DR15, DQ6, DQ7 A26, B61, DR11 A24, A26, B35, B45, DR7 A3, A11, B49, DR1, DR4 A30, B37, B44, DR4, DR15, DQ6, DQ8 A11, B35, B49, DR4, DQ8 A26, B38, B48, DR4, DR14, DQ8 A2, A30, B48, B50, DR8, DQ3, DQ4 A30, B48, B50, DR4, DR8, DQ3, DQ4 A2, A3, B7, B37, DR1, DR13 A29, B13, B14, DR7 A11, B35, B75, DR9, DQ3, DQ4 A2, B60, DR12, DQ7

1.2 1.35 1.65 2.03 1.12 2.5 1.2 1.2 1.06 1.5 2.6 1.2 2 1.8 1.8 1 1.5 1.5 3

None None None ACR ACR ACR None None ACR None ACR None ACR None None None ACR, AHR ACR ACR, AHR

Negative Direct Direct Indirect (post AR) Negative Direct & Indirect (pre AR) Negative Negative Direct (pre AR) Negative Direct (pre AR) Negative Direct (pre AR) Negative Direct Direct Direct (pre AR) Direct (post AR) Direct (pre AR)

Abbreviations: Tx ⫽ transplantation; MM ⫽ number of human leukocyte antigen mismatches, A, B, DR; ACR ⫽ acute cellular rejection; AHR ⫽ acute humoral rejection. a Cutoff value ⫽ 0.07% over background.

[3] and compatible with the hypothesis that alloreactivity reflects cross-reactivity to the universe of peptide: MHC complexes derived from environmental antigens. We used the FCCS assay to prospectively monitor 19 renal allograft recipients for the development of antidonor alloreactivity via the direct and indirect allorecognition pathways. This study is unique in that we performed a simultaneous kinetic evaluation of both the indirect and direct recognition pathways in the whole blood of renal transplant recipients. Our data indicate that development of Th1-type donor-specific alloreactivity identifies a subpopulation of patients at increased risk of acute rejection and graft failure. Thus, the mean frequencies of IFN-␥ producing donor-reactive T cells were higher in patients with AR than in patients without evidence of rejection. The incidence of early graft dysfunction as reflected in mean serum creatinine values ⬎1.5 after the third posttransplant month was also higher in patients who produced responses to donor alloantigens compared to patients without evidence of donor alloreactivity, supporting a role for Th1-type cytokine producing alloreactive T cells in the process of acute and chronic rejection. Since most recipients responded equally to third party alloantigens, the data suggest that immune reactivity to donor alloantigens is a characteristic distinctive to the group of patients with rejection and early graft dysfunction. Alternatively, the absence of donor reactivity in patients with stable graft function may reflect the development of hyporesponsive-

ness to donor alloantigens [16, 28]. These data are in accord with other clinical studies in renal, heart, lung and liver allografts that linked donor hyporesponsiveness with better clinical outcome [16]. Although the mechanisms underlying hyporesponsiveness to organ allografts are not completely understood, clonal deletion and T-cell anergy mediated by donor-specific suppressor/regulatory T cells are suggested models. Based on studies by SuciuFoca et al. [29 –31] and preliminary data by our group, we favor the latter mechanism whereby tolerized endothelial cells of the graft play a pivotal role in inducing the generation of hyporesponsive T cells. Both in vitro and in vivo studies show that CD8⫹CD28⫺FoxP3⫹ T suppressor cells (Ts), which are specific for donor class I antigens, can induce the downregulation of costimulatory molecules and upregulation of inhibitory molecules ILT3 and ILT4 on endothelial cells of the donor [31–36]. These endothelial cells become tolerogenic, inducing Th anergy and stimulating alloantigen specific CD4⫹ CD25⫹ T regulatory cells. Both recipients of kidney and liver allografts displaying T suppressor and T regulatory cells have reduced incidence of rejection and require lower immunosuppression [33, 37]. In this study, we observed that transplant recipients display lower frequencies of alloreactive T cells when compared with healthy control responders. There are two plausible explanations for this observation. First, testing of healthy individuals was performed on whole blood in the absence of immunosuppressive drugs, whereas post-

