Allograft rejection remains a major cause of morbidity and mortality after lung transplantation and is associated with increased gene expression in human lung alografts (1). T cells are a major cell type involved in transplant rejection and are the target of current immunosuppression strategies (2). We have recently shown that CD4+ T-cell proinflammatory cytokines were significantly reduced in peripheral blood and bronchoalveolar lavage (BAL) from stable lung transplant patients consistent with adequate immunosuppression (3, 4). Importantly, we also showed that inhibition of proinflammatory cytokines by CD8+T cells in these patients was less effective (3, 4). Systemic drug levels were within “therapeutic range,” suggesting that intracellular T-cell cytokine levels from blood may give a better indication of immunosuppression than drug levels.
Analysis of cells and cytokines in BAL has also previously been used as an indicator of transplant rejection and may be more relevant for assessing lung transplantation (5–7). Although changes in CD4:CD8 in BAL have been associated with transplant rejection (5, 6), surface phenotyping gives little information regarding cell function and in particular cytokine production by various leukocyte subsets. Analysis of soluble inflammatory cytokines in BAL and blood may not be as reliable as analysis of intracellular cytokines using flow cytometry (8); this may be attributed to the presence of soluble cytokine receptors (9). Our previous reports of differential inflammatory cytokine production in blood, BAL and intraepithelial T cells from bronchial brushings (BB) from stable lung transplant patients (3, 4, 10) suggests that the same level of immunosuppression may not occur in all of the various lung/blood compartments. There have been no previous studies of intracellular cytokines in T cells from blood, BAL, and intraepithelial T cells from bronchial brushings (BB) during lung transplant rejection.
We hypothesized that T-cell proinflammatory cytokines would be increased in the airways of lung transplant patients during acute rejection episodes. To investigate this hypothesis, whole blood, BAL, and BB from stable lung transplant patients and lung transplant patients with evidence of biopsy-proven rejection were stimulated in vitro and intracellular cytokine production by CD8+ and CD4+ T-cell subsets were determined using multiparameter flow cytometry.
MATERIALS AND METHODS
Patient and Control Groups
Ten lung transplant recipients with histopathological evidence of current acute rejection were invited to participate in the study and fully informed consent was obtained following institutional ethics approval. Demographic details of these patients are shown in Table 1. Twenty-five lung transplant recipients with no clinical or histopathological evidence of current acute or chronic rejection, scheduled for routine surveillance assessment were also invited to participate in the study. Twelve of these patients were assessed on several occasions. All patients were submitted to the same protocol and analysis performed retrospectively. All patients were tested to exclude cytomegalovirus (CMV; histopathologically, rapid viral culture and CMV polymerase chain reaction [PCR] of BAL), mycoplasma (enzyme immunoassay of BAL), bacterial and fungal infection (BAL culture). All transplant patients were at least 2 months posttransplant. Acute rejection was assessed by experienced pathologists and graded according to International Society of Heart and Lung Transplantation guidelines (A≥1, B=0) (11, 12) and were 12±6.8 months posttransplant. All patients enrolled in the stable control group were classified as A0B0 and were 10±7.2 months posttransplant.
Immunosuppression therapy comprised combinations of either cyclosporin A (CsA) or tacrolimus (Tac) with prednisolone, and azathioprine or mycophenolate mofetil. Trough plasma drug levels of either CsA or Tac were within or above recommended therapeutic ranges (range for CsA: 80–250 μg/L; Tac: 5–20 μg/L). Predisposing pathology and other patient demographics are shown in Table 2. Venous blood was collected into 10 U/mL preservative free sodium heparin (DBL, Sydney, Australia) and blood, BAL, and BB samples were maintained at 4°C until processing.
Full blood counts, including white cell differential counts, were determined on blood specimens using a CELL-DYN 4000 (Abbot Diagnostics, Sydney, Australia). Blood films were stained by the May-Grunwald-Giemsa method and white cell differential counts checked by morphological assessment microscopically. BAL cell counts were determined using a hemocytometer using standard techniques.
Aspirated BAL samples (3×50 mL aliquots) were collected as previously described (4) and transferred to 50-mL polypropylene tubes. For each collection from an individual, the first aliquot was processed for microbiological and viral testing. BAL specimens 2 and 3 were pooled and cell counts determined as described above and cells were then pelleted by centrifugation at 500×g for 5 min. Supernatant was discarded and cells resuspended at 4×105 cells/ml in Roswell Park Memorial Institute (RPMI) 1640 media supplemented with 125 U/mL penicillin and 125 U/mL streptomycin (Gibco, New York).
