Human CMV infection is frequent in kidney transplant recipients (KTR). Pretransplant Ag-specific T cells and adaptive NKG2C+ NK cells associate with reduced incidence of infection in CMV+ KTR. Expansions of adaptive NKG2C+ NK cells were reported in posttransplant CMV-infected KTR. To further explore this issue, NKG2C+ NK, CD8+, and TcRγδ T cells were analyzed pretransplant and at different time points posttransplant for ≥24 mo in a cohort of CMV+ KTR (n = 112), stratified according to CMV viremia detection. In cryopreserved samples from a subgroup (n = 49), adaptive NKG2C+ NK cell markers and T cell subsets were compared after a longer follow-up (median, 56 mo), assessing the frequencies of CMV-specific T cells and viremia at the last time point. Increased proportions of NKG2C+ NK, CD8+, and TcRγδ T cells were detected along posttransplant evolution in viremia(+) KTR. However, the individual magnitude and kinetics of the NKG2C+ NK response was variable and only exceptionally detected among viremia(–) KTR, presumably reflecting subclinical viral replication events. NKG2C+ expansions were independent of KLRC2 zygosity and associated with higher viral loads at diagnosis; no relation with other clinical parameters was perceived. Increased proportions of adaptive NKG2C+ NK cells (CD57+, ILT2+, FcεRIγ) were observed after resolution of viremia long-term posttransplant, coinciding with increased CD8+ and Vδ2 γδ T cells; at that stage CMV-specific T cells were comparable to viremia(–) cases. These data suggest that adaptive NKG2C+ NK cells participate with T cells to restore CMV replication control, although their relative contribution cannot be discerned.

Cytomegalovirus establishes a prevalent and life-long infection, generally asymptomatic in healthy individuals. Different immune mechanisms contribute to control the pathogen, which enters a latency state undergoing sporadic reactivations (1, 2). CMV becomes an important threat in congenital infection and in immunocompromised patients. In kidney transplant recipients (KTR), immunosuppression favors CMV infection, which has been associated to reduced graft and patient survival (35). Antiviral prophylaxis is indicated in high-risk CMV recipients transplanted from CMV+ donors (CMV D+/R), as well as in patients undergoing intensive immunosuppression (6). CMV+ KTR at intermediate risk may experience viral reactivation or reinfection. In that case, monitoring viremia along the first months posttransplant allows selective preemptive antiviral therapy (6). Low viremia values of uncertain clinical significance may be detected by sensitive quantitative nucleic acid amplification testing (QNAT), but there is no consensus on a precise threshold for therapeutic intervention (6, 7).

In CMV+ KTR, the risk of posttransplant infection and restoration of CMV control depend on the individual fitness of the immune system (8). In this regard, pretransplant frequencies of T cells specific for IE-1 and pp65 CMV Ags have been associated with a reduced risk of posttransplant infection (912), whose resolution was reported to correlate with increased cytotoxic Vδ2 γδ T cells (13). Together with T cells, NK cells are known to contribute to immune defense against CMV infection, which promotes the adaptive differentiation and persistent expansion of a mature CD94/NKG2C+ NK cell population with a distinctive phenotypic and functional profile (1418). Common adaptive NK cell–associated features include 1) lack of the homologous CD94/NKG2A inhibitory receptor; 2) expression of CD57 and inhibitory receptors for HLA class I molecules (i.e., killer Ig-like receptors [KIRs] and Ig-like transcript 2 [ILT2; also termed LIR1 or LILRB1]); and 3) downregulation of activating receptors (i.e., NKp30, NKp46), signaling adaptors (e.g., FcεRI γ-chain), and transcription factors (e.g., prolymphocytic leukemia zinc finger transcription factor [PLZF]). This pattern of response to the viral infection is detected to a variable degree in healthy blood donors, being particularly prominent in immunocompromised individuals (15, 1921). A deletion of the KLRC2 gene–encoding NKG2C has been identified in populations of different ethnic origins (2224), and the numbers of NKG2C+ NK cells have been reported to be greater in KLRC2 homozygous compared with hemizygous CMV+ healthy individuals (24, 25). We reported that pretransplant adaptive NKG2C+ NK cells in KTR were associated with a reduced incidence of symptomatic CMV infection (26, 27), suggesting that they may contribute with T cells to contain infection progression, rather than impairing initial viral replication. Development of adaptive NKG2C+ NK cells in allogeneic hematopoietic stem cell transplantation was related with protection against CMV reactivation (28, 29). Expansions of adaptive NKG2C+ NK cells in response to posttransplant CMV infection in KTR have been described, yet information on that process is limited (3032). In the current study, we analyzed the evolution of NKG2C+ NK, CD8+, and TcRγδ T cells for ≥24 mo in a cohort of CMV+ KTR (n = 112), stratified according to the incidence of posttransplant CMV viremia. In a subgroup (n = 49), additional adaptive NKG2C+ NK cell markers, T cell subsets, frequencies of CMV-specific T cells, and viremia were assessed at the end of a longer follow-up period (median, 56 mo).

Patients undergoing kidney transplantation at Hospital del Mar (Barcelona) between February 2013 and June 2017 were enrolled. KTR at intermediate risk of CMV infection (D+R+ or DR+) who were followed for at least 24 mo and received the same maintenance immunosuppressive regimen (tacrolimus, mycophenolic acid, and steroids) with anti-CD25 mAb induction were included (n = 112) (26). Cases at low risk of CMV infection (DR) and those at high risk receiving antiviral prophylaxis (i.e., CMV D+R) were excluded. Patients were transplanted according to negative complement-dependent cytotoxicity cross-match with donor lymphocytes, as previously described (33). PBMC cryopreservation and immunophenotyping was routinely performed prior to transplantation and at different time points afterward (∼3, 6, 12, 24, and 36 mo). In a subgroup (n = 49), an additional analysis was carried out, coinciding with a later clinical visit (median, 56 mo; interquartile range [IQR], 46–62) assessing adaptive NK cell markers, T cell subpopulations, CMV-specific T cell frequencies, and viremia. The study was conducted following the Declaration of Helsinki guidelines and approved by Parc de Salut Mar Ethical Research Board (2018/7873I). All patients signed written informed consent for the use of peripheral blood samples for research purposes.

