Biology of Blood and Marrow Transplantation
Volume 16, Issue 7 , Pages 994-1004, July 2010

Protective Immunity Transferred by Infusion of Cytomegalovirus-Specific CD8+ T Cells within Donor Grafts: Its Associations with Cytomegalovirus Reactivation Following Unmanipulated Allogeneic Hematopoietic Stem Cell Transplantation

  • Xiao-Hua Luo

      Affiliations

    • Department of Onco-Haematology, Children's Hospital of Chongqing Medical University, Chongqing, China
    • ,
  • Xiao-Jun Huang

      Affiliations

    • Peking University Institute of Hematology, People's Hospital, Beijing 100044, China
    • Corresponding Author InformationCorrespondence and reprint requests: Xiao-Jun Huang, Peking University, People's Hospital, Institute of Hematology, 11 Xizhimen South Street, Beijing 100044, China.
    • ,
  • Kai-Yan Liu

      Affiliations

    • Peking University Institute of Hematology, People's Hospital, Beijing 100044, China
    • ,
  • Lan-Ping Xu

      Affiliations

    • Peking University Institute of Hematology, People's Hospital, Beijing 100044, China
    • ,
  • Dai-Hong Liu

      Affiliations

    • Peking University Institute of Hematology, People's Hospital, Beijing 100044, China

Received 25 November 2009; accepted 5 February 2010. published online 17 February 2010.

Article Outline

Human cytomegalovirus (CMV)-specific cytotoxic T lymphocyte (CTL) immune response must be reconstituted for long-term protection against CMV relapse and disease in hematopoietic stem cell transplantation (HSCT) recipients. We phenotypically quantitated absolute numbers of CMV-pp65 peptide-specific CTLs (CTLCMV) in 50 related donor unmanipulated allografts infused into HLA-matched or -mismatched recipients and examined the incidence of CMV reactivation. High CTLCMV with terminally differentiated effector CD45RO-CD62L- cell (TEMRA) phenotype in the allografts were associated with reduced risk of CMV reactivation, in the presence of sufficient CD45RO+CD62L- cell (TEM) infusion (≥0.208 × 106/kg). Early after transplantation, there was significant expansion of CTLCMV with the central memory CD45RO+CD62L+ cell (TCM) phenotype when CMV was reactivated. The frequencies of CTLCMV TNaive (CD45RO-CD62L+), TCM, and TEM at day 90 posttransplantation and of CTLCMV TEMRA at day 60 posttransplantation were greater in recipients with higher infusions of CTLCMV TEMRA, suggesting protective immunity transferred by infusion of CTLCMV within allografts. Moreover, the majority of the CTLCMV identified in the recipients early after HSCT was of donor origin. Our findings support that measuring levels of CTLCMV and its subsets in the donor grafts and manipulating these cells early after transplantation may help control CMV reactivation, which is closely correlated with immune reconstitution and differentiation of CTLCMV subsets.

Key Words: Allogeneic hematopoietic stem cell transplantation, CMV reactivation, Antigen-specific CTL, Donor grafts

 

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Introduction 

Human cytomegalovirus (CMV) infection remains a challenging problem after allogeneic hematopoietic stem cell transplantation (HSCT) 1, 2. The use of prophylactic and preemptive antiviral chemotherapy strategies has significantly reduced the morbidity and mortality of CMV infection 3, 4, 5; however, CMV-specific cytotoxic T lymphocyte (CTL) immune response must be reconstituted to confer long-term protection against CMV relapse and disease. Studies using tetramer staining combined with fluorescent antibodies to the variable region of the T cell receptor and single-nucleotide DNA polymorphism analysis suggest that CMV-specific CD8+ T lymphocytes detected in CMV-seropositive recipients of allogeneic HSCT from CMV-seropositive donors are likely of donor origin and expand in the recipient after antigenic stimulus because of CMV reactivation 6, 7. Gratama et al. [8] reported CMV-specific T lymphocytes infused with the grafts correlated inversely with the number of preemptive courses of ganciclovir administered after HSCT. These reports indicate that protective immunity can be successfully transferred to patients through infusion of donor-derived antigen-specific CD8+ CTLs within allografts if the donor has immunity to the specific antigen.

Few immunocompromised HSCT recipients with a CMV-specific CD8+ cell level >2-10 × 106/L developed CMV disease 8, 9. Studies of the correlation between CTL frequency and CMV reactivation have provided inconsistent results, however 10, 11, 12. These studies suggest that control of human CMV infection might require the presence of functional CMV-specific T cells, rather than the ability to recover sufficient numbers. Although adoptive transfer of CMV-specific T cells has shown promising results for controlling or preventing CMV reactivation, the most desirable phenotypic profile for T cells remains unclear 13, 14, 15, 16.

