CMV-Specific CD8+ T-Cell Dynamics in the Blood and the Lung Allograft Reflect Viral Reactivation Following Lung Transplantation

Authors


Abstract

Despite the potentially high burden of cytomegalovirus (CMV)-related disease following lung transplantation, the role of the cytotoxic T-lymphocyte (CTL) response to CMV in this patient group is ill-defined. We assessed the CMV-specific T-cell response in the blood and lung allograft of immunosuppressed lung transplant recipients receiving antiviral prophylaxis and following their withdrawal. While the proportion of CMV-specific CTL varied between patients, in the absence of CMV reactivation the level of CMV-specific CD8+ T cells in the blood remained stable over time. In the majority of patients CMV-specific cells could be detected in the lung allograft, often in the absence of viral DNA. Additionally, following primary CMV lung infection, CMV-specific CD8+ T cells were detected no earlier than 100 days post-transplantation but still prior to the detection of viral DNA in the lung allograft. Together these findings suggest that very low levels of CMV replication are sufficient to both prime and recruit CMV-specific CD8+ T cells to the MHC-mismatched lung allograft. The direct detection of CMV-specific T cells with an effector phenotype in the lung allograft suggests a protective antiviral function. This study provides a framework upon which the association between CMV and chronic allograft rejection can be further studied.

Introduction

More than 20 years after the first successful lung transplant cytomegalovirus (CMV) continues to negatively influence the post-operative course of transplant recipients. During this time, the focus of the impact of CMV has shifted from direct effects, such as CMV pneumonitis, to an appreciation of indirect effects, which include chronic allograft rejection (1,2), currently the major obstacle to long-term improvement in outcomes following lung transplantation (3).

Clinically, broad spectrum non-specific immunosuppression regimens are routinely used to limit graft rejection by alloreactive T cells. However, this may result in impaired antiviral T-cell responses that only partially control CMV replication, leading to viral reactivation and the establishment of a chronic inflammatory state that may promote chronic allograft rejection (4). Whilst improvements in antiviral prophylaxis strategies have clearly reduced the incidence of fulminant CMV-disease syndromes, subclinical CMV reactivation is still common (5,6) and remains a significant risk factor for accelerated chronic allograft rejection in lung transplant recipients (1).

In the setting of transplantation, CMV reactivation, inflammation and T-cell reactivity are likely to be intimately linked, as highlighted by studies that show that CMV reactivation is more likely to occur in the setting of acute allograft rejection (7), and reports that acute rejection may be aggravated following CMV reactivation, possibly as a consequence of infiltrating virus-specific CD8+ T cells (8).

Given the likely interrelationships between CMV reactivation, lung inflammation and alloreactivity there has been a pressing need for studies that move beyond clinico-epidemiological observation. Recently, insights into the replicative behavior of CMV have arisen from the introduction of molecular diagnostic techniques to measure CMV viral load and these have been applied in the setting of lung transplantation (5,9). We provide the first assessment of CMV-specific cytotoxic T lymphocytes (CTLs) both in the blood and lung allograft during the early post-transplant period. The findings reveal the complex interplay between immunosuppression, antiviral prophylaxis, CMV reactivation and their impact on the acquisition and maintenance of CD8+ T-cell immunity in the context of solid organ transplantation.

Materials and Methods

Lung transplant recipients

Twenty-two patients (14 male and 9 female, age range 20–64 years) had appropriate human leukocyte antigen (HLA) typing (HLA-A2+, n = 14; HLA-B8+, n = 6 and positive for both HLA-A2+ and -B8+, n = 2) for cross-sectional analysis of CMV-specific CTL subsets using HLA-restricted MHC class I tetramers. A longitudinal analysis was performed on a smaller subset of 14 patients, given that at 90 days post-transplant 8 patients moved interstate and were thus unavailable for further analysis. Donor (D) and recipient (R) CMV serostatus profiles were as follows: D+/R+, n = 10; D+/R–, n = 7; D–/R+, n = 3; D–/R–, n = 2. All lung transplant recipients received standard triple-therapy immunosuppression consisting of prednisolone, azathioprine and cyclosporine. At-risk patients (donor and/or recipient seropositive for CMV) received antiviral prophylaxis for 5 months. The units protocol is for all patients to undergo surveillance bronchoscopy at regular intervals (2 weeks and 1, 2, 3, 6, 9 and 12 months post-transplantation) or according to clinical need with bronchoalveolar lavage (BAL) and transbronchial biopsy sampling. All patients gave written consent and the study was approved by the Alfred hospital ethics committee.