CMV DNAemia was systematically monitored following transplantation by standardized diagnostic QNAT (COBAS AmpliPrep, Cobas TaqMan; Roche Diagnostics) every 2 wk for the first 3 mo posttransplant and whenever infection was clinically suspected. Viremia was defined based on QNAT detection in plasma samples of viral loads above the lower limit of quantification (LLOQ) (137 IU/ml) or by histopathological examination. Viremia(+) patients were stratified as asymptomatic or symptomatic according to clinical guidelines (6). The latter category included cases with viral syndrome or with histopathological evidence of invasive disease. Symptomatic cases and those asymptomatic with DNAemia > 500 copies per milliliter (455 IU/ml) received antiviral therapy. In the subgroup studied in more detail (n = 49), DNAemia was surveyed at the end of follow-up (median, 56 mo posttransplant), employing a different technique (Abbott Diagnostics, Lake Forest, IL) with a reduced LLOQ (31.2 IU/ml), which was adopted by the clinical diagnostics laboratory during the study (2018).

As previously described (27), blood samples were obtained by venous puncture in EDTA and directly analyzed by flow cytometry; in addition, PBMC were separated by Ficoll-Hypaque density gradient centrifugation and cryopreserved. For flow cytometry analysis, samples were pretreated with human aggregated IgG (100 µg/ml) to block FcR. In fresh samples, direct analysis of NK, CD8, CD4, and TCRγδ T cells was systematically performed using BD Multitest CD3/CD8/CD45/CD4 (BioLegend), anti–TCRγδ-PE (11F12; BioLegend), and anti–CD56-FITC (NCAM16.2; BD Biosciences). NKG2C expression was detected by indirect immunofluorescence using anti-NKG2C (MAB1381; R&D Systems) and PE-conjugated F(ab')2 goat anti-mouse secondary Ab. Data were acquired on an FACS Canto II Cytometer (BD Biosciences).

Extensive immunophenotyping was performed in thawed PBMC samples as described (26) using the following monoclonal Abs: anti–CD3-allophycocyanin-Cy7(OKT3; BioLegend), anti–CD45-Alexa Fluor 700 (2D1; eBioscience), anti–CD56-allophycocyanin-Cy7(NCAM; BioLegend), anti–CD3-PerCP(SK7; BD Biosciences), anti-TCR pan γ/δ-PE/Dazzle594 (B1; BioLegend), anti-TCR Vδ2-FITC (IMMU389; Beckman-Coulter), anti–NKG2C-PE (FAB138P; R&D systems), anti–NKG2A-Pacific Blue (Z199; provided by Dr. A. Moretta), anti–ILT2-PeCy7 (GHI/75; BioLegend), anti–CD57-FITC (HCCD57; BioLegend), anti–CD57-allophycocyanin (HCD57; BioLegend), anti–CD56-allophycocyanin (CMSSB; eBioscience), anti–CD4-allophycocyanin (RTA-T4; BD Pharmingen), anti–CD8-BV510 (RPA-T8; BD Biosciences), and anti–CD56-BV510 (NCAM16.2; BD Bioscience). For intracellular staining, cells were fixed and permeabilized (FIX & PERM Cell Fixation & Cell Permeabilization Kit; Invitrogen), according to manufacturer instructions, followed by intracellular staining with anti-FcεRIγ–FITC (polyclonal; Merck MilliporeSigma) and anti–PLZF-PECF594 (R17-809; BD Biosciences). Data were acquired on an LSR Fortessa Flow Cytometer (BD Biosciences) and analyzed using FlowJo software (10.0.7; TreeStar). Gating strategy is shown in Supplemental Fig. 1.

Multidimensional flow cytometry analysis using t-distributed stochastic neighbor embedding (t-SNE) was implemented in manually gated NK and T cells as described (17, 26, 34). KLRC2 (NKG2C) gene deletion was assessed on genomic DNA samples as described (23).

PBMCs were incubated overnight at 37°C in polypropylene tubes with complete medium (RPMI 1640 supplemented with 10% heat-inactivated FBS, 100 U/ml penicillin, 100 µg/ml streptomycin, and 1 mM sodium pyruvate) and placed at 1 × 106 cells per well in 96-well plates. T cell activation was assessed by flow cytometry detection of cytokine production, as described (26) Briefly, samples were incubated with RPMI alone, anti-CD3 mAb or 1 µg/ml of peptide libraries from CMV pp65 or IE-1 Ags (Peptivator; Miltenyi Biotec) for 6 h. During the last 5 h, brefeldin A (Sigma-Aldrich) and anti-CD49d (BD Biosciences) were added to a final concentration of 10 μg/ml and 1 g/ml, respectively. Subsequently, cells were stained with fluorochrome-conjugated Abs and a minimum of 105 T cells were analyzed on an LSR Fortessa flow Cytometer (26).

Statistical analysis was performed in R version 3.5.1. Categorical variables are expressed as percentages and continuous variables as mean and SD or median and IQR. Univariate analysis was performed by χ2 test, Spearman correlation, Wilcoxon, or Fisher exact test, as appropriate. Two-sided p values <0.05 were considered significant.

Generalized estimating equations population-averaged model was used to analyze in the whole cohort the evolution of NK and T cell populations along the follow-up time points to compare the evolution of study groups an interaction term group x time was added.