Cytokine Secreting Alloreactive T Cells

1121

Alloresponses versus acute rejection AR +

8

p = 0.06

Number of patients

7

AR -

Number of patients

A

6 5 4 3 2 1 0

Allo+

Acute rejection versus graft funciton

D

9

Allo-

9 8

Poor GF

Stable GF

7

p = 0.07

6 5 4 3 2 1 0

AR-

Frequency IFN-γ/20,000 CD3+ cells 16

DM

p = 0.04

14

TM

Allo responses in early versus late acute rejections

12 10 8 6

E

4 2 0

AR+

AR-

Number of patients

B

Number IFN-γ/20,000 CD3+ cells

AR+

8 7 6 5 4 3

6/8

Pre AR Post AR

4/4

2/4

2 1 0

C

Number of patients

Alloresponses versus graft function 9

p = 0.01

8

Poor GF

Early AR

Lat e AR

t ot al AR

Stable GF

7 6 5 4 3 2 1 0

Allo+

Allo-

FIGURE 5 Patients with acute rejection display higher frequencies of alloreactive T cells to donor alloantigens. (A) Number of patients displaying alloimmune responses via the direct and indirect pathways in patients with and without acute rejection. (B) Mean frequencies of interferon (IFN)-␥⫹/20,000 CD3⫹ cells after stimulation with donor or third-party alloantigens. The numbers for each group represent mean frequencies of IFN-␥ secreting CD3⫹ T cells in response to donor-specific allostimulation (DM) versus stimulation with third-party (TM) in patients with (AR⫹) or without (AR⫺) acute rejection. (C) Number of transplant recipients within each group who exhibited a positive antidonor flow cytometric cytokine secretion assay. Patients were grouped according to stable function (mean creatinine after third posttransplant month ⬍1.5 mg/L) or poor function (mean creatinine after third posttransplant month ⬎1.5 mg/L). (D) Correlation between acute rejection and graft function. Number of transplant recipients within each group who exhibited a positive antidonor response. Patients grouped according to stable function (mean creatinine after third posttransplant month ⬍1.5 mg/L) or poor function (mean creatinine after third posttransplant month ⬎1.5 mg/L). (E) The kinetics of alloimmune responses in early versus late acute rejection. Patients were grouped according to the time we have observed alloimmune responses in relation to the onset of the acute rejection. Early AR occurs during the first 3 months posttransplant; late AR occurs after the third posttransplant month. p values ⬍ 0.05 are considered significant.

transplant patient testing was performed on whole blood in the presence of immunosuppressive agents. Second, the patients’ alloresponses to the donor are lower because of the development of donor-specific hyporesponsiveness. However, comparison of the frequencies of IFN-␥ secreting T cells stimulated via the direct activation pathway to third-party cells showed lower frequencies in patients than controls suggesting that hyporesponsiveness was not a likely explanation for the differences observed. We conclude that the decreased alloreactivity in patients is most likely due to immunosuppressive drugs. We found that transplant recipients displayed T-cell reactivity to donor alloantigens via both the direct and indirect recognition pathways. IFN-␥ responses to donor cells via the direct allorecognition pathway were found in

57% of the patients, whereas only 10% of the patients responded to membrane-bound cell free donor antigen preparations via the indirect recognition pathway. The finding that rejection is not always accompanied by both direct and indirect allorecognition can be explained by the different requirements for stimulation of these pathways. Shedding of graft antigens is probably the main stimulus for the indirect pathway, whereas directly activated T cells are stimulated by graft APCs. Our finding that only 10% of patients showed donor-specific indirect recognition in samples primarily collected during the first posttransplant year suggests that the alloresponse during the first year after transplantation is indeed biased toward development of Th1-type direct allorecognition as previously suggested [10, 28, 38, 39]. Our data dif-