The foregoing findings demonstrate that the magnitude of CMV-specific CD8+ T cell response alone is not an appropriate predictor of clinical outcome. A recent study [17] found that different memory T cell subsets, defined according to phenotype, had different functional capabilities and roles 18, 19, and that their relative contributions to the overall size of the response may relate to overall efficacy. This led us to propose that the general phenotype of the CMV-specific CD8+ T cell response also might be linked to its antiviral capacity.

Based on CD45RO and CD62L surface marker expression 18, 19, 20, the following linear differentiation of antigen-experienced CD8+ T cells has been proposed: naive CD45RO-CD62L+ T (TNaive) cells, central memory CD45RO+CD62L+ T (TCM) cells, effector memory CD45RO+CD62L- T (TEM) cells, and terminally differentiated effector CD45RO-CD62L- T (TEMRA) cells. TEM and TEMRA cells can produce high levels of perforin, whereas virus-specific TCM cells have little direct antiviral activity, but likely serve to replenish and sustain CD8+ TEM cell populations through secretion of interleukin-2 and proliferation. A study of human immunodeficiency virus (HIV) infection found a link between HIV-1 specific CD8+ TEMRA cell level in early infection and control of HIV-1 viremia [17]. Distinct patterns of CD8+ memory cell subsets also might affect the risk of CMV reactivation [21]. In the present study, we explored whether immunophenotypic composition of donor grafts, especially levels of CMV-specific CD8+ T cells, was correlated with the overall potency of the response and predicted control of CMV infection in 50 HLA-matched or -mismatched related HSCT recipients.

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Materials and Methods 

Patients and Donors 

We studied 50 consecutive patients and their donors who had either HLA-A∗0201 or HLA-A∗2402 alleles, in whom the recipients with a malignant hematologic disorder underwent unmanipulated allogeneic HSCT from an HLA-matched or -mismatched related donor between March 2007 and April 2008 at Peking University Institute of Hematology. All of these patients were CMV serologically donor-positive/recipient-positive (D+/R+) pretransplantation, and were given both granulocyte colony-stimulating factor (G-CSF)-primed peripheral blood grafts (G-PBs) and G-CSF–primed bone marrow grafts (G-BMs) at transplantation. The HLA-mismatched transplant recipients had no HLA-identical related or unrelated donors.

We included all of the donors typed as HLA-A∗0201+ or HLA-A∗2402+ using high-resolution techniques. The combined frequencies of these 2 alleles encompass >30% of the Chinese population 22, 23. All recipients and their donors under HLA-matched or -mismatched related transplantation were matched on these 2 alleles. The Institutional Review Board of Peking University approved this study, and all 50 recipients and their respective stem cell donors provided written informed consent. Because of sample limitations, CMV-specific CTL reconstitution could be analyzed in only 44 of the recipients. Clinical characteristics of these recipients and their donors are summarized in Table 1.

Table 1. Clinical Characteristics of Recipients and Donors
All patients, n50
Acute lymphoblastic leukemia, n10
Acute myelogenous leukemia, n23
Biphenotypic acute leukemia, n1
Chronic myelogenous leukemia12
Myelodysplastic syndrome, n4
High risk/standard risk, n∗13/37
Patient age, years, median (range)33.5 (14-55)
Males/females, n30/20
Donor graft composition,
CD34+, 106cells/kg2.020 (0.708-50.143)
CD3+, 108cells/kg1.536 (0.229-3.504)
CD8+, 108cells/kg0.589 (0.091-1.506)
CD8+pentamer+ (CTLCMV), 106cells/kg0.671 (0.011-16.677)
CTLCMV CD45RO-CD62L+ (TNaive), 106cells/kg0.015 (0-0.605)
CTLCMV CD45RO+CD62L+ (TCM),106cells/kg0.008 (0-0.302)
CTLCMV CD45RO+CD62L- (TEM), 106cells/kg0.208 (0-3.324)
CTLCMV CD45RO-CD62L- (TEMRA), 106cells/kg0.177 (0-13.343)

CTLCMV indicates cytomegalovirus-pp65 peptide-specific cytologic T lymphocyte; n, number.

∗High risk: Patients with acute leukemia beyond second remission, not in remission, or Philadelphia chromosome–positive; myelodysplastic syndrome–refractory anemia with excess of blasts in transformation or myelodysplastic syndrome–acute myelogenous leukemia; chronic myelogenous leukemia in accelerated phase or blast crisis phase; or biphenotypic acute leukemia. Standard risk: all others.