An immunophenotypic analysis was sequentially carried out pretransplant and at different time points posttransplant in PBMCs from a cohort of CMV+ KTR (n = 112) who did not receive antiviral prophylaxis and were followed up for at least 24 mo. Demographic and clinical information is summarized in Table I. NKG2C+ NK cells were assessed in patients stratified according to the incidence of CMV viremia as defined in the Materials and Methods. Given the scope of the study, six cases with undetectable NKG2C+ NK cells and a confirmed homozygous KLRC2 deletion were excluded from subsequent analyses. As compared with pretransplant levels, the proportions of NKG2C+ NK cells were significantly increased in viremia(+) KTR at 12 and 24–36 mo posttransplant (Fig. 1A), with differences being more evident in symptomatic cases (Fig. 1B). However, a substantial individual variability in the evolution of the NKG2C+ NK cells was noticed in viremia(+) KTR (Fig. 1C) displaying early/delayed increases, steady levels, or late decrements; representative profiles are illustrated in (Fig. 1D. By contrast, NKG2C+ NK cell increments were only exceptionally observed among KTR categorized as viremia(–) (5 out of 68).

FIGURE 1.

Dynamics of NKG2C+ NK cells in KTR. (A) Frequencies of NKG2C+ NK cells were assessed pretransplant and at 12 and 24–36 mo posttransplant in PBMC from a cohort of CMV+ KTR (n = 106) stratified according to the incidence of posttransplant CMV viremia. (B) NKG2C+ NK cells in viremia(+) KTR (n = 41) stratified as symptomatic or asymptomatic. (C) Individual variations between NKG2C+ NK cell frequencies pretransplant and at 24 mo posttransplant in KTR stratified according to viremia detection. (D) Line graphs showing representative evolution patterns of the frequencies of NKG2C+ NK cells in viremia(+) KTR. (E) Frequencies of NKG2C+ NK cells pretransplant and 24 mo posttransplant in KTR with >30% NKG2C+ NK cells pretransplant (n = 26); dashed lines correspond to viremia(+) cases. Graphs include median and IQR. Paired (A, B, and E) and unpaired Wilcoxon (C). *p ≤ 0.05.

FIGURE 1.

Dynamics of NKG2C+ NK cells in KTR. (A) Frequencies of NKG2C+ NK cells were assessed pretransplant and at 12 and 24–36 mo posttransplant in PBMC from a cohort of CMV+ KTR (n = 106) stratified according to the incidence of posttransplant CMV viremia. (B) NKG2C+ NK cells in viremia(+) KTR (n = 41) stratified as symptomatic or asymptomatic. (C) Individual variations between NKG2C+ NK cell frequencies pretransplant and at 24 mo posttransplant in KTR stratified according to viremia detection. (D) Line graphs showing representative evolution patterns of the frequencies of NKG2C+ NK cells in viremia(+) KTR. (E) Frequencies of NKG2C+ NK cells pretransplant and 24 mo posttransplant in KTR with >30% NKG2C+ NK cells pretransplant (n = 26); dashed lines correspond to viremia(+) cases. Graphs include median and IQR. Paired (A, B, and E) and unpaired Wilcoxon (C). *p ≤ 0.05.

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Table I.

Characteristics of the studied KTR cohort

All Cases (n = 112)No CMV Viremia (n = 71)CMV Viremia (n = 41)p Value
Recipient age (y), mean (SD) 55.3 (13) 52.5 (12.4) 60.1 (12.7) 0.003 
Female recipient, n (%) 40 (35.7) 23 (32.4) 17 (41.5) 0.447 
Donor age (y), mean (SD) 57.5 (14.4) 54.2 (14.3) 62.9 (12.9) 0.004 
Female donor, n (%) 57 (50.9) 35 (49.3) 22 (53.7) 0.860 
Live donor, n (%) 23 (20.5) 19 (26.8) 4 (9.8) 0.050 
Retransplant, n (%) 9 (8) 8 (11.3) 1 (2.4) 0.151 
Peak PRA > 5%, n (%) 16 (14.3) 12 (16.7) 4 (9.8) 0.262 
Pretransplant PRA > 5%, n (%) 9 (8) 5 (7) 4 (9.8) 0.779 
Pretransplant DSA, n (%) 2 (1.8) 2 (2.8) 0.532 
Delayed graft function, n (%) 25 (22.3) 15 (21.1) 10 (24.4) 0.870 
Biopsy-proven acute rejection (treated), n (%) 13 (11.6) 7 (9.9) 6 (14.6) 0.543 
Biopsy-proven acute rejection, n (%) 16 (14.3) 10 (14.1) 6 (14.6) 
Pre-KT CMV serostatus, n (%)    0.128 
 D+/R+ 99 (88.4) 60 (84.5) 39 (95.1)  
 D/R+ 13 (11.6) 11 (15.5) 2 (4.9)  
Symptomatic CMV, n (%)   25 (61)  
CMV PCR at diagnosis (IU/ml), median (IQR)   430 (197–736)  
Peak CMV PCR (IU/ml), median (IQR)   576 (258–1789)  
CMV infection time after KT (d), median (IQR)   56 (40–72)  
Time from CMV infection diagnosis to first negative PCR (wk), median (IQR)   3 (2.6–4.7)  
 Graft loss, n (%) 28 (25) 16 (22.5) 12 (29.3) 0.499 
 Death-censored graft loss, n (%) 8 (7.1) 3 (4.2) 5 (12.2) 0.139 
Renal functiona     
 Serum creatinine, mean (SD) 1.7 (0.9) 1.6 (0.9) 1.9 (0.8) 0.009 
 Estimated GFR, mean (SD) 50.7 (26) 57 (27) 38 (18) 0.002 
 Urine protein/creatinine ratio, median (IQR) 133.2 (66.4–324) 122.5 (66–247) 241.6 (76.8–507.5) 0.230 
All Cases (n = 112)No CMV Viremia (n = 71)CMV Viremia (n = 41)p Value
Recipient age (y), mean (SD) 55.3 (13) 52.5 (12.4) 60.1 (12.7) 0.003 
Female recipient, n (%) 40 (35.7) 23 (32.4) 17 (41.5) 0.447 
Donor age (y), mean (SD) 57.5 (14.4) 54.2 (14.3) 62.9 (12.9) 0.004 
Female donor, n (%) 57 (50.9) 35 (49.3) 22 (53.7) 0.860 
Live donor, n (%) 23 (20.5) 19 (26.8) 4 (9.8) 0.050 
Retransplant, n (%) 9 (8) 8 (11.3) 1 (2.4) 0.151 
Peak PRA > 5%, n (%) 16 (14.3) 12 (16.7) 4 (9.8) 0.262 
Pretransplant PRA > 5%, n (%) 9 (8) 5 (7) 4 (9.8) 0.779 
Pretransplant DSA, n (%) 2 (1.8) 2 (2.8) 0.532 
Delayed graft function, n (%) 25 (22.3) 15 (21.1) 10 (24.4) 0.870 
Biopsy-proven acute rejection (treated), n (%) 13 (11.6) 7 (9.9) 6 (14.6) 0.543 
Biopsy-proven acute rejection, n (%) 16 (14.3) 10 (14.1) 6 (14.6) 
Pre-KT CMV serostatus, n (%)    0.128 
 D+/R+ 99 (88.4) 60 (84.5) 39 (95.1)  
 D/R+ 13 (11.6) 11 (15.5) 2 (4.9)  
Symptomatic CMV, n (%)   25 (61)  
CMV PCR at diagnosis (IU/ml), median (IQR)   430 (197–736)  
Peak CMV PCR (IU/ml), median (IQR)   576 (258–1789)  
CMV infection time after KT (d), median (IQR)   56 (40–72)  
Time from CMV infection diagnosis to first negative PCR (wk), median (IQR)   3 (2.6–4.7)  
 Graft loss, n (%) 28 (25) 16 (22.5) 12 (29.3) 0.499 
 Death-censored graft loss, n (%) 8 (7.1) 3 (4.2) 5 (12.2) 0.139 
Renal functiona     
 Serum creatinine, mean (SD) 1.7 (0.9) 1.6 (0.9) 1.9 (0.8) 0.009 
 Estimated GFR, mean (SD) 50.7 (26) 57 (27) 38 (18) 0.002 
 Urine protein/creatinine ratio, median (IQR) 133.2 (66.4–324) 122.5 (66–247) 241.6 (76.8–507.5) 0.230 
a