Blood and BAL samples

Blood samples were collected at the time of routine surveillance bronchoscopy. BAL samples were obtained as previously described (10). BAL samples were passed through a 100 μm filter, washed and resuspended (1 × 107 cells/mL) in supplemented RPMI 1640.

Generation of tetrameric complexes

Tetramers specific for A2- (NLVPMVATV) or B8- (QIKVRVDMV) restricted epitopes of CMV were generated as described previously (11) with minor modifications. Tetramers were titrated to optimize staining and specificity was confirmed with patients of known HLA haplotypes. The limit of detection by tetramers was 0.05% of CD8+ T cells.

Ex vivo detection of CMV-specific CTL using MHC class I tetramers

Freshly isolated 100 μL whole blood samples (or 100μL BAL samples) were incubated with PE-conjugated tetrameric complexes in combination with the following directly conjugated monoclonal antibodies (mAbs): CD8-PerCP, CD27-FITC and CD28-APC (all Becton Dickinson, San Jose, CA). Red blood cells were removed with FACS Lysing solution (Becton Dickenson) before analysis on a FACS Calibur (Becton Dickinson) flow cytometer, set on a lymphocyte-gated population, and using FlowJo software (Treestar, San Carlos, USA). Results are presented as percentage of tetramer positive cells within the CD8+ population.

Ex vivo detection of CMV-specific CTL by intracellular γ-IFN staining

Intracellular cytokine analysis was performed essentially as previously described (12). Peripheral blood mononuclear cells (PBMCs) or BAL cells were stimulated with 1 μm HLA-A2- (NLVPMVATV [13]) and B8- (QIKVRVDMV [14]) restricted peptides for 6 h in the presence of Brefeldin A (10 μg/mL) (Sigma Aldrich, Sydney, Australia). Cells were then washed prior to staining with γ-IFN-FITC and CD8-PerCP. All assays were performed in duplicate. Cells were incubated with PMA (Sigma Aldrich, Australia, 10 ng/mL)/ionomycin (1 μM, Sigma Aldrich, Australia) and PBS as positive and negative controls, respectively. A subset of patients had intracellular staining performed on whole blood as previously described (15), in parallel with the PBMC assay, to assess the possible impact of immunosuppressive drugs. Cells were analyzed on a FACS Calibur and results expressed as percentage γ-IFN+ cells within the CD8+ population.

Quantification of CMV load

CMV load was measured in plasma and BAL samples with the semi-automated COBAS Amplicor CMV monitor test (Roche Diagnostic Systems, NSW, Australia) as described elsewhere (16). CMV load was measured in plasma as copies per milliliter and in the BAL as copies per milliliter of BAL supernatant.

Statistical analysis

Numerical data were expressed as medians and ranges. Group comparisons were performed using the Spearman's rank correlation coefficient. Between-group analysis was made with the non-parametric Mann-Whitney and Wilcoxon ranked test for unpaired and paired data, respectively. Statistical significance was defined as p < 0.05.

Results

CMV-specific CTL can be detected in the blood of immunosuppressed lung transplant recipients

To dissect out the relationship between cellular immunity to CMV and viral replication, 22 HLA-A2 and/or -B8 lung transplant recipients were analyzed for the presence of CMV-specific CD8+ T cells in the blood using MHC class I tetramers (Figure 1A) or by staining for intracellular IFN-γ following peptide stimulation (data not shown). An initial cross-sectional study was performed on all patients early post-transplant (<90 days) while they were still receiving antiviral prophylaxis. CMV-specific CD8+ T cells could be detected in 12/22 lung transplant recipients, with frequencies ranging from 0.05% to 18.1% (Figure 1B). Of the 10 lung transplant recipients with undetectable CMV-specific CTL, two were at low risk of CMV disease as both donor and recipient were seronegative for CMV. Conversely, seven CMV seronegative recipients were at ‘high-risk’ of CMV disease having received a transplant from a seropositive donor. In three of these ‘high-risk’ patients CMV-specific CTL were apparent following the cessation of antiviral prophylaxis at 5 months post-transplant, while in the other four patients' analysis was limited to the first 90 days post-transplantation because long-term follow-up was at a distant center. In a parallel analysis, there was a tight correlation between the frequencies of CMV-specific CTL as assessed by tetramer staining or by intracellular staining for IFN-γ following in vitro stimulation with peptides encoding specific T-cell epitopes (r = 0.90, p < 0.001), with the frequency of γ-IFN producing CMV-specific CD8+ T cells (following CMV-specific stimulation) being 75% (median, range 35–97%) of the frequency of circulating CMV-specific CTL as measured by tetramer binding (data not shown).