All patients received myeloablative conditioning and standard prophylaxis for graft-versus-host disease (GVHD) according to HLA disparity, as reported previously 24, 25. Acute GVHD (aGVHD) treatment was provided as described previously 26, 27. In brief, aGVHD was treated with methylprednisolone (0.5-1 mg·kg−1·d−1). GVHD manifesting in the skin was treated with a combination of methylprednisolone and methotrexate or methotrexate alone. In patients with inadequate or no response to primary therapy for GVHD, 1 mg·kg−1 of anti-Tac monoclonal antibody (Daclizumab; Roche, Basel, Switzerland) was administered i.v. on days 1, 4, and 8, and subsequently at 7-day intervals, for a total of 3-6 doses.

Graft Harvesting and Graft Composition 

Donors were mobilized with recombinant human G-CSF (rhG-CSF) (Filgrastim; Kirin, Japan), 5 mg/kg daily, injected s.c. for 5 or 6 consecutive days. Stem cells from BM were collected on the fourth day of mobilization, and peripheral blood stem cells (PBSCs) were collected on day 5-6. The target mononuclear cell (MNC) count was 3-4 × 108 per kg of recipient weight. BM was infused to patients within 3 hours after collection, and PBSCs were infused fresh and unmanipulated.

Samples from grafts (G-PBs and G-BMs) were stained with monoclonal antibodies for flow cytometry analysis of surface antigens CD45, CD3, CD8, and CD34. CD34+ cell enumeration was performed at Peking University People's Hospital, as described by Luo et al. [28]. The absolute numbers of CD3+ and CD8+ T cells were calculated and divided by actual patient weight to determine the cell dose per kg. Data acquisition and analyses were performed using CellQuest software (BD Biosciences, San Jose, CA).

Enumeration of CMV-Specific CTLs 

Venous blood samples were collected with sodium heparin before HSCT and at 30 days, 60 days, and 90 days post-HSCT. To serve as reference samples, venous blood samples were obtained from 12 HSCT donors before mobilization. HLA-A∗0201/NLVPMVATV (human CMV pp65 495-504) or HLA-A∗2402/QYDPVAALF (human CMV pp65 341-349) phycoerythrin-labeled Pro pentamers (ProImmune, Oxford, United Kingdom) was added to unfractionated whole blood at room temperature for 10 minutes. The secondary antibodies (CD8-Percp, CD45RO-APC, and CD62L-FITC; BD Biosciences) and corresponding isotype antibodies were added for the final 20 minutes of incubation, followed by lysis of red cells using ammonium chloride lysing solution. Cells were then fixed in 1% paraformaldehyde in phosphate-buffered saline and analyzed within 12 hours on a FACScan flow cytomer (BD Biosciences). After acquisition of 20,000 CD8+ lymphocytes (bright, low to intermediate forward scatter, and low side scatter), the proportion of CD8+ T cells binding pentamers was assessed with the unstained control subtracted (Figure 1 in the supplementary data). The absolute number of circulating CMV-specific CD8+ T cells was calculated from the proportion of CD8+ T cells binding pentamers and the simultaneously obtained absolute CD8+ T cells count. The phenotypically distinct memory CD8+ T cell subsets were characterized by the expression of CD45RO and CD62L as well. Blood samples from normal donors, negative for the restricting HLA type, served as additional negative controls.

In the stem cell grafts, the number of transplanted CMV-specific CD8+ T cells/kg body weight of recipient was calculated from the proportion of CD8+ T cells binding pentamers plus the simultaneously established number of CD8+ T cells/kg transplanted. The absolute numbers of CTL subsets (TNaive, TCM, TEM, and TEMRA) were calculated accordingly.

CMV Testing and Preemptive Treatment 

Donor/recipient serologic status was assessed by enzyme-linked immunosorbent assay before transplantation and by plasma CMV DNA testing, using real-time polymerase chain reaction (PCR) to monitor for CMV reactivation, throughout the study period (kits purchased from Sino-American Biotech, Beijing, China). CMV-positive patients (>6 × 102 copies/mL) usually received preemptive i.v. ganciclovir 5 mg/kg twice daily for 10-14 days or until CMV tests were negative. In patients with neutropenia, 2 doses of foscarnet 90 mg/kg i.v. were administered in place of ganciclovir. All patients received prophylactic ganciclovir (5 mg/kg twice daily i.v.) from day -9 to day -2 and acyclovir (400 mg orally) from day +1 to day +180. A patient's first positive CMV PCR result was considered an event for the purpose of this analysis, and time to first positive CMV test result was calculated from the date of HSCT until the date of the first positive test. CMV disease was diagnosed according to previously published criteria [29].