Last available data at follow-up end.

DSA, donor‐specific Ab; GFR, glomerular filtration rate; KT, kidney transplantation; PRA, panel reactive Ab (by complement‐dependent cytotoxicity).

As pretransplant NKG2C+ NK cells have been associated with a reduced incidence of symptomatic infection (26, 27), KTR displaying high pretransplant proportions of this subset (>30%, n = 26), which included three viremia(+) cases, were separately analyzed (Fig. 1E). An overall reduction of NKG2C+ NK cells was observed in viremia(–) KTR (n = 23; solid lines). Yet, steady levels or minimal variations of NKG2C+ NK cells as well as increased or decreased proportions were detected. Of note, two of the three viremia(+) cases in this group (dashed lines) displayed marked late decrements of NKG2C+ NK cells (Fig. 1E).

The putative influence of KLRC2 copy number on the magnitude of posttransplant NKG2C+ NK cell expansions was considered. In line with previous observations (27), greater proportions of NKG2C+ NK cells in KLRC2wt/wt as compared with KLRC2wt/del KTR were detected pretransplant, but no differences were observed at 24 mo posttransplant. Moreover, variations of NKG2C+ NK cells in viremia(+) cases were comparable in KLRC2 homozygous and hemizygous individuals (Supplemental Fig. 2).

To explore the relation of the adaptive NK cell response with clinical variables, viremia(+) KTR were stratified in two groups, according to the relative magnitude of NKG2C+ NK cell expansions reached at 24 mo (Table II). The “expanders” category included cases with increased posttransplant proportions of NKG2C+ NK cells at least doubling pretransplant levels and with an Δ% NKG2C+ ≥ 10% of total NK cells. According to these criteria, the Δ% NKG2C+ NK cells median (IQR) was 25% (1530) for expanders and –1% (–6% to 3%) for nonexpanders. No significant differences between both groups were detected for most clinical parameters, except for significantly greater viremia detected at diagnosis in expanders (Table II). This observation suggested an association of viral load with the magnitude of the NKG2C+ NK cell response, although no significant correlations were detected by regression analysis between the Δ% NKG2C+ NK cells and CMV viremia peak (r = 0.15, p = 0.38) or at diagnosis (r = 0.3, p = 0.076).

Table II.

