Deciphering Effects of Uncontrolled Cytomegalovirus Replication on Immune Responses in Cytomegalovirus DNA-Positive Renal Transplant Recipients

Abstract

Cytomegalovirus (CMV) is a highly prevalent virus and a common cause of morbidity in solid organ transplant patients. It is also known for its long-lasting imprint on the immune system, expanding populations of highly differentiated T cells and natural killer (NK) cells with novel phenotypes. However, it is unclear whether these cells mark success or failure in the management of an active infection. We assessed CMV reactivation in 54 renal transplant recipients (RTRs) by measuring CMV DNA in plasma samples. Function and phenotype of T cells and NK cells were then assessed in seven RTR with detectable CMV DNA. The patient with highest CMV viral load (P1) displayed increased NK cell function and abundant highly differentiated T cells. We compare P1 with the other six patients and review possible scenarios of cross-regulation between NK cells and T cells.

Introduction

Cytomegalovirus (CMV) is a common opportunistic infection with a seroprevalence of 50–80% worldwide (10). It can remain quiescent with periodic reactivations triggered by immune deficiency and/or inflammation. In immunosuppressed individuals such as solid organ transplant recipients, active infections are common and have clinical consequences (12). CMV reactivation and the immune responses invoked create characteristic and novel populations of natural killer (NK) cells and T cells (5). It is unclear whether these novel populations regulate each other to control CMV or induce adverse outcomes.

Activated NK cells can activate adaptive immune responses through direct and indirect mechanisms (11). For example, interferon gamma (IFNγ) production by activated NK cells can drive Th1 differentiation and recruit CD8⁺ T cells to the lymph node (22). In addition, NK cells can promote dendritic cell (DC) maturation and activation via cell to cell contact or via IFNγ and tumor necrosis factor alpha (TNFα) production (1). Once mature, activated DC can stimulate T cell expansion, which ultimately confers protection against infection (15). Reciprocally, T cells can activate or regulate NK cell responses. For example following Leishmania infection, interleukin (IL)-2 production by CD4⁺ T cells activates and induces production of IFNγ by NK cells (6). Moreover, T regulatory (Treg) cells can influence NK cell activation by depriving them of IL-2 (18).

Activated NK cells can reduce T cell function via IL-10 and transforming growth factor beta production (16,23) or by direct lysis. For example in perforin-deficient mice, murine cytomegalovirus (MCMV) induced production of IL-10 by NK cells that suppressed CD8⁺ T cell effector function and enhanced persistent infection (19). In addition, NK cells can kill activated T cells that upregulate ligands for NK cell receptors, such as the stress ligands that bind to NKG2D (8).

To prevent NK cell-mediated killing, T cells can express inhibitory receptors. For example, activated CD4⁺ T cells upregulate Human Leukocyte Antigen (HLA)-E, the ligand for NKG2A (26). Blocking of NKG2A or HLA-E increases NK cell killing of these CD4⁺ T cells (4). NK cells can also limit T cell responses by lysing DC or by decreasing the ability of DC to present antigen (4,8). For example, following MCMV infection, infected DC prime naive T cells in both Ly49H⁻ and Ly49H⁺ mice. However, Ly49H⁺ NK cells induced by MCMV kill antigen-bearing DC, reducing the generation and maintenance of MCMV-specific T cells (3). The negative regulation of T cell responses by NK cells may dampen excessive immune responses following an infection and prevent damage to host tissues, or can hinder the control of infections.

Overall, these findings reveal an intricate association between T cell and NK cell responses following any infection. In this study, we present a case study and review evidence of cross-talk between NK cells and T cells in renal transplant patient in the early stages of a CMV infection that she was ultimately unable to control. The data are compared with 6 other renal transplant recipient (RTR) who were positive for CMV DNA, but were able to control their infections.

Patients and Methods

Fifty-four RTRs with a median (range) age of 54 (27–71) years were recruited from February 2013 to April 2013 from Royal Perth Hospital (RPH), Western Australia. The median (range) time posttransplantation was 8 (2 –18) years. All patients were clinically stable on maintenance immunosuppressive therapy (33 on Tacrolimus, 12 on Sirolimus, 9 on Cyclosporin with Mycophenolate, and/or Prednisolone), with no apparent CMV disease or reactivation and no antiviral treatment within 6 months of recruitment. Fifty milliliters blood and a saliva sample were collected from each patient. The patient presented as a case study (P1) filled the criteria of the study when recruited in 2013. All participants provided written informed consent and the project was approved by the Human Research Ethics Committees of RPH, the University of Western Australia and Curtin University.