Statistical Methods 

Initial univariate exploration of the baseline covariates was performed using the 2-sided Fisher's exact test, Mann-Whitney U test, or Wilcoxon's test. To determine which factors independently correlated with CTLCMV content in allografts, a multivariate linear regression model was constructed using donor characteristics including age (< or ≥38 years, according to median donor age), sex, weight (< or ≥68.5 kg, according to median donor weight), pregnancy, Epstein-Barr virus serostatus, and hepatitis B virus serostatus. Patient variables included age, sex, underlying disease, HLA disparity, BM transplantation (BMT) disease risk, and aGVHD. The cell type variables were CD34+, CD3+, CD8+, CTLCMV, CTLCMV TNaive, CTLCMV TCM, CTLCMV TEM, and CTLCMV TEMRA. All variables that were either statistically significant in univariate analysis or potentially important with respect to CMV reactivation (eg, content of CTLCMV and its subsets in donor grafts) were included in final models. The cell type variables were included with a cutoff at the 50th percentile. GVHD was modeled as a time-varying covariate. The covariates in the final model were evaluated for collinearity, using the variance inflation factor. Statistical analyses were done with Stata 10.0 (Stata Corp, College Station, TX) and GraphPad Prism software 5.01 (GraphPad, San Diego, CA).

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Results 

More CTLCMV Cells in G-PBs Compared with G-BMs 

Of the total CD8+ T cell population, a median of 0.48% (0-7.52%) were pentamer-positive in G-BMs, compared with 1.165% (0-22.22%) in G-PBs. No pentamer-positive T cells were detected in 14 G-BMs and 1 G-PBs. G-PBs had higher proportions of CTLCMV and CD45RO-CD62L- (TEMRA) (P <.001 and .03, respectively), whereas G-BMs had a higher proportion of CD45RO-CD62L+ (TNaive) (P = .003) (Table 2).

Table 2. Graft Product Composition
Cell TypeBone Marrow G-BM,G-PBPBSCs G-BM,G-PBP
CD34+, 106/kg0.67 (0.12-7.31)1.27 (0.16-42.83)<.001
CD3+, 108/kg0.15 (0.06-1.23)1.31 (0.08-3.37)<.001
CD8+, 108/kg0.05 (0.02-0.54)0.49 (0.03-1.39)<.001
CD8+pentamer+(CTLCMV), 106/kg0.01 (0-0.81)0.55 (0-16.68)<.001
CTLCMV CD45RO-CD62L+ (TNaive, %)7.86 (0-55.43)1.04 (0-50.91).003
CTLCMV CD45RO+CD62L+ (TCM, %)3.49 (0-39.33)1.63 (0-66.67).56
CTLCMV CD45RO+CD62L- (TEM, %)40.00 (3.13-94.15)42.44 (0-96.28).48
CTLCMV CD45RO-CD62L- (TEMRA, %)30.17 (0.14-78.57)41.84 (0-100).03

CTLCMV indicates cytomegalovirus-pp65 peptide-specific cytologic T lymphocyte.

On multivariate linear regression analysis, only donor age was negatively associated with CTLCMV content in allografts (β, -1.638; 95% confidence interval [CI], -3.116 to ∼-0.159; P = .031; data not shown); thus, this covariate was included in the subsequent analysis for CMV reactivation. Other premobilization donor parameters (ie, sex, weight, pregnancy, hepatitis B virus serostatus, and Epstein-Barr virus serostatus) were not associated with CTLCMV content differences in allografts.

CMV Reactivation During the First 100 Days after HSCT 

A total of 632 CMV-DNA PCR assays were performed during the first 100 days after HSCT, with a median of 11 assays per patient (range, 2-26). An initial positive CMV-DNA was detected in 31 patients (62%) at a median of 31 days after HSCT (range, 4-68 days). The crude incidence of CMV reactivation during this period were similar among patients across age, sex, underlying disease, steroid use, and BMT disease risk group (Table 3). Covariates associated with increased CMV-PCR positivity after HSCT were recipients of HLA-mismatched HSCT (P = .004) and those who experienced grade II-IV aGVHD (P = .049). There was no significant differences in graft composition, including CTLCMV content, between patients with or without CMV reactivation (P = .38-.94).

Table 3. Characteristics of Crude CMV Reactivation to Day +100 Post-HSCT
CharacteristicsnCMV reactivation, n%P
Total503162
Recipient age .61
<35 years261765.38
≥35 years241458.33
Recipient sex .41
Female201155.00
Male302066.67
Primary disease .61
Acute leukemia∗332266.67
Chronic myelogenous leukemia12650.00
Other†5360.00
HLA match .004
Mismatched donor342676.47
Matched donor16531.25
BMT disease risk group .53
Standard risk372259.46
High risk13969.23
Acute GVHD .049
Grade II-IV131184.62
None or grade I372054.05
Steroid therapy (≤ day 100) .36
>0≤1 mg/kg/day452964.44
>1 mg/kg/day5240.00
Graft composition
CD34+ <2.020 × 106/kg251456.00.38
CD34+ ≥2.020 × 106/kg251768.00
CD3+ <1.536 × 108/kg251664.00.77
CD3+ ≥1.536 × 108/kg251560.00
CD8+ <0.589 × 108/kg251664.00.77
CD8+ ≥0.589 × 108/kg251560.00
CTLCMV <0.671 × 106/kg251664.00.77
CTLCMV ≥0.671 × 106/kg251560.00
CTLCMV TNaive <0.015 × 106/kg241562.50.94
CTLCMV TNaive ≥0.015 × 106/kg261661.54
CTLCMV TCM <0.008 × 106/kg251560.00.77
CTLCMV TCM ≥0.008 × 106/kg251664.00
CTLCMV TEM <0.208 × 106/kg251560.00.77
CTLCMV TEM ≥0.208 × 106/kg251664.00
CTLCMV TEMRA <0.177 × 106/kg251768.00.38
CTLCMV TEMRA ≥0.177 × 106/kg251456.00