Characteristics of CMV viremia(+) KTR according to NKG2C+ NK expansion

CMV Viremia (n = 38)NKG2C+ Nonexpander (n = 25)NKG2C+ Expandera (n = 13)p Value
Recipient age (y), mean (SD) 59.5 (13) 61.2 (14) 56.2 (11) 0.212 
Female recipient, n (%) 15 (39.5) 9 (36) 6 (46) 0.728 
Live donor, n (%) 4 (10.5) 4 (16) 0.278 
Retransplantation, n (%) 
Pretransplant DSA, n (%)    
Delayed graft function, n (%) 10 (26) 6 (24) 4 (31) 0.709 
Biopsy-proven acute rejection (treated), n (%) 6 (16) 5 (20) 1 (8) 0.642 
Pre-KT CMV serostatus, n (%)    
 D+/R+ 36 (95) 24 (96) 12 (92)  
 D/R+ 2 (5.3) 1 (4) 1 (8)  
Symptomatic CMV, n (%) 23 (60.5) 15 (60) 8 (61.5) 
CMV PCR at diagnosis (IU/ml), median (IQR) 471 (216–759) 286 (209–550) 1000 (471–8946) 0.008 
Peak CMV PCR (IU/ml), median (IQR) 597 (279–1829) 478 (266–1094) 1024 (494–35,613) 0.050 
CMV infection time after KT (d), median (IQR) 53.5 (40–71) 57 (39–79) 50 (45–66) 0.914 
Time from CMV infection diagnosis to first negative PCR (wk), median (IQR) 3.1 (2.6–4.7) 3 (2.6–4.7) 3.3 (2.1–4.7) 0.926 
Graft loss, n (%) 11 (29) 8 (32) 3 (23) 0.714 
Death-censored graft loss, n (%) 5 (13) 4 (16) 1 (8) 0.643 
Renal functionb 
 Serum creatinine, mean (SD) 2 (0.8) 2 (0.8) 1.9 (0.7) 0.782 
 Estimated GFR, mean (SD) 37.9 (19) 36 (19) 40 (20) 0.620 
 Urine protein/creatinine ratio, median (IQR) 241 (80–510) 263 (90–437) 126 (56–705) 0.587 
CMV Viremia (n = 38)NKG2C+ Nonexpander (n = 25)NKG2C+ Expandera (n = 13)p Value
Recipient age (y), mean (SD) 59.5 (13) 61.2 (14) 56.2 (11) 0.212 
Female recipient, n (%) 15 (39.5) 9 (36) 6 (46) 0.728 
Live donor, n (%) 4 (10.5) 4 (16) 0.278 
Retransplantation, n (%) 
Pretransplant DSA, n (%)    
Delayed graft function, n (%) 10 (26) 6 (24) 4 (31) 0.709 
Biopsy-proven acute rejection (treated), n (%) 6 (16) 5 (20) 1 (8) 0.642 
Pre-KT CMV serostatus, n (%)    
 D+/R+ 36 (95) 24 (96) 12 (92)  
 D/R+ 2 (5.3) 1 (4) 1 (8)  
Symptomatic CMV, n (%) 23 (60.5) 15 (60) 8 (61.5) 
CMV PCR at diagnosis (IU/ml), median (IQR) 471 (216–759) 286 (209–550) 1000 (471–8946) 0.008 
Peak CMV PCR (IU/ml), median (IQR) 597 (279–1829) 478 (266–1094) 1024 (494–35,613) 0.050 
CMV infection time after KT (d), median (IQR) 53.5 (40–71) 57 (39–79) 50 (45–66) 0.914 
Time from CMV infection diagnosis to first negative PCR (wk), median (IQR) 3.1 (2.6–4.7) 3 (2.6–4.7) 3.3 (2.1–4.7) 0.926 
Graft loss, n (%) 11 (29) 8 (32) 3 (23) 0.714 
Death-censored graft loss, n (%) 5 (13) 4 (16) 1 (8) 0.643 
Renal functionb 
 Serum creatinine, mean (SD) 2 (0.8) 2 (0.8) 1.9 (0.7) 0.782 
 Estimated GFR, mean (SD) 37.9 (19) 36 (19) 40 (20) 0.620 
 Urine protein/creatinine ratio, median (IQR) 241 (80–510) 263 (90–437) 126 (56–705) 0.587 
a

The expanders category included cases with increased posttransplant proportions of NKG2C+ NK cells at least doubling pretransplant levels and with a Δ% NKG2C+ ≥ 10% of total NK cells. According to these criteria the Δ% NKG2C+ NK cells median (IQR) was 25% (15–30%) for expanders and –1% (–6 to 3%) for nonexpanders.

b

Last available data at follow-up end.

DSA, donor‐specific Ab; GFR, glomerular filtration rate; KT, kidney transplantation.

A more detailed analysis was conducted in available cryopreserved samples from a KTR subgroup, including viremia(+) (n = 23) and viremia(–) (n = 26) cases, who were further studied at a late regular clinical visit (median, 56 mo; IQR, 46–62) (Supplemental Tables I, II). As shown in (Fig. 2A, the evolution patterns of NKG2C+ NK cells at ≥36 mo in KTR stratified according to the incidence of viremia are consistent with observations in the whole cohort. A subset of KTR displaying marked increases of NKG2C+ NK cells was clearly differentiated from cases showing minimal changes, steady, or decreased proportions.

FIGURE 2.

Expression of adaptive NKG2C+ NK cell differentiation markers in CMV viremia(+) KTR. (A) Frequencies of NKG2C+ NK cells detected pretransplant and >36 mo (median, 56; IQR, 46–62 mo) posttransplant in KTR (n = 49) stratified according to posttransplant viremia detection. Paired Wilcoxon. (B) Flow cytometry plots comparing the expression of adaptive NKG2C+ NK cell markers analyzed pretransplant and at the end of the long-term follow-up in infected KTR. Cases are representative of two different profiles, displayed in the accompanying line graphs.

FIGURE 2.

Expression of adaptive NKG2C+ NK cell differentiation markers in CMV viremia(+) KTR. (A) Frequencies of NKG2C+ NK cells detected pretransplant and >36 mo (median, 56; IQR, 46–62 mo) posttransplant in KTR (n = 49) stratified according to posttransplant viremia detection. Paired Wilcoxon. (B) Flow cytometry plots comparing the expression of adaptive NKG2C+ NK cell markers analyzed pretransplant and at the end of the long-term follow-up in infected KTR. Cases are representative of two different profiles, displayed in the accompanying line graphs.

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To further evaluate in this subgroup the impact of posttransplant infection on the development of adaptive NKG2C+ NK cells, expression of differentiation markers (i.e., lack of NKG2A and CD57 and ILT2 expression and FcεRγ downregulation) was assessed in PBMC samples obtained pretransplant and at the last follow-up time point, analyzing also in the latter PLZF downregulation. (Fig. 2B displays flow cytometry analyses of two representative viremia(+) KTR. The evolution of adaptive NKG2C+ NK cell markers was compared according to viremia detection in KTR. Given the scope of the analysis, cases already displaying high pretransplant proportions of adaptive NK cells (>30%) (Fig. 1E) that would mask the changes were excluded. As shown in (Fig. 3A, proportions of NKG2C+ NK cells with adaptive phenotypic features (i.e., NKG2A, CD57+, FcεRIγ, and ILT2+) were significantly increased posttransplant in viremia(+) KTR. Conversely, their pretransplant levels were greater in viremia(–) cases, consistent with the reported association of this subset with a reduced risk of CMV infection (26). Altogether, these results revealed a marked individual variability in the long-term evolution of canonical NKG2C+ adaptive NK cells following posttransplant CMV infection. By tSNE analysis, three clusters of NKG2C+ NKG2A NK cells, prominent in viremia(+) cases, could be discriminated according to CD57, ILT2, FcεRγ, and PLZF expression, likely representing distinct adaptive NK cell differentiation stages (Fig. 3B). By contrast, NKG2C NKG2A+/− subsets expressing PLZF and FcεRγ as well as different levels of CD57 and ILT2 predominated in viremia(–) KTR.