Peripheral blood mononuclear cells (PBMCs) were isolated from heparin-treated blood using Ficoll–Paque density gradient centrifugation and were cryopreserved in liquid nitrogen. Plasma samples were stored at −80°C.

Plasma samples were screened for CMV DNA in the Department of Microbiology (RPH) using commercial kits (Abbott Diagnostics, Lake Forest, IL) and were able to quantitate >20 copies/mL. This value was used as a cutoff to identify CMV DNA positive RTRs. CMV immunoglobulin G (IgG) levels were determined by enzyme-linked immunosorbent assay using CMV glycoprotein B (gB) antigen and presented in arbitrary units (20).

PBMC were thawed, rested overnight at 37°C, and then stained for surface and intracellular markers of NK and T cells using multiparametric flow cytometry (20,21). To assess antibody-dependent cellular cytotoxicity (ADCC), CMV gB antigen was coated onto 96-well plates, incubated overnight at 4°C. On the next day, samples of heat-inactivated autologous plasma (diluted 1 in 300) were added to the coated wells and incubated overnight at 4°C. Wells were washed and PBMC were added with BV786 anti-CD107a (H4A3; BD Biosciences) for 1 h, followed by brefeldin A and monensin (BD Biosciences) for a further 5 h at 37°C. Lineage markers were then stained as before (20,21).

To assess T cell responses, PBMC (1 × 10⁶) were stimulated with overlapping 15mer pp65 or immediate early-1 (IE-1) protein CMV peptides (1 μg/mL for each peptide; JPT Peptide Technologies GmbH, Germany) or anti-CD3 (0.1 μg/mL; Mabtech, Sweden) in the presence of anti-CD28 (0.1 μg/mL) and anti-CD49d (0.1 μg/mL; BD Biosciences). BV786 anti-CD107a was added to all wells. After 2 h, GolgiPlug and GolgiStop (BD Biosciences) were added for a further 5 h (21).

To characterize replicating CMV, DNA was extracted from lithium heparin buffy coats and saliva pellets (Favorgen, Taiwan). Nested primers targeting UL18 and UL40 were retrieved (14) and assays were optimized using DNA extracted from HCMV-infected fibroblasts. Reactions (20 μL) contained each primer (10 μM), PCR buffer with 35 mM MgCl₂, 40 mM dNTPs, Platinum Taq DNA Polymerase (Invitrogen, Carlsbad, CA), DNA (diluted 1:2) for the outer PCRs, and the outer PCR product for the inner PCRs. Cycling conditions for outer PCRs were as follows: 1 cycle of 5 min at 95°C followed by 35 cycles of 30 sec at 95°C, 30 sec at 60°C, and 1 cycle of 1 min at 72°C. Inner PCRs were run through only 30 cycles. Amplicons were separated on 1% agarose gels and underwent Sanger sequencing at the Australia Genome Research Facility. Sequences were analyzed using Geneious version 10.0.3.

Case study

Following end-stage kidney disease secondary to Alport's syndrome, a 53-year-old female (P1) received a kidney from a deceased donor in 1994 (Fig. 1). Both donor and recipient were CMV seropositive at the time of transplant. The recipient did not receive antiviral prophylaxis, and had no known history of CMV disease. Within 6 months posttransplantation, the patient became positive for CMV immunoglobulin M, but viral culture was not performed to confirm active CMV replication. When recruited for our study in February 2013, the recipient was stable on cyclosporin, Mycophenolate and Prednisolone, with no antiviral therapy and no clinical symptoms of CMV infection or any other disease. Her renal function was stable (creatinine 137 μmoles/L) and cyclosporine trough levels were within the recommended range. Hospital records revealed that in May 2013, she was diagnosed with an invasive cutaneous facial squamous cell carcinoma (SCC) with local infiltration causing sinusitis and cranial nerve palsies. This was treated with radiotherapy. In September 2013, she was admitted with a febrile illness, with a diagnosis of multipathogen pneumonia, being positive in a bronchial lavage for Aspergillus and Pneumocystis jiroveci and positive in bronchial lavage and plasma for CMV. She was commenced on intravenous (IV) antibiotic therapy (Tazocin, azithromycin, cotrimoxazole) and IV ganciclovir, with a stepdown to oral cotrimoxazole and valganciclovir. In July 2014 she presented with sepsis secondary to orbital cellulitis attributed to her progressive SCC. Imaging revealed extensive osseous and intracranial involvement ultimately leading to her death.