CTLCMV indicates cytomegalovirus-pp65 peptide-specific cytologic T lymphocyte; CMV, cytomegalovirus; GVHD, graft-versus-host disease; HSCT, hematopoietic stem cell transplantation.

∗Acute lymphoblastic leukemia and acute myelogenous leukemia.

†Biphenotypic acute leukemia and myelodysplastic syndrome.

CTLCMV TEMRA and TEM in Donor Grafts Correlates with Risk of CMV Reactivation 

Our multivariate proportional hazards model included donor age, donor–recipient HLA mismatch or match, aGVHD (grade 0-I vs II-IV) as a time-varying covariate, and graft composition (ie, CTLCMV TNaive, CTLCMV TCM, CTLCMV TEM, and CTLCMV TEMRA). CTLCMV was excluded because of marked colinearity in this model (variance inflation factor >10), and because CTLCMV was not significant in multivariate analysis (P = .593), even after the exclusion of its subsets (P = .795). In multivariate analysis (Table 4), HLA-matched (adjusted hazard ratio [HR], 0.212; 95% CI, 0.078-0.579; P = .002) remained significantly associated with CMV reactivation. CTLCMV TEM and CTLCMV TEMRA in allografts became significant for CMV reactivation in the adjusted model (adjusted HR, 4.386; 95% CI, 1.095-17.573; P = .037 and 0.233; 95% CI, 0.065-0.831; P = .025, respectively).

Table 4. Proportional Hazards Modeling of the Risk of CMV Reactivation after Allogeneic HSCT
CharacteristicMultivariate HR (95% CI)P
Donor age ≥38 years0.799 (0.346-1.848).600
HLA-matched donor0.212 (0.078-0.579).002
Acute GVHD grade II-IV0.996 (0.985-1.007).450
Graft composition
CTLCMV TNaive: CD45RO+CD62L- ≥0.015 × 106/kg0.840 (0.299-2.361).741
CTLCMV TCM: CD45RO-CD62L+ ≥0.008 × 106/kg0.799 (0.250-2.559).706
CTLCMV TEM: CD45RO+CD62L+ ≥0.208 × 106/kg4.386 (1.095-17.573).037
CTLCMV TEMRA: CD45RO-CD62L- ≥0.177 × 106/kg0.233 (0.065-0.831).025

CTLCMV indicates cytomegalovirus-pp65 peptide-specific cytologic T lymphocyte; GVHD, graft-versus-host disease; CMV, cytomegalovirus; HSCT, hematopoietic stem cell transplantation.

Most of the CMV-specific CD8+ T cells in donor grafts displayed either a TEMRA or a TEM phenotype. Thus, there was a close and inverse correlation between the two in the proportions of G-PBs and G-BMs (r = 0.57, P < .01 and r = 0.46, P = .046, respectively; Figure 2 in the supplementary data), but a positive correlation between the two in the absolute numbers of total grafts (r = 0.62, P < .01). This is a classic example of confounding, because higher TEMRA infusions were more likely in patients with higher TEM infusions at transplantation; thus, the crude univariate analysis for subsets of CTLCMV was severely confounded. In a separate analysis, we compared the absolute numbers of TEMRA cells infused in patients with or without CMV reactivation according to TEM content in allografts (above median or vice versa). We compared absolute numbers of TEMRA cells in the same way. Figure 1 shows that patients without CMV reactivation had more CTLCMV TEMRA infused at transplantation (P = .042). In contrast, no difference was found for CTLCMV TEM in the subgroup analysis. These findings suggest that the TEM level might be as crucial as TEMRA level in influencing risk of CMV reactivation.

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  • Figure 1 

    Relationship between TEMRA and TEM in allografts and CMV reactivation after HSCT. (A) The absolute numbers of TEMRA in allografts were significantly higher in patients with CMV reactivation in the presence of sufficient infusion of TEM (≥0.208 × 106/kg). (B) The absolute numbers of TEM in allografs were not significantly different in patients with CMV reactivation and those without CMV reactivation, regardless of the TEM content infused. Only significant P values are shown. “High” and “low” were introduced as above and below the median value, respectively (TEMRA, 0.177 × 106/kg; TEM, 0.208 × 106/kg).