FIGURE 3.

Effect of CMV viremia on the long-term distribution of adaptive NKG2C+ NK cell markers. (A) Frequencies of NKG2C+ NK cells displaying adaptive NK cell–associated markers, pretransplant and at the end of the follow-up, in KTR with available pretransplant information (n = 31), segregated according to posttransplant CMV viremia. Graphs include median and IQR. (B) Multidimensional Barnes-Hut t-SNE analysis of six parameters was performed on manually gated NK cells at the end of the follow-up in PBMC samples from KTR (n = 34). Single-parameter plots show protein expression levels on the t-SNE field. Kernel density plots show events in the t-SNE field compiled separately for samples from patients stratified according to detection of posttransplant CMV viremia. Cases with high baseline levels (>30%) of NKG2C+ NK cells were excluded from both analyses. Paired Wilcoxon for pretransplant and posttransplant comparisons and unpaired Wilcoxon for comparison between infected and noninfected patients. *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001.

FIGURE 3.

Effect of CMV viremia on the long-term distribution of adaptive NKG2C+ NK cell markers. (A) Frequencies of NKG2C+ NK cells displaying adaptive NK cell–associated markers, pretransplant and at the end of the follow-up, in KTR with available pretransplant information (n = 31), segregated according to posttransplant CMV viremia. Graphs include median and IQR. (B) Multidimensional Barnes-Hut t-SNE analysis of six parameters was performed on manually gated NK cells at the end of the follow-up in PBMC samples from KTR (n = 34). Single-parameter plots show protein expression levels on the t-SNE field. Kernel density plots show events in the t-SNE field compiled separately for samples from patients stratified according to detection of posttransplant CMV viremia. Cases with high baseline levels (>30%) of NKG2C+ NK cells were excluded from both analyses. Paired Wilcoxon for pretransplant and posttransplant comparisons and unpaired Wilcoxon for comparison between infected and noninfected patients. *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001.

Close modal

Although no clinical evidence of CMV infection was recorded at the end of the long-term follow-up, viremia was assessed in parallel to the last immunophenotypic analysis. In this survey, out of the 49 KTR studied, low DNAemia levels were detected in cases initially identified as viremia(+) (Supplemental Table II; 982, 1034, 1224) and viremia(–) (Supplemental Table II; 1105), all displaying scarce proportions of NKG2C+ cells, as well as in a patient with early increased NKG2C+ NK cells (Supplemental Table II; 1106). Although limited to a single point analysis, these data do not support an association between late viremia and adaptive NKG2C+ NK cell expansions, and suggest a putative contribution of the latter to long-term CMV control.

Vδ2 γδ T cells have been associated with control of posttransplant CMV infection in KTR (13), and expansions of CMV-specific CD8+ T cells were reported even in the absence of CMV disease (35). Based on immunophenotyping data in fresh samples from the whole cohort, an integrated analysis of posttransplant evolution of NKG2C+ NK, CD8+, and total TcRγδ T cells was carried out, comparing KTR stratified according to viremia (Fig. 4A–C). Reflecting the variability of the response described above, the changes of NKG2C+ NK cells were not significant when evaluated separately within each group [viremia(+) and viremia(–)], but these differed significantly from each other. In contrast, significant differences were also observed for the evolution of CD8+ and total TcRγδ T cells (Fig. 4B–C), which were increased in viremia(+) KTR; to a lesser extent, changes in CD8+ T cells were also noticed in viremia(–) cases. The evolution of CD8+ T cells was comparable regardless of the magnitude of NKG2C+ NK cell expansions as defined before (Fig. 4D); by contrast, TcRγδ T cells significantly increased in NKG2C expanders (Fig. 4E), suggesting a relation between the response of both populations to CMV infection.

FIGURE 4.

Comparative posttransplant evolution of NKG2C+ NK, CD8+, and TcRγδ T cells in relation with CMV viremia. Generalized estimating equations population–averaged model was used to analyze in the whole cohort the evolution along the follow-up time points of NKG2C+ NK (A), CD8+ (B), and total TcRγδ T cells (C). To compare the evolution study groups the stratifying variable (i.e., viremia detection) was analyzed as an interaction term. A similar analysis was carried out to compare the evolution of CD8+ (D) and total TcRγδ T cells (E) in viremia(+) KTR stratified according to the magnitude of NKG2C+ NK cell expansion (see results and Table II footnote). The p values marked for each curve correspond to the comparisons between data obtained at baseline and the different time points. Statistical comparisons between study groups are separately indicated.

FIGURE 4.

Comparative posttransplant evolution of NKG2C+ NK, CD8+, and TcRγδ T cells in relation with CMV viremia. Generalized estimating equations population–averaged model was used to analyze in the whole cohort the evolution along the follow-up time points of NKG2C+ NK (A), CD8+ (B), and total TcRγδ T cells (C). To compare the evolution study groups the stratifying variable (i.e., viremia detection) was analyzed as an interaction term. A similar analysis was carried out to compare the evolution of CD8+ (D) and total TcRγδ T cells (E) in viremia(+) KTR stratified according to the magnitude of NKG2C+ NK cell expansion (see results and Table II footnote). The p values marked for each curve correspond to the comparisons between data obtained at baseline and the different time points. Statistical comparisons between study groups are separately indicated.