FIG. 1.

Case study timeline.

NK and T cell profiles of RTR with detectable CMV DNA

CMV DNA was detected in plasma from 7/54 (13%) RTR, with viral loads of 22–2,717 copies/mL. P1 had the highest burden of CMV (2,717 copies/mL) and the burden of CMV correlated with time posttransplant (Spearman's test; r = 0.8, p = 0.02) but not with age (r = 0.35, p = 0.50). P1 also had the lowest CMV gB IgG titer and the highest frequency of NK cells with expressing the activating receptor (NKG2C), the inhibitory receptor (LIR-1) and lacking the signaling adaptor molecule (FcRγ⁻). Although expression of NKG2C correlates with CMV seropositivity, we have shown that proportions of FcRγ⁻ LIR-1⁺ NKG2C⁻ NK cells are higher in CMV DNA⁺ and CMV seropositive patients than FcRγ⁻ LIR-1⁺ NKG2C⁺ or FcRγ⁻ LIR-1⁻ NKG2C⁺ NK cells (20). When proportions of NK cells expressing FcRγ⁻ LIR-1⁺ NKG2C⁻ were compared among the seven CMV DNA⁺ RTR, P1 had the highest frequency of these cells (Table 1). Moreover, in an ADCC assay, her FcRγ⁻ LIR-1⁺ NKG2C⁻ NK cells displayed the highest expression of cytotoxic marker (CD107a) and TNFα following stimulation with her own plasma, anti-CD16 (used as a positive control) or K562 cells (used to assess NK cell cytotoxicity; Table 1). In addition, her CD4⁺ T cell counts (80 cells/μL) and CD4:CD8 ratio (0.08) were low relative to the normal range used in clinical care (457–1,498 and 0.72–4.18, resp.), while her CD8⁺ T cell and NK cell counts were high or normal (CD8: 1,020 cells/μL, normal range 205–1,013; NK cells: 190 cells/μL, normal range 93–575). The patient had a low frequency of CD4⁺ T cells and these were enriched for terminally differentiated (CD27⁻ CD45RA⁺) T_EMRA cells (Table 2). Most of her CD4⁺ effector memory T cells (CD4⁺ CD27⁻ CD45RA⁻ CD57⁺) expressed NK cell inhibitory receptors, LIR-1 and KLRG1. These cells expressed CD107a (a marker of cytotoxicity) and TNFα efficiently upon stimulation with CMV pp65 antigen. However, P1 had the lowest proportion of terminally differentiated (CD27⁻ CD45RA⁺) CD8⁺ T cells.

Table 1.

Cytomegalovirus DNA Levels, Antibody Titers, and Natural Killer Cell Phenotypes in Cytomegalovirus DNA⁺ Renal Transplant Recipient (n = 7)

	P1	P2–P7
Age	72	53 (41–66)
CMV DNA copies/mL of plasma	2,717	105 (22–349)
Antibody titer against gB antigen (AU × 10⁻³)	35	50 (39–66)
NK cell phenotype and function
CD56^dim NK cells (% of lymphocytes)	13	8 (4–14)
NKG2C⁺	42	16 (1.5–38)
LIR-1⁺	87	65 (54–74)
FcRγ⁻	88	31 (8.9–45)
FcRγ⁻ LIR-1⁺ NKG2C⁻ (% of CD56^dim NK cells)	41	17 (4–24)
FcRγ⁻ LIR-1⁺ NKG2C⁻ CD107a⁺ TNFα⁺ (following patient plasma stimulation)	11	1 (0–1.7)
FcRγ⁻ LIR-1⁺ NKG2C⁻ CD107a⁺ TNFα⁺ (following anti-CD16 stimulation)	12	1 (0–2.8)
FcRγ⁻ LIR-1⁺ NKG2C⁻ CD107a⁺ TNFα⁺ (following K562 stimulation)	6.9	1 (0.05–2.3)

AU, arbitrary unit; CMV, cytomegalovirus; gB, glycoprotein B; NK, natural killer; TNFα, tumor necrosis factor alpha.