Fiberoptic bronchoscopy was performed as previously described (10). Cells were obtained from the third- or fourth-order bronchi with several passages of the brush into each airway so as to avoid bleeding. Cells were deposited by washing the brush in 5 mL RPMI in 10 mL conical polypropylene tubes (Johns Professional Products, Sydney) and kept on ice until processed.
Absolute Lymphocyte and CD4+ and CD8+ T-Cell Counts
One hundred microliters of peripheral blood, BAL, and BB were stained with appropriately diluted fluorescently conjugated monoclonal antibodies to CD8 fluorescein isothiocyanate (FITC; BD Biosciences [BD], Sydney, Australia), CD4 PE (BD) and CD3 PC5 (Beckman Coulter, Sydney, Australia) as previously described (3, 4, 10). Samples were analyzed by gating using forward scatter (FSC) versus side scatter (SSC) to exclude platelets and debris. Gated cells were analyzed with CD45/CD14 (BD) to ascertain that cells were of lymphoid origin as previously reported (3). Control staining of leukocytes with antimouse immunoglobulin (Ig) G1-FITC (BD)/IgG1a-PE (BD)/IgG1-PC5 (Beckman Coulter) was performed on each sample and background readings of <2% were obtained. A minimum of 5,000 (from blood cultures) and 3,000 CD3 positive, low SSC events from BAL and BB samples were acquired in list-mode format for analysis.
T-Cell Cytokine Production
T cell cytokine production was assessed as previously described (3, 4, 10). Two-milliliter aliquots of prepared BAL, BB, or 1-mL aliquots of blood (diluted 1:2 with RPMI 1640 medium) were placed in a 10-mL sterile conical PVC tubes (Johns Professional Products, Sydney, Australia). Phorbol myristate (25 ng/mL; Sigma, Sydney, Australia) and ionomycin (1 μg/mL; Sigma) were added to stimulate T-cell cytokine production. Brefeldin A (10 μg/mL) was added as a “Golgi block” (Sigma) and the tubes reincubated in a humidified 5% CO2/95% air atmosphere at 37°C. At 16 hours, 100 μL of 20 mM ethylenediamine tetraacetic acid/phosphate-buffered saline was added to the culture tubes which were vortexed vigorously for 20 sec to remove adherent cells. To lyse red blood cells in the blood cultures, 2 mL of FACSlyse solution (BD) was added and tubes incubated for 10 min at room temperature in the dark. After centrifugation at 500×g for 5 min and decanting, 0.5 ml 1:10 diluted FACSperm (BD) was added to each blood, BAL and BB culture tube, mixed, and incubated a further 10 min at room temperature in the dark. Two milliliters of 0.5% bovine serum albumin (Sigma) in Isoton II (Beckman Coulter) was then added and the tubes centrifuged at 300×g for 5 min. After decanting supernatant, Fc receptors were blocked with 10 μL of human immunoglobulin (Intragam, CSL, Parkville, Australia) for 10 min at room temperature. Five microliters of appropriately diluted anti-CD8 (BD) and anti-CD3 PC5 (Coulter/Immunotech) phycoerythrin (PE)-conjugated anticytokine monoclonal antibodies to IL2, IL4, interferon (IFN)-γ, tumor necrosis factor (TNF)-α (BD) and transforming growth factor (TGF)-β (IQ Products, Groningen, Nederlands) or isotype control monoclonal antibody was added for 15 min in the dark at room temperature. Two mls of 0.5% bovine serum albumin in Isoton II was then added and the tubes centrifuged at 300×g for 5 min. After decanting, cells were analyzed within 1 h on a FACSCalibur flow cytometer using CellQuest software (BD). Samples were analyzed by live gating using FL3 staining versus side scatter (SSC). A minimum of 5,000 (from blood cultures) and 3,000 CD3 positive, low SSC events from BAL and BB samples were acquired in list-mode format for analysis. Control staining of cells with antimouse IgG1-PE/IgG-PC5 was performed on each sample and background readings of <2% were obtained.
Data are reported as means±SD. Statistical analysis was performed using the nonparametric Mann-Whitney test using SPSS software. Differences between groups with P<0.05 were considered significant.
Absolute Blood CD4+ and CD8+ T-Cell Counts
There was no significant difference between the absolute blood leukocyte counts of stable and rejecting patient groups (7.8±4.3 and 7.9±4.6×109/L, for stable patients and patients with acute allograft rejection respectively, P>0.05). There was no significant difference in the absolute lymphocyte counts for patient groups (1.5±0.7 and 1.5±0.6×109/L for stable patients and patients with acute allograft rejection respectively, P>0.05).
There was no significant difference in the absolute T cell count for patient groups (1.1±0.8 and 1.3±0.6×109/L, for stable patients and patients with acute allograft rejection respectively, P>0.05).