Close modal

TcRαβ (CD4+, CD8+), TcRγδ (Vδ2, Vδ2+), and a minor NKG2C+ T cell subset reported in CMV+ healthy subjects (14) were also analyzed in the subgroup of 49 KTR with extended follow-up described above. Pre-/posttransplant phenotypes were compared according to the incidence of CMV viremia. As shown in (Fig. 5A, significantly increased CD8+ and reduced CD4+ T cell subsets, as well as raised proportions of Vδ2 γδ T cells and a minor NKG2C+ Vδ2 subset were detected in viremia(+) cases long-term after transplant. By contrast, no significant differences of Vδ2+ γδ or total NKG2C+ T cells were observed. Of note, differences were also noticed comparing the levels of CD4+ and Vδ2 γδ T cells in viremia(–) cases.

FIGURE 5.

Effects of CMV viremia on the long-term distribution of T cell subsets. (A) The frequencies of CD4+, CD8+, NKG2C+, and TCRγδ (Vδ2+ and Vδ2−) T cell subsets were assessed by flow cytometry, pretransplant, and at the end of the follow-up (median, 56 mo), in KTR with available pretransplant information (n = 31), stratified according to CMV viremia detection. Graphs include median and IQR. (B) Multidimensional Barnes-Hut t-SNE analysis of six parameters was performed on manually gated T cells at the end of the follow-up in PBMC samples from KTR (n = 34), as described in (Fig. 3. Paired Wilcoxon for pretransplant and posttransplant comparisons and unpaired Wilcoxon for comparison of patients according to viremia detection. *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001.

FIGURE 5.

Effects of CMV viremia on the long-term distribution of T cell subsets. (A) The frequencies of CD4+, CD8+, NKG2C+, and TCRγδ (Vδ2+ and Vδ2−) T cell subsets were assessed by flow cytometry, pretransplant, and at the end of the follow-up (median, 56 mo), in KTR with available pretransplant information (n = 31), stratified according to CMV viremia detection. Graphs include median and IQR. (B) Multidimensional Barnes-Hut t-SNE analysis of six parameters was performed on manually gated T cells at the end of the follow-up in PBMC samples from KTR (n = 34), as described in (Fig. 3. Paired Wilcoxon for pretransplant and posttransplant comparisons and unpaired Wilcoxon for comparison of patients according to viremia detection. *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001.

Close modal

tSNE analysis graphically discriminated differences between cases, stratified according to viremia, in the distributions of discrete clusters of CD4+, CD8+, and Vδ2 γδ T cell populations, as well as of a minor NKG2C+ Vδ2 γδ T cell subset (Fig. 5B).

The specific T cell response to pp65 and IE-1 immunodominant CMV Ags was also assessed at the end of the follow-up. As shown in Supplemental Fig. 3, no differences in the frequencies of pp65- and IE1-specific T cells were perceived between KTR stratified according to viremia. Altogether, these data illustrated the complex influence exerted by CMV infection on the NK and T cell compartments long-term after transplantation, which likely cooperate in controlling CMV, rendering it difficult to ascertain the individual contribution of adaptive NKG2C+ NK cells in this process.

Control of CMV replication is altered in seropositive KTR, often leading to viral reactivation and/or reinfection. Factors related with the transplantation procedure (e.g., cold ischemia) may trigger viral replication, whereas immunosuppression promotes infection progression (36). The risk of posttransplant viremia, its clinical impact, and control ultimately depend on the individual fitness of the immune system. In this regard, frequencies of CMV pp65- and IE-1–specific T cells inversely correlate with incidence of posttransplant infection, consistently with their central role in keeping the pathogen at bay (912). Recently, pretransplant adaptive NKG2C+ NK cells were also associated with a reduced incidence of symptomatic infection in KTR (26, 27), suggesting that they may contribute with T cells to contain infection progression, rather than hampering initial viral replication. Development of adaptive NKG2C+ NK cells in hematopoietic stem cell transplantation was related with protection against CMV reactivation (28, 29). Expansions of adaptive NKG2C+ NK cells to posttransplant CMV infection in KTR have been reported but information on this process is limited (3032).

In this study, adaptive NKG2C+ NK cells were analyzed, at different times posttransplant, in a cohort of CMV+ KTR undergoing a homogeneous immunosuppressive regimen without antiviral prophylaxis. Adaptive NKG2C+ NK cell expansions were detected in viremia(+) KTR, yet with marked individual differences in their magnitude and kinetics at the individual level. In some cases, NKG2C+ NK cells increased early and sharply after viremia detection, persisting elevated along the follow-up, whereas increments appeared delayed and progressive in others. These profiles may reflect that NKG2C+ NK cells compensate an inefficient T cell–mediated control of CMV reactivation/reinfection, which might take place during the first months after transplant, when viremia was systematically monitored, but also at later times.

Conversely, other viremia(+) KTR showed minor changes or steady levels of adaptive NKG2C+ NK cells, comparable to the profile commonly observed in viremia(–) patients, presumably indicating a prompt and stable control of CMV replication. Of note, NKG2C+ NK cell expansions were exceptionally detected in few cases categorized as viremia(–), likely reflecting overlooked subclinical CMV replication events after interruption of monitoring.

As pretransplant levels of adaptive NKG2C+ NK cells were associated with reduced incidence of symptomatic infection (26) it is plausible that posttransplant development of an adaptive NK cell response may also contribute to restore CMV control in KTR. Consistent with this hypothesis, viremia was undetectable at the late follow-up time point in most KTR, including cases with adaptive NKG2C+ NK cell expansions. Moreover, in that survey four out of five cases with late subclinical viremia displayed relatively low proportions (<20%) of NKG2C+ NK cells. Although these data suggest that NKG2C+ NK cell expansions might contribute to maintain CMV control, concomitant increases of CD8+ and TcRγδ T cell subsets were also detected along posttransplant evolution in viremia(+) KTR, who displayed late frequencies of IE-1– and pp65-pecific T cells comparable to viremia(–) patients. Effects of posttransplant CMV infection in the T cell compartment have been previously reported in KTR (37). Thus, the complex imprint of CMV infection on the immune system did not allow to discern the relative roles played by the different lymphocyte subsets, which likely cooperate in controlling the pathogen.