Table 2.

T Cell Frequencies and Function Following pp65 and IE-1 Stimulation in Cytomegalovirus DNA⁺ Renal Transplant Recipients (n = 7)

	P1	P2–P7
T cell phenotype
CD4 T cells (% of lymphocytes)	5.8	45 (25–56)
CD27⁻ CD45RA⁺ (T_EMRA % of CD4)	66	2 (0.4–4.4)
CD27⁻ CD45RA⁻ (T_EM % of CD4)	5	8 (0–18.7)
CD8 T cells (% of lymphocytes)	72	18 (7–27)
CD27⁻ CD45RA⁺ (T_TEMRA % of CD8)	0.8	44 (13–72)
CD27⁻ CD45RA⁻ (T_EM % of CD8)	3.2	11 (2–30)
LIR-1⁺ KLRG1⁺ % of CD57⁺ CD4⁺ T_EMRA	72	31 (3–74)
LIR-1⁺ KLRG1⁺ % of CD57⁺ CD4⁺ T_EM	80	15 (1.8–32)
LIR-1⁺ KLRG1⁺ % of CD57⁺ CD8⁺ T_TEMRA	95	72 (28–98)
LIR-1⁺ KLRG1⁺ % of CD57⁺ CD8⁺ T_EM	90	66 (39–84)
T cell response following pp65 stimulation
CD107a % of LIR-1⁺ KLRG1⁺ CD4 T_EM	9.6	3 (1.4–4.3)
TNFα % of LIR-1⁺ KLRG1⁺ CD4 T_EM	1.5	0.07 (0–0.2)
CD107a % of LIR-1⁺ KLRG1⁺ CD8 T_EM	4.3	3 (0.4–6.4)
TNFα % of LIR-1⁺ KLRG1⁺ CD8 T_EM	2.6	0.5 (0–1.5)
CD107a % of LIR-1⁺ KLRG1⁺ CD8 T_EMRA	1.8	1 (0.1–2.4)
TNFα % of LIR-1⁺ KLRG1⁺ CD8 T_EMRA	0.2	0.05 (0–0.12)
T cell response following IE-1 stimulation
CD107a % of LIR-1⁺ KLRG1⁺ CD4 T_EM	0.3	0.3 (0–1.2)
TNFα % of LIR-1⁺ KLRG1⁺ CD4 T_EM	2.6	0.3 (0–1.3)
CD107a % of LIR-1⁺ KLRG1⁺ CD8 T_EM	1.3	1.5 (0–3.7)
TNFα % of LIR-1⁺ KLRG1⁺ CD8 T_EM	4.3	1.7 (0–7.4)
CD107a % of LIR-1⁺ KLRG1⁺ CD8 T_EMRA	0.9	0.6 (0–1.4)
TNFα % of LIR-1⁺ KLRG1⁺ CD8 T_EMRA	0.3	0.3 (0.02–1.1)

IE-1, immediate early-1.

Modulation of protective T cells by NK cells

NK cells can inhibit the duration and effectiveness of virus-specific CD4⁺ and CD8⁺ T cell responses by reducing the antigen load via the direct killing of target cells or the killing of DCs. During MCMV infection, infected DCs prime naive T cells in both Ly49H⁻ and Ly49H⁺ mice. In brief, Ly49H⁺ NK cells expand in the presence of MCMV compared to Ly49H⁻ mice and transmit activating signals upon binding to m157 (an MCMV-encoded MHC-I homolog). In Ly49H⁺ mice, where the NK cell response is greater, the NK cells kill antigen bearing DCs, reducing the generation and maintenance of virus-specific T cells; while in Ly49H⁻ mice, higher numbers of both MCMV-specific CD8⁺ and CD4⁺ IFNγ⁺ T cells are observed (3,29).