CTLCMV Infusion at Transplantation and CTLCMV Reconstitution after HSCT 

CTLCMV reconstitution was prospectively monitored by immunophenotyping in 44 patients. Absolute counts of CTLCMV and its subsets were calculated at regular intervals (days +30, +60, and +90), whereas donor blood samples were evaluated at day −3 before mobilization as controls. The absolute numbers of CTLCMV and its subsets after HSCT were compared in patients with or without CMV reactivation (Figure 2). The absolute numbers of CTLCMV with TCM phenotype were significantly higher at day +30 and day +60 post-HSCT in patients with CMV reactivation (P <.05). No statistically significant difference in the percentage of circulating CTLCMV subsets was found between the 2 groups (Figure 3 in the supplementary data). The absolute counts of CD8+ T cells and CTLCMV were not statistically different between the 2 groups (Figure 2B and C); however, analysis of the correlation between CTLCMV reconstitution and CTLCMV TEMRA input showed higher numbers of CTLCMV TNaive, CTLCMV TCM, and CTLCMV TEM in peripheral blood at day +90 and CTLCMV TEMRA at day +60 when HSCT recipients received more CTLCMV TEMRA cells in allografts (P <.05; Figure 3).

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  • Figure 2 

    Expansion of CTLCMV TCM cells on CMV reactivation post-HSCT. Using flow-based frequency enumeration and absolute lymphocyte counts, absolute numbers of CTLCMV cells and subsets were calculated for healthy donors (n = 17) and HSCT recipients (n = 44) at regular intervals on days +30, +60, and +90. (A) The CTLCMV with TCM phenotype was significantly higher at days +30 and +60 post-HSCT in patients with CMV reactivation compared with patients without CMV reactivation. (B and C) The absolute counts of CD8+ T cells and CTLCMV were not statistically significantly different between the 2 groups. Bars indicate median values. Only significant P values are shown.

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  • Figure 3 

    Relationship between number of CTLCMV TEMRA cells transplanted and the recovery kinetics of circulating CTLCMV subsets after HSCT. Bars indicate median values. The frequencies of circulating CTLCMV TNaive, CTLCMV TCM, and CTLCMV TCM at day +90 and CTLCMV TEMRA at day +60 post-HSCT were higher in HSCT recipients with more CTLCMV TEMRA infused. Only significant P values are shown. “High TEMRA infusion” and “low TEMRA infusion” was introduced as above or below the median value of TEMRA cells in allografts (ie, 0.177 × 106/kg).

Most CTLCMV Identified in the Recipient Early after HSCT Were of Donor Origin 

To determine whether CTLCMV originate from donor cells or from recipient cells, the percentages of CTLCMV and CMV DNA copies were calculated for each post-HSCT sample time point. Recipients A and B underwent human CMV D+/R+ HLA-mismatched related donor allogeneic HSCT. Both recipients were HLA-A∗0201–positive, and both donors were HLA-A∗2402–positive. HLA-A∗0201/NLVPMVATV pentamer was used to trace CTLCMV from recipients, and HLA-A∗2402/QYDPVAALF pentamer was used to do so from donors. Figure 4 shows that most CTLCMV originated from donors early post-HSCT. CTLCMV from recipients remained at low levels until day +180, whereas CTLCMV from donors decreased slowly to its lowest point at day +90, and increased thereafter. Two patients experienced CMV reactivation between day +30 and day +60, and subsets of CTLCMV differed between donors and recipients.

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  • Figure 4 

    (A and B) Sources of CTLCMV responses in peripheral blood in relation to CMV infection as measured by CMV-DNA copies. The number of days after HSCT is shown on the x-axis. CTLCMV (HLA-A∗0201+ CD8+ T cells [patient origin; black] and HLA-A∗2402+ CD8+ T cells [donor origin; red]) are expressed as percentage of circulating CD8+ T cells (left, y-axis). CMV reactivation is expressed as CMV-DNA copies in peripheral blood (right, y-axis, green). (C-F) Relative proportions of CTLCMV cells in 4 distinct maturation stages (from least mature to more mature: TNaive, TCM, TEM, TEMRA) assessed by flow cytometry. The majority of CTLCMV cells were from donors early post-HSCT, and subsets of CTLCMV differed between donors and patients.