The adaptive NKG2C+ NK cell response to posttransplant viral infection in CMV+ KTR may result from expansion of a preexisting pool of this subset, and/or from de novo differentiation. Although the sensitivity of adaptive NK cell development to different immunosuppressive drugs has not been specifically addressed (38), observations in our cohort indicate that the process is, at least partially, resistant to the common immunosuppressive regimen. Moreover, NKG2C+ NK cell expansions detected in a group of high-risk CMV seronegative KTR suffering posttransplant infection (M. Ataya and M. López-Botet, unpublished observations) indicate that their de novo differentiation may at least partially overcome the effects of standard immunosuppression.

CMV infection appears the primary determinant for triggering an adaptive NKG2C+ NK cell response, and thus its magnitude may be indirectly modulated by factors related with pathogen control. As reported (39), preemptive antiviral therapy may reduce antigenic stimulation of the immune system, thus potentially blunting as well adaptive NKG2C+ NK cell development. In the same line, control of donor-derived CMV reinfection may be less efficient than that of viral reactivation (40), potentially favoring adaptive NK cell development. In contrast, the magnitude of the adaptive NK cell response may be also determined by host/viral variables directly related with the underlying mechanisms, which remain incompletely defined. In this regard, we found no association with KLRC2 copy number, reported to influence the numbers of circulating adaptive NKG2C+ NK cells in other settings (24, 25, 27). Two additional variables not explored in this study deserve attention. Because activation through CD16 induces proliferation and Ab-dependent cell–mediated cytotoxicity of adaptive NKG2C+ NK cells (18, 4143), their numbers might be influenced by the concentration of CMV-specific IgG, as well as by polymorphisms conditioning CD16-IgG affinity (44). Moreover, the adaptive NKG2C+ NK cell response has been proposed to be triggered by CD94/NKG2C recognition of HLA-E bound to peptides from the CMV UL40 molecule (45) that mimic endogenous HLA class I–derived leader sequence nonamers, whose polymorphism influences CD94/NKG2C affinity for HLA-E (46). Prospective studies on viremia samples are required to assess whether variability of the UL40 peptide modulates the magnitude of the adaptive NKG2C+ NK cell development in KTR.

Additional observations in this study may contribute to better understanding adaptive NK cell development in KTR. First, despite that the increment of NKG2C+ NK cells did not correlate with CMV copy numbers by regression analysis, the group defined as NKG2C expanders displayed a significantly greater viral load at diagnosis. Remarkably NKG2C+ NK cell expansions were not perceived in three out of six cases suffering CMV tissue invasive disease. The lack of a response coincided with low viremia values (e.g., Supplemental Table II, cases 945, 1054), previously reported in gastrointestinal CMV infection (47). These data suggest that development of the adaptive NK cell response may be influenced by the tissue location and extension of viral replication (i.e., epithelial versus hematopoietic tissues). Second, CMV DNAemia below LLOQ levels was detected early in some KTR with high pretransplant proportions of adaptive NKG2C+ NK cells, who did not receive antiviral therapy and did not display viremia in the late posttransplant survey (Supplemental Table II, cases 1002, 1056, 1079). These observations are consistent with the reported association of pretransplant adaptive NKG2C+ NK cells with reduced incidence of symptomatic CMV infection, suggesting that rather than preventing early viral replication, they contribute to contain infection progression (26). Third, although adaptive NKG2C+ NK cell expansions of KTR generally persisted, in some cases, they declined at late time points. Given that healthy individuals maintain such expansions for years (25), effective CMV control is an unlikely explanation for these decrements. As an alternative interpretation, it is conceivable that such late reductions of adaptive NK cells may reflect their altered turnover (48), resulting from the combined effects of sustained immunosuppression and exhaustion under CMV pressure.

Altogether, these observations provide a broader perspective on the course of the adaptive NKG2C+ NK cell response to CMV in KTR, illustrating its individual variability. We hypothesize that the magnitude of the response might be directly related with an inefficient/delayed T cell–mediated CMV replication control, potentially modulated by additional factors as detailed above. This interpretation is based on the observation that NKG2C+ NK cell expansions were mostly detected in viremia(+) cases and further supported by previous studies relating: 1) adaptive NKG2C+ expansions with defective T cell–mediated responses to CMV and 2) pretransplant CMV-specific T cells and adaptive NK cells with a reduced risk of posttransplant viremia. In contrast, following their development, adaptive NKG2C+ NK cells may contribute with T cells to restore CMV control. Altogether, our results indicate that analysis of adaptive NKG2C+ NK cells in combination with CMV-specific and TcRγδ Vδ2 T cells contributes to assess the development of the immune response to the viral infection in KTR.

We thank Sara Alvarez and Anna Faura (Nephrology Research Support team) for coordinating sample acquisition from KTR, Xavier Duran for statistical analysis, and Dulce Soto and Gemma Heredia for technical support in the Immunology Laboratory.

This work was supported by grants from Fundació La Marató de TV3 (201822-10), Agencia Estatal de Investigación–FEDER (PID2019-110609RB-C21-C22/AEI/10.13039/501100011033), Instituto de Salud Carlos III, Fondo de Investigaciones Sanitarias FIS-FEDER (PI13/00598, PI16/00617, PI20/00090), and RedinRen-FEDER (RD16/0009/0013).

The online version of this article contains supplemental material.

Abbreviations used in this article

IQR

interquartile range

KTR

kidney transplant recipient

LLOQ

lower limit of quantification

PLZF

prolymphocytic leukemia zinc finger transcription factor

QNAT

quantitative nucleic acid amplification testing

t-SNE

t-distributed stochastic neighbor embedding

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The authors have no financial conflicts of interest.

Supplementary data