Absolute BAL CD4+ and CD8+ T-Cell and Leukocyte Counts
There was no significant difference between the absolute BAL leukocyte count for patient groups (0.60±0.60 and 0.62±0.59×109/L, for stable patients and patients with acute allograft rejection, respectively, P=0.396 and 0.328). There was no significant difference in the absolute T cell counts for patient groups (0.02±0.03 and 0.02±0.02×109/L, for stable patients and patients with acute allograft rejection respectively, P>0.05). There was no significant increase in the absolute macrophage counts for patient groups (0.490±0.112 and 0.530±0.114×109/L, for stable patients and patients with acute allograft rejection respectively, P<0.05). There was no significant increase in the absolute neutrophil counts for patient groups (0.089±0.098 and 0.071±0.068×109/L, for stable patients and patients with acute allograft rejection respectively, P<0.05).
Absolute BB CD4+ and CD8+ T-Cell Counts
There was no significant difference in the percentage of T cells in BB for patient groups (2.39±1.75 and 3.30±2.37%, for stable patients and patients with acute allograft rejection respectively, P>0.05); no significant difference in the absolute T cell counts in BB for patient groups (0.021±0.018 and 0.029±0.021×109/L, for stable patients and patients with acute allograft rejection respectively, P>0.05); and no significant difference in the total T cell yield in BB for patient groups (0.105±0.086 and 0.145±0.098×106 cells, for stable patients and patients with acute allograft rejection respectively, P>0.05).
T-Cell Cytokine Production From Blood From Transplant Patients
There was no significant difference in the percentages of CD8+ or CD8− (CD4+) T cells in the blood of stable patients and patients with acute allograft rejection (Table 3). The percentage of CD4−CD8− and CD4+CD8+ T cells was <3% for both patient groups.
There was no significant difference between the percentage of CD8+ or CD8− (CD4+) T cells producing IFNγ, IL-2, IL-4, or TNFα in the blood of transplant patient groups (P>0.05; data in Table 3). There was a significant decrease in the percentage of CD8+ and CD8− (CD4+) T cells producing TGFβ in patients with rejection compared with stable transplant patients (Table 3). Representative dot plots showing the percentage of blood CD8+ and CD8− (CD4+) T cells producing TGFβ in a stable patient and a patient with acute allograft rejection are shown in Figure 1.
T-Cell Cytokine Production From BAL From Transplant Patients
There was no significant difference in the percentages of CD8+ or CD8− (CD4+) T cells in the BAL of stable patients and patients with acute allograft rejection (58±14 and 56±14% for CD8+ and 43±13 and 46±14% for CD4+, for stable patients and patients with acute allograft rejection respectively, P<0.05). The percentage of CD4−CD8− and CD4+CD8+ T cells was <3% for both patient groups.
There was no significant difference between the percentage of CD8+ or CD8− (CD4+) T cells producing IL-2 or IL-4 in the BAL of transplant patient groups (P>0.05; Fig. 2). There was a significant increase in the percentage of both CD8+ and CD8− (CD4+) T cells producing IFNγ and TNFα in patients with acute allograft rejection compared with stable patients. Representative dot plots showing the percentage of BAL CD8+ and CD8− (CD4+) T cells producing IFNγ and TNFα in a stable patient and a patient with acute allograft rejection are shown in Figure 3. T cell yields were insufficient to quantify TGFβ production by T-cell subsets in BAL samples.
Intraepithelial T-Cell Cytokine Production From BB From Transplant Patients
There was no significant difference in the percentages of CD4+ or CD8+ intraepithelial T cells in stable patients and patients with acute allograft rejection (Table 4). The percentage of CD4-CD8− and CD4+CD8+ T cells was <3% for both patient groups.
There was no significant difference between the percentage of CD8+ or CD8− (CD4+) intraepithelial T cells producing IFNγ, IL-2, IL-4, or TNFα in the BB of transplant patient groups (P>0.05; Table 4). There was, however, a trend in the percentage of total intraepithelial T cells producing IFNγ and TNFα in patients with allograft rejection compared with the stable transplant group. T-cell yields were insufficient to quantify TGFβ production by T-cell subsets in BB samples.