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Discussion 

Even in the antiviral era, CMV serostatus is a potent predictor of both CMV disease and death after HSCT [30]. CMV disease has been reported in 17.6% of D+/R+, 20.9% of D-R+, 5.3% of D+/R-, and 1.1% of D-/R- HSCTs [1]. When selecting donors, a CMV-seropositive donor to a CMV-seropositive recipient is clearly preferable, aided by increased levels of multifunctional CMV-specific T cells [30]. In this regard, evaluation of CTLCMV in donor grafts provides the most direct assessment of donor immunity to CMV antigen transferred to recipients.

Like other biologic markers, CTLCMV can be influenced by numerous factors. In elderly individuals, seropositivity for human CMV leads to the development of oligoclonal populations of CMV-specific CTLs. These CMV-specific CTLs have a highly polarized membrane phenotype, typical of effector memory cells [31]. Moreover, the majority of the clonally expanded virus-specific CD8+ cells in the elderly are dysfunctional 32, 33, explaining our finding that elderly donors had few CTLCMV TNaive in G-PBs.

Several previous studies have presented persuasive evidence that CD8+ CTLs play an important role in controlling human CMV infection 8, 9, 12. Comparable clinical trials using CTLCMV infusions have demonstrated the safety and efficacy of immunotherapy for CMV infection in HSCT 13, 15, 34. However, in line with other reports 10, 11, 12, univariate and multivariate analyses revealed no significant relationship between frequency of CTLCMV and CMV reactivation. This finding leaves open the possibility that other measurable aspects of CTLCMV response, including the contribution of key CD8+ memory T cell subsets, could be linked to control of CMV. We found that a high number of infused CTLCMV TEMRA is conversely correlated with the risk of early CMV reactivation after allogeneic HSCT with sufficient CTLCMV TEM in the allograft. Both TEM and TEMRA are effector memory cells for virus infection, and CD8+ TEMRA cells have greater cytotoxic function than CD8+ TEM cells 35, 36. Most evidence suggests that less-differentated or interleukin-2–producing polyfunctional cells are superior with respect to memory T cell function. Naïve or central memory CD8+ T cells confer superior antitumor/antivirus immunity compared with effector memory T cells 37, 38, 39, 40; however, studies have shown that CD45RO and CD45RA of memory CD8+ T cells are interchangeable 41, 42. CD45RA+ expression has been shown to be regulated upon antigenic stimulation, CD8+ as CD45RA+ T cell clones become CD45RO+, and then gradually reacquire CD45RA expression. The proliferation of all CMV tetramer-staining cells in long-term carriers suggests that these revertant memory cells can proliferate perfectly well [43], and that at least some TEMRA cells are able to upregulate telomerase activity [44]. In addition, in our HSCT setting, HSCs may promote the expansion and function of adoptively transferred antigen-specific CD8+ T cells, a process not affected by the activation state of transferred T cells [45]. A study of 18 recipients receiving T cell–depleted grafts (some with later T cell add-back) found an association between lower numbers of differentiated CMV-specific donor cells and decreased recipient CMV reactivation [46]. The difference possibly could result from characteristic changes in T cells within allografts after G-CSF stimulation (vs T cells from donor peripheral blood before HSCT) 47, 48, but more likely results from much higher T cell infusions including CTLCMV, without in vitro T cell depletion, in our HSCT protocol. This finding indicates that CTLCMV TEMRA in donor grafts play a role in the immune control of CMV reactivation after HSCT.

Regular monitoring of immune reconstitution demonstrated significantly increased circulating CTLCMV TCM, but not CTLCMV TEMRA or its other subsets, in samples obtained at day +30 and day +60 post-HSCT from recipients with CMV reactivation (on median day +31). A possible explanation for this finding is that TEMRA may serve as backup for a functional subset of CTLCMV, which responds to CMV infection, and so CTLCMV TEMRA in peripheral blood is not associated with CMV reactivation. As a matter of fact, virus-specific CTL TCM can expand adequately on virus encounter, sustain TEM and TEMRA cell populations, and control virus infection. This, it is likely that the TEMRA cells are ready to rapidly intervene on reencounter with antigen, whereas precursor cells will expand and ensure continuous replenishment of the effector cell pool [49]. Whether the reduced risk of virus infection is the result of high CTLCMV TEMRA content in donor grafts, increased circulating CTLCMV TCM after HSCT, or both is not known, however. Investigate a possible correlation between input of CTLCMV TEMRA cells and reconstitution of CTLCMV TCM cells early post-HSCT would be of interest.