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fers, however, from previous reports that showed higher frequencies of T cells responding to donor allopeptides during the first year after transplantation [14, 40]. This disparity may be explained by the different test methods employed (i.e., LDA analysis, ELISPOT, proliferation, flow cytometry), the alloantigen preparations used, or evolution of the immune response during the course of transplantation. For example, Suciu-Foca and colleagues expanded the pool of indirectly primed T cells by first culturing T cells in IL-2 before allopeptide stimulation and LDA analysis. They observed activation of the indirect recognition pathway as early as 1 week after transplantation and showed a significant correlation between allopeptide reactivity in the periphery and incidence of acute and chronic rejection [5, 14, 40]. Their reported frequency range of in vivo activated T cells specific for donor peptides using the LDA assay was reported as 1.5–10 cells/million, which is more sensitive than the FCCS assay. It is also possible that the source of alloantigens (i.e., peptides versus cell-free donor antigens) may differ in their abilities to interact with APCs and trigger T-cell activation. Although several studies have documented the use of allopeptides as potent stimuli of indirect recognition, studies by Lechler and colleagues have clearly shown that cell-free membrane antigen preparations can be used to pulse APCs in lieu of allopeptides [2, 41] because only CD4⫹ T cells previously primed through the indirect pathway respond in measurable numbers. Our results are consistent with these findings and showed that indirectly primed T cells were not reactivated when exposed to donor cells but rather only produced IFN-␥ in response to stimulation with cell-free donor antigens or allopeptides. From a practical standpoint, we reason that T-cell priming with cell-free donor antigens maximizes the likelihood of detecting alloresponses and is a more cost-effective way to stimulate T cells than allopeptides because all of the donor alloantigens (class I and class II) are present in the single preparation and available for antigen processing and presentation by host APCs. Another possible explanation for our observation of low frequencies of indirectly primed T cells is the timing of these studies. Only three previous groups have attempted to measure alloresponses via the direct and indirect pathways in the same group of patients [2, 40, 42]. The synopsis of these studies is that the direct pathway is predominant during the early posttransplant period but diminishes with time, whereas raised frequencies of indirectly primed T cells correlated with chronic rejection. Because our study primarily focused on samples collected during the first year after transplantation, the probability of detection of indirectly activated T cells should increase as more time elapses. Further studies are underway to confirm this possibility. Examination of the temporal relationship between

Y. Korin et al.

development of donor-specific alloimmune responses (both direct and indirect) and rejection showed that the majority of patients (67%) with acute rejection displayed donor reactivity in the peripheral blood before diagnosis of rejection. Because alloreactive T cells were detectable in the circulation of most patients before the diagnosis of rejection, our data suggest that monitoring of the indirect and direct pathways may be valuable for the early prediction of rejection. The use of immunosuppressive agents such as Thymoglobulin and Campath pose a potential limitation to performing cell-based immune monitoring assays because T-lymphocyte counts in the peripheral blood of treated patients may be very low during the early period after transplantation. In this study, three patients were treated with Thymoglobulin and 16 were treated with Simulect. Analysis of the impact of Thymoglobulin on the performance of the FCCS assay showed that all three patients treated with Thymoglobulin had one time point early after treatment where there were insufficient cells to analyze the FCCS data (i.e., less than 1000 CD3⫹ T cells in the gate). In contrast, only a single time point from 4 of 16 patients treated with Simulect could not be studied because of an insufficient number of cells. These results indicate that treatment with T-cell depleting immunosuppressive agents is a potential obstacle to performing the FCCS assay. In conclusion, this study provides a clinically useful approach to analyze and expand our understanding of the role of direct and indirect recognition in allograft rejection. T-cell alloreactivity in the periphery can be rapidly measured by the FCCS assay in 24 h, making this a feasible, noninvasive approach for identifying patients at risk of rejection, guiding immunosuppressive drug therapy, and monitoring response to therapy. This technique can also be applied to the study of the immune response to nominal proteins and peptides with direct application to vaccine research and clinical trials. ACKNOWLEDGMENTS This work was supported by National Institute of Allergy and Infectious Diseases Grant RO1 AI 42819 (to E.F.R). We thank Dr. J Michael Cecka for his constructive insights and editorial expertise in completion of this manuscript.

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28.

29.

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