Figure 1.

CMV-specific CTL in the blood of lung transplant recipients at risk of CMV reactivation. Whole blood from lung transplant recipients were stained with CMV-tetramer and CD8-specific mAb and then analyzed by flow cytometry. (A) Dot plot showing CD8+ and tetramer staining of peripheral blood mononuclear cells from a CMV-seropositive lung transplant recipient. (B) Cross-sectional analysis of the frequency of CMV-specific tetramer cells in 12 HLA-A2 or HLA-B8 lung transplant recipients. All patients were within 90 days of transplant, receiving antiviral prophylaxis and were at risk of CMV reactivation as assessed by their donor (D)/recipient (R) CMV serostatus (D+/R+, n = 10; D–/R+, n = 2). Data shown represent the percentage of CMV-tetramer positive cells of gated CD8+ cells.

Frequencies of CMV-specific CD8+ T cells reflect viral reactivation profiles in CMV seropositive lung transplant recipients

To determine the impact of both immunosuppression and antiviral prophylaxis on CMV-specific immunity, 10 patients were followed longitudinally from the time of transplantation.

In 9/10 CMV seropositive lung transplant recipients, the proportion of CMV-tetramer positive CD8+ T cells in the blood remained relatively stable over time, despite the cessation of antiviral prophylaxis at 5 months post-transplant (Figure 2, dashed line). In five patients, the proportion of CMV-specific CTL was also assessed in pre-transplant blood samples. There was no significant difference in the frequency of CMV-specific CD8+ T cells between the pre- and post-transplant time points, suggesting that the introduction of broad spectrum immunosuppression had little impact on CMV-CTL frequency. In contrast, the CMV-specific CTL response to CMV reactivation was illustrated in the one CMV seropositive lung transplant recipient who demonstrated CMV DNA in the lung allograft (31 000 copies/mL BAL) on cessation of antiviral prophylaxis. The frequency of CMV-specific CD8+ T cells was higher following viral reactivation in the lung allograft (3.1% vs. 6.8% CMV-specific CTL pre- and post-CMV reactivation, respectively) (Figure 2, solid line).

Figure 2.

In the absence of CMV reactivation CMV-specific CTL frequencies remain stable. Whole blood from 10 lung transplant recipients were stained at the indicated times (post-transplantation) with CMV-tetramer and CD8-specific mAb and the frequency of CMV-specific CD8+ T cells determined by flow cytometry. Data from patients who had no detectable CMV-reactivation are indicated with a dotted line. One patient experienced CMV reactivation at day 182 (bold arrow) post-lung transplant and the corresponding data are indicated by bold line.

CMV-specific CTL can be detected in the lung allograft in the absence of CMV reactivation

As viral replication in the lung allograft is associated with chronic rejection, it was important to determine whether CMV-specific CTL were also present in the lung. Of the 12 patients who had CMV-specific CTL in the blood, nine also had CMV-tetramer positive CD8+ T cells detected in the lung allograft, as measured by BAL sampling (Table 1). At no instance were CMV-specific T cells detected in the BAL but not in the blood. In general, the proportion of CD8+ T cells that were CMV-specific tetramer positive was higher in the blood than in the BAL. In four of these patients, the donor lung shared either HLA-A2 or -B8 with the recipient and thus tetramer positive T cells could recognize both CMV-derived peptides presented by both the allograft and infiltrating recipient antigen-presenting cells (APCs). However, in five lung transplant recipients who had CMV-specific CTL detected in the lung allograft the donor lung lacked either HLA-A2 or -B8. Finally, the size of the BAL CMV-specific CTL pool in any one patient was not related to donor/recipient CMV serostatus or episodes of CMV reactivation (Table 1).

Table 1. Characteristics of lung transplant recipients with CMV-tetramer positive CD8+ T-cells measured in both the blood and BAL
Patient1Post-transplant (days)Recipient HLA2Donor HLACMV serostatus (D/R)CMV reactivationBlood-CMV specific CTL3 (%)BAL CMV- specific CTL3 (%)
  1. 1Cross-sectional analysis of CMV-specific CD8+ T cells in the BAL was performed at the time points specified above. Longitudinal analysis of CMV-specific CTL for patients 1–2 and 3–7 are also represented in Figures 3 and 2, respectively.