Donor graft is an important source of hematopoietic cells that not only give rise to neutrophils, natural killer cells, dendritic cells, monocytes/macrophages and B cells, but also provide an important source of T cells, because de novo recovery of T cells (from hematopoietic cells) early after HSCT is inadequate. CMV-specific CD8+ T cell regeneration has been reported to depend on the size of the T cell input, but to not be a consequence of nonspecific lymphopenia-driven expansions of CD3+ and CD8+ T cell compartments 50, 51. Consistent with this and other reports 6, 7, most CTLCMV cells detected in our patients early after HSCT were from donors with some persistent recipient CTLs, illustrating the transfer and expansion of CMV-specific donor T cells. Pretransplantation, CMV-specific T cells protect recipients of T cell–depleted grafts against CMV-related complications [52], whereas infusion of unmanipulated grafts without in vitro T cell depletion using our HSCT protocol might compromise the role of recipient CTLs. Thus, the model for protection from CMV infection after HSCT should focus on the antigen-expansion or direct effect of these donor CTLCMV cells.

We found higher frequencies of CTLCMV TNaive, TCM, and TEM at day +90 and CTLCMV TEMRA at day +60 post-HSCT in recipients with higher CTLCMV TEMRA cell infusions. The association between CTLCMV TEMRA and CTLCMV TEM in donor grafts suggests that infusion of CTLCMV TEMRA cells helps accelerate immune recovery of CTLCMV and its subsets, and then helps control CMV reactivation. Furthermore, accumulating data suggest that CTLCMV TEMRA are by no means terminally differentiated, and the reverse situation may hold for memory CD8+ T cells 36, 53. Northfield et al. [17] showed that HIV-1–specific CD8+ T cells were phenotyped in the early phase of infection, and that a larger number of HIV-1–specific CD8+ TEMRA was associated with a lower future viral load set point. These authors did not continuously examine the HIV-1–specific CD8+ T cell response, however. A study of CMV infection found that CMV-specific TEMRA contributed to immunity not only through direct execution of effector functions, but also by yielding progeny in situations of viral reinfection or reactivation [44]. Questions pertaining to a similar reverse situation in CTLCMV cells of HSCT recipients remains to be answered [21]. Higher numbers of CD8+ TEMRA cells infused at transplantation could yield progeny after CMV reactivation, resulting, when accompanied by sufficient CD8+ TEM cell infusion, in better control of virus infections. This result differs from findings in most adoptive immunotherapies with mixtures of CD62L+ and CD62L- cells 54, 55, and more extensive investigations are needed to elucidate the underlying biological mechanisms.

The immune response to a complex pathogen, such as CMV, involves a large number of components, with different individuals responding to different epitopes. Although the response to pp65 may be prominent in HLA-A2– or HLA-A24–positive individuals, other viral epitopes, such as IE-1, can contribute as well, and in the context of other restriction elements may even predominate 56, 57.

One further issue of relevance for the use of defined epitopes in an adaptive immunotherapy strategy must be the role of major histocompatibility class II restricted T helper cell responses. Although not all CD8 cell responses have the same dependence on T cell help, efficient CD4 T cell responses may nonetheless be required for optimal and long-term memory responses [19]. Further studies of IE-1–specific CTLs and human CMV-specific CD4+ cells within donor grafts could help elucidate their role in CMV reactivation after HSCT. Of note, T cell responses occur following primary infection selection of T cells into memory and long-term maintenance [58]. The level of antigen stimulation may play a role in the phenotype of the reactive cells. In this respect, an inverse correlation between viral load and the frequencies of CMV-specific CTLs might be expected. Indeed, we found no significant associations in phenotype between them (data not reported), similar to previous studies 10, 11, 12. An analysis of the interplay between viral burden and size of the CMV-specific CD8+ T cell response requires simultaneous analysis of its diversity, however. Both the peak and duration of viremia can be evaluated only discontinuously in the population.

In summary, our data suggest that the CTLCMV TEMRA content of donor grafts can predict the risk of CMV reactivation after HSCT in the presence of sufficient TEM cell infusion, which is closely correlated with the immune reconstitution and differentiation of CTLCMV subsets. Protective immunity to CMV transferred during HSCT is determined by the characteristics of CTLCMV cells within donor grafts. These results have important implications for adoptive cell therapy and vaccine design for the control of CMV infection.

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Acknowledgments 

This work was supported by the Hi-Tech Research and Development Program of China (Grant 2006AA02Z4A0), the China National Funds for Distinguished Young Scientist's (Grant 30725038), and the Funds for Creative Research Groups of China (Grant IRT0702).

Authorship statement: Xiao-Hua Luo performed the research, analyzed the data, and wrote the manuscript. Xiao-Jun Huang designed the research, analyzed the data, and wrote the manuscript. Kai-Yan Liu, Lan-Ping Xu, and Dai-Hong Liu wrote and supported the clinical protocol and edited the manuscript.

Financial disclosure: The authors have no conflicts of interest to disclose.

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Supplementary Material 

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 Financial disclosure: See Acknowledgments, page 1002.

PII: S1083-8791(10)00063-7

doi:10.1016/j.bbmt.2010.02.007

Biology of Blood and Marrow Transplantation
Volume 16, Issue 7 , Pages 994-1004, July 2010

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