This is the first comprehensive report of intracellular Th1/Th2/Th3 pro- and anti-inflammatory cytokines in blood, BAL and BB (intraepithelial) T cells from lung transplant patients undergoing acute rejection episodes. We show that acute rejection episodes are associated with decreased Th3 cytokine production, TGFβ by T cells in blood and increased Th1 pro-inflammatory cytokines, and IFNγ and TNFα in airway-derived T cells compared with stable transplant patients. Peripheral blood CD4+ T-regulatory cells (Tregs) have been shown to be important in maintaining graft tolerance (13) and are potent negative regulators of Th1 proinflammatory cytokines (14). We now show that both CD8− (CD4+) and CD8+ T-cell subsets producing the T-regulatory cytokine TGFβ are decreased in the peripheral blood of patients undergoing acute rejection compared with stable lung transplant patients. Unfortunately, T-cell yields were insufficient in BAL and BB specimens to quantify TGFβ producing T-cell subsets in these compartments and this would be an important adjunct to this present study. One could speculate that as the Th1 proinflammatory cytokines IFNγ and TNFα were increased in airway-derived T cells in patients with allograft rejection, Treg numbers may be reduced; our ongoing studies are investigating this possibility. In contrast to our findings, a previous study showed a significant expansion of CD4+CD25++ CD69-peripheral blood Tregs during acute lung rejection episodes (15). However, surface phenotyping gives little information regarding cell function and in particular cytokine production by cells. Our findings of increased Th1 proinflammatory cytokines in airway-derived T cells are consistent with previous studies that showed increased gene expression for proinflammatory cytokines IL-2, IFNγ, and TNFα in human allograft biopsies (1) during acute rejection episodes. In this regard, we showed increased IFNγ and TNFα (but not IL-2) by CD4+ and CD8 + T cell subsets in BAL and a trend in intraepithelial T cells in BB. Our findings of unchanged IL-2 production by intraepithelial and BAL T cells are in conflict with the reported increased gene expression of this cytokine in lung biopsies (1) and may reflect differential expression of IL-2 within the subepithelium. Consistent with our findings, one previous study showed increased levels of TNFα mRNA in rejecting lung allografts but not increased TNFα serum levels (16) and another reported a correlation between increased BAL IFNγ (but not IL-2) mRNA (17) during acute rejection episodes.
Our current results together with the previous report of the synergistic action of IFNγ and TNFα in activating both large and small airway epithelial cells (18) may have important implications for lung allograft rejection.
Taken together, these findings suggest that current immunosuppression protocols are effective at reducing Th1 proinflammatory cytokine levels in the blood but are inadequate at reducing levels in the airways during rejection episodes. The use of flow cytometry to quantify intracellular Th1/Th2/Th3 levels of inflammatory cytokines in T-cell subsets in the various compartments during rejection episodes allows precise identification of specific cell subsets producing these cytokines and is quicker and less invasive than techniques involving mRNA PCR and lung biopsy.
Our results suggest that use of drugs that effectively reduce airway T-cell IFNγ and TNFα may improve current protocols for reducing acute graft rejection in lung transplant patients. The utility of surveillance bronchoscopy in lung transplantation is controversial. Our results may give support for monitoring immunosuppression levels by measurement of intracellular inflammatory cytokine levels in T-cell subsets in blood, BAL, and BB rather than measurement of trough serum levels because these drug levels were within therapeutic range. Aerosolized cyclosporine A treatment has been used successfully and safely in reducing inflammatory cytokines in refractory acute rejection (19). This therapy may be of benefit in treating lung transplant patients identified with high percentages of inflammatory cytokine producing T cells in BAL and BB by targeting the lungs while minimizing systemic side effects of immunosuppressive agents (20).
However, one must be vigilant in not overimmunosuppressing patients; a Th1 proinflammatory response is important in host defense against bacterial infection in the lungs. We have previously shown that a group of lung transplant patients with significantly decreased levels of Th1 proinflammatory cytokines were infected with pathogenic microorganisms (21). The degree to which transplant recipients are immunosuppressed influences their risk of infection and rejection (22). The restoration of Th1 responses has been shown to be an important predictor of fungal infection outcome in stem cell transplantation patients (23).
Monitoring the balance of intracellular Th1 cytokines between levels associated with infection and rejection may improve morbidity in our patient group. Sequential monitoring of T-lymphocyte gene expression has previously been used to predict early postrenal allograft rejection (24). We are currently undertaking a study of longitudinal profiles of intracellular T-cell subset inflammatory cytokines in the various compartments in lung transplant patients to alert clinicians to changes in inflammatory cytokines associated with possible infection and rejecting episodes.
In conclusion, acute lung transplant rejection is associated with decreased intracellular T-cell TGFβ in blood and increased intracellular IFNγ and TNFα in BAL CD4+ and CD8+ T cells. Drugs that effectively reduce airway T-cell IFNγ and TNFα proinflammatory cytokine production may improve current protocols for reducing acute graft rejection in lung transplant patients. The clinical relevance of this work is being further pursued with longitudinal follow-up of this group and a much larger patient cohort.
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