  2. 2All patients were studied with either a HLA-A2 or HLA-B8 CMV-specific tetramer to enumerate CMV-specific CD8+ T cells in the blood and BAL.

  3. 3Percentage of CD8+ T cells that were CMV-specific tetramer positive.

1189A2, A2, B44, B60A2, A32, B41, B41+/–Yes7.45.4
2287A2, A2, B44, B44A1, A3, B8, B35+/–Yes201.2
335A2, A23, B4, B60A2, A24, B18, B62+/+No0.32.8
484A1, A13, B8, B44A30, A31, B39, B60–/+No18.12.5
584A2, A30, B13, B44A2, A19, B7, B44–/+No5.72.3
6189A2, A3, B13, B70A3, A9, B49, B18+/+No0.80.2
7428A1, A1, B8, B8A2, A31, B44, B60+/+Yes3.10.9
8371A1, A24, B8, B49A1, A30, B8, B35+/+No0.60.7
9266A1, A3, B7, B8A11, A26, B27, B35+/–No0.60.5

Expansion of CMV-specific CD8+T cells precedes CMV reactivation in primary CMV-mismatched lung transplant recipients

A second pattern of CMV-specific immunity was seen in primary CMV-mismatched lung transplant recipients. Detailed analysis of these two patients gave an insight into how the primary immune response to CMV developed not only in the blood but also in the lung allograft (Figure 3). While both these patients were HLA-A2+, the first patient received a lung allograft from a HLA-A2 donor, whilst the second patient received a totally HLA-mismatched allograft.

Figure 3.

The appearance of CMV-specific CTL in the lung allograft is delayed following lung transplantation, but precedes the appearance of measurable CMV reactivation, and persists following viral clearance. Longitudinal analysis of two lung transplant recipients who were primary CMV mismatches (D+/R–) is shown. Both patients were HLA-A2+ and were studied with a HLA-A2 CMV-specific tetramer. The first patient received a lung allograft from a HLA-A2 donor, whilst the second patient received a totally HLA-mismatched allograft. At the indicated time points, post-transplantation whole blood and cells obtained from BAL were stained with CMV-tetramer and CD8-specific mAb and then analyzed by flow cytometry. The CMV viral load in the BAL (–▪–), the percentage of CMV-tetramer positive CD8+ T cells in the blood (–▴–) and in the BAL (–○–) are shown on the y-axis. Detection of histological evidence of CMV infection (arrowheads) and period of antiviral prophylaxis (shaded rectangle) are indicated.

In the first patient, despite the fact that CMV was introduced at the time of transplant, virus-specific CD8+ T cells were first detected in the blood and in the lung allograft 120 days post-transplant. However, despite the presence of CMV-specific CTL, viral reactivation (20 000 copies/mL BAL) was observed following withdrawal of valganciclovir. Subsequently, the patient was diagnosed with CMV pneumonitis, on day 188, at which point the high viral load (100 000 copies/mL BAL) was associated with the detection of CMV inclusions on transbronchial biopsy. During this period CMV was not detected in the blood. Corresponding to the increase in viral antigenemia in the lung allograft, the frequency of CMV-specific CTL increased both in the blood and BAL at this point. The patient was treated with a 2-week course of i.v. ganciclovir, and at repeat bronchoscopy CMV DNA was not detected in the lung allograft. Of note, CMV-specific CTL continued to circulate through the lung allograft despite undetectable CMV DNA levels (as measured by quantitative PCR), and indeed, at 480 days failed to control low-level viral reactivation (9100 copies/mL BAL) in the lung allograft.

A similar pattern was observed in the second patient, in whom CMV-specific CD8+ T cells were detected in the blood prior to CMV reactivation in the lung allograft and represented a significant proportion of the CD8+ T-cell population (range 11.5–21% of total CD8+ T cell) even following viral clearance. In contrast, the frequency of the HLA-A2-restricted CMV-specific T cells was much lower, but still detectable, in the HLA-A2-mismatched lung allograft.

Progressive down-regulation of co-stimulatory molecules on CMV-specific CD8+T cells following primary CMV infection

A linear pathway of CD8+ T-cell differentiation has been proposed in which activation of naïve CD8+ T cells leads to the sequential down-regulation of CD28 and CD27 (17). Following primary CMV infection, very few CMV-specific T cells in the blood expressed CD28 even at the earliest time points, while a progressive reduction in CD27 expression was observed (Figure 4A). Extending the phenotypic analysis to all 14 lung transplant recipients with demonstrable CMV-specific immunity, it is illustrated that the expression of CD28 on blood CMV-specific CD8+ T cells was quite variable with particular low expression seen in the two patients with primary CMV infection (Figure 4B, solid line). Additionally, in a number of patients we were able to demonstrate that CD28 expression on blood CMV-specific CD8+ T cells was not significantly different to its expression on the same cells in the BAL. Finally, we went on to demonstrate that the size of the CMV-specific CTL pool in the blood was correlated to the overall expression of CD28 on the CD8+ T-cell population as a whole (r =–0.78, p < 0.001, Figure 4C).

Figure 4.

Following primary CMV infection CMV-specific CD8+ T cells have an effector phenotype. (A) whole blood from lung transplant recipients was stained at the indicated time points with CMV tetramer, mAb specific for either CD8 and CD28 or CD8 and CD27 and analyzed by flow cytometry. Plots show the percentage of CMV tetramer positive CD8+ cells that express either CD27 or CD28. The initial detection of a CMV DNA by quantitative PCR is indicated by the arrowhead. (B) Cross-sectional analysis of CD28 expression on blood CMV-specific CD8+ T cells from 14 lung transplant recipients. Paired analysis performed at the same time on BAL CMV-specific CTL showed low levels of CD28 expression following primary CMV infection post-transplant (n = 2 patients, bold line) compared to the higher expression seen in transplanted patients who were CMV serospositive at the time of transplant (n = 2 patients, dashed line). (C) The expression of CD28 on CD8+ T cells in any one CMV-seropositive lung transplant recipient is inversely related to the size of the CMV-specific CTL pool in that individual (r =–0.78, p < 0.001).

Discussion

Despite the significant morbidity associated with CMV, the CMV-specific T-cell response has not been described following lung transplantation. The longitudinal assessment of CMV-specific CTL in a cohort of lung transplant recipients on broad spectrum immunosuppression revealed that not only could CMV-specific CTL be detected but that distinct patterns of CMV-specific immunity could be identified that were related to the dynamics of CMV replication in any one individual. The frequency of tetramer positive CD8+ T cells in lung transplant recipients was similar to that of γ-IFN producing T cells; a tight correlation suggestive of an intact and functioning antiviral immune response given that cytokine production has been shown to be correlated to other markers of cytotoxicity (18).

By prospectively following patients from the time of transplantation, we have been able to assess the impact that immunosuppressive drugs, antiviral prophylaxis and CMV reactivation have on determining patterns of CMV-specific T-cell immunity. In the absence of CMV reactivation, the frequency of CMV-specific CTL in the blood remained stable during the early post-transplant period, despite changing levels of immunosuppressive and antiviral drugs. In this setting, viral replication is either being suppressed by a functional antiviral immune response and/or the antiviral drug, valganciclovir. The observation that viral DNA was not detected in either the blood or BAL following the withdrawal of antiviral drugs suggests that viral latency is maintained by an active immune response (Figure 2). The impact of CMV reactivation was illustrated in one patient in whom the frequency of CMV-specific CTL in the blood increased after CMV reactivation occurred on withdrawal of antiviral prophylaxis.

The longitudinal analysis of CMV-mismatched lung transplant recipients allowed an insight into how the primary immune response to CMV develops not only in the blood but also in the lung allograft (Figure 3). Although CMV was introduced via the donor lung, at the time of transplant the appearance of CMV-specific CTL was delayed in both patients. This is in contrast to what has been described following primary CMV infection in renal transplant recipients where in the absence of routine antiviral prophylaxis, CMV-specific CD8+ T cells can usually be detected within 30 days of transplant (19). The delayed development of CMV-specific CTL demonstrated in the current study most likely reflects the ability of antiviral drugs to suppress viral replication and hence the antigenic load, allowing CMV to remain ‘hidden’ from the circulating naïve T-cell pool. Although less likely, the immunosuppressive effects of the antiviral prophylaxis may also contribute to a delay in the development of CMV-specific CTLs (20). Finally, while not analyzed in the current study, CMV-specific CD4+ T cells are known to be impaired following lung transplantation, and thus any functional impairment of T helper cells will impact not only on CD8+ T-cell differentiation but also on CMV reactivation (21). The data, however, demonstrate that despite low levels of CMV-antigen present at this time, a CMV-specific T-cell response can nevertheless be initiated despite extensive immunosuppression.

The presence of CMV-specific CTL in the lung allograft is of particular interest in light of the seminal work of Doherty and Zinkernagel (22). The paradigm that virus-specific CTL can only eliminate infected cells by recognizing viral peptides bound to self-MHC class I molecules raises several important questions in the lung transplant setting, where HLA matching does not routinely occur. In the MHC-incompatible lung allograft, it is unclear how self-restricted CMV-specific CTL can contribute to controlling the spread of virus. Thus in our study, it was of particular interest to discover that we were able to visualize CMV-specific CD8+ T cells restricted by HLA class I alleles of the recipient in the transplanted lung. The role of these HLA-A2 and -B8-restricted T cells in the allograft is unclear. Their presence may simply represent a non-specific influx of lymphocytes in response to inflammation (23). Alternatively, the T cells may recognize viral antigens that are either directly or cross-presented by host bone marrow-derived cells that have seeded in the lung allograft (24). Finally, these T cells may be actively contributing to graft rejection via cross-reactivity with allogenic MHC class I molecules (25,26).

In recent years, the phenotypic analysis of viral-specific T cells has led to significant advances in our understanding of how the immune system responds to chronic persistent antigens. Given that most of these studies describe T-cell responses in the blood, there is a paucity of information on the function of human viral-specific T cells in peripheral target organs. The need to perform regular surveillance bronchoscopy in lung transplant recipients therefore provides an ideal opportunity to study viral-specific immune responses in the lung. Contrary to what might have been expected, CMV-specific CD8+ T cells in the BAL appeared no more differentiated (CD28) than those in the blood, suggesting that cytotoxic CD28CD8+ T cells were not being redistributed to and sequestered at the site of active infection, as has been described in animal models (27).

It is recognized that antigen-experienced CD8+ T cells exposed to persistent antigen downmodulate the costimulatory molecules CD28 and CD27 (28). We studied the expression of CD27 and CD28 on CMV-specific CD8+ T cells following primary CMV infection, and showed a progressive decrease in CD27 expression possibly reflecting the acquisition of effector functions, such as cytotoxicity (29). In contrast to CD27 expression, the cell surface expression of CD28 was already down-regulated prior to the first analysis (Figure 4A). This differs from the more gradual down-regulation of CD28 on CMV-specific CTL following primary CMV disease in renal transplant recipients (19). Given that differentiation to an effector phenotype is driven by antigenic load (30), this difference in CD28 down-regulation may be explained by the relatively high inoculum of CMV that is introduced with the donor lung at the time of transplant (31). The potential impact of antigenic load also needs to be considered given the variable expression of CD28 on CMV-specific CTL that was seen on cross-sectional analysis (Figure 4B). The development of antiviral T-cell immunity following primary CMV infection in the immunosuppressed lung transplant recipient may be driven by high CMV viral load (as was the case in patient 1), which in turn explains the low levels of CD28 on CMV-specific CTL (Figure 4B, solid lines). Finally demonstrating that the size of the CMV-specific CTL pool influences the net expression of CD28 on the CD8+ T-cell population in the peripheral blood is relevant given a growing body of work suggesting that expansion of a CD28CD8+ pool may be associated with immune dysfunction (32).

In summary, in the present study we have described for the first time the dynamics of CD8 T-cell immune responses to CMV, both in the peripheral blood and lung allograft of a cohort of lung transplant recipients with and without CMV reactivation. In primary CMV-mismatched lung transplant recipients, we have shown that CMV-specific CTL develop prior to viral reactivation and persist in the absence of detectable viral DNA. Finally, the persistence of CMV-specific CTL with an effector phenotype in both the HLA-matched and -mismatched donor lung allograft in the absence of detectable viral reactivation, suggests that these cells may have a role in controlling subclinical CMV reactivation.

Acknowledgments

The authors wish to thank Sue Fowler for clinical assistance with patients in this study; Dr. Jie Lin who constructed the tetramers in the lab; Dr. Lucy Sullivan and Magda Wojtasiak for proofreading of the manuscript; and the patients who kindly agreed to participate in this study. Glen Westall is a recipient of a National Health and Medical Research Council (NHMRC) postgraduate research scholarship. This study was supported by grants received from the Marion and EH Flack Trust and the National Health and Medical Research Council.

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