Our work suggests that active CMV may induce antibody response, which then crosslinks Fc receptors on NK cells to initiate the ADCC response. Given the efficient ADCC responses in P1 (Table 1), lysis of DCs by adaptive NK cells (FcRγ⁻ LIR-1⁺ NKG2C⁻) may have contributed to the low frequencies of CD8⁺ T_EMRA cells or CD4⁺ T cells—some of which would have been CMV-specific. Conversely low frequencies of adaptive NK cells may be linked to higher T cell responses, as was observed in another CMV DNA+ RTR (P2). Relative to P1, P2 had low proportions of FcRγ⁻ LIR-1⁺ NKG2C⁻ (11.7%), low ADCC using patient plasma and positive control (0.02–0.04%), and low NK cell cytotoxicity against K562 (0.05%), but displayed high CD107a expression (5.6–11.2%) and IFNγ production (1.7–3.7%) by CD8⁺ T_EM cells expressing LIR-1 and KLRG1 against both pp65 and IE-1 antigen, suggesting that strong CD4⁺ and CD8⁺ T cell responses against CMV may impede the expansion of adaptive NK cells. It is also possible that adaptive NK cells do not kill CMV-activated T cells, but limit T cell responses by reducing the antigen load.

Modulation of NK cells by T cells

CD4⁺ T cells are also implicated in the control of CMV infection. In Ly49H⁻ mice, depletion of CD4⁺ T cells and not CD8⁺ T cells increased the MCMV replication in the salivary gland (3). Here, the low numbers of CD4⁺ T cells may have allowed CMV replication, which in turn drove the expansion of adaptive NK cells. Most CD4⁺ T cells expressed a highly differentiated phenotype (LIR-1⁺ KLRG1⁺ CD4⁺ T_EM) with high cytotoxic responses against structural antigen (CMV pp65). This may reflect an attempt to control active CMV replication. Upon CMV reactivation, IE-1 is the first protein synthesized and presented on the surface of infected cells, so IE-1-specific T cells may activate rapidly to curtail CMV replication. Poor responses to IE-1 may contribute to active replication and higher burden of CMV in P1. CMV-specific CD4⁺ T cells are also critical in licensing DC to generate the expansion of effector and memory CD8⁺ T cells required to prevent lytic CMV infection (13). Low frequencies of circulating effector CD4⁺ T cells suggest that this process may be impaired in P1—reducing frequencies of CD8⁺ T_EMRA cells and thus increasing CMV replication.

Diversity of CMV may be critical

Despite heightened immune response, a high CMV viral load was observed in P1. We therefore sequenced UL18 and UL40 genes from samples of saliva and buffy coat (predominantly blood granulocytes). These genes are functional homologs of HLA-G (the ligand of LIR-1) and the HLA-E (ligand of NKG2A and NKG2C), and may help CMV to escape NK cell killing (25). The sequences showed that the same strain of virus predominated in both samples. This contained known mutations in UL18 that may increase binding capacity to LIR-1 [positions 32 (aspartic acid to glycine, nucleotides 94–96), 33 (aspartic acid to glutamic acid, nucleotides 97–98 and 102), 60 (lysine to arginine, nucleotides 178–180), and 61 (alanine to threonine, nucleotides 181–183)]. However, the UL18 sequenced from P1 also had several other mutations downstream (Supplementary Fig. S1) (7), which may further influence the affinity for LIR-1 (28). Furthermore, sequence chromatographs of both the UL40 and UL18 genes from the buffy coat revealed the presence of at least two viral species. Mixed CMV infections have been associated with increased viral load and are often accompanied to infection with other herpesviruses (2). This may explain her poor clinical course. Moreover, infection with multiple viral species may have challenged her immune response and therefore replication persisted despite increased NK cell responses.

The patient's clinical course should be considered

Few months postrecruitment for our study, P1 was diagnosed with SCC. We cannot rule out the possibility that the changes seen in NK cells can be due to an onset of SCC, but no studies have linked expansions of adaptive NK cells with cancer. In fact, studies are now harnessing CMV-induced adaptive NK cells for cancer immunotherapy (27), and the changes observed in NK cells have been attributed to CMV in other studies (17,30), so the SCC may not be critical here.

The low CD4⁺ T cell count in P1 may reflect the chronic exposure to immunosuppressive drugs rather than cross regulation by NK cells. CMV reactivation is also common in the elderly (24), reflecting and/or driving the accumulation of terminally differentiated T cells (9). Thus, older age of P1 and the long period elapsing since transplantation may be associated with her poor T cell responses to CMV antigens. This may be the primary effect with other changes following on.

Conclusions

Changes in NK cell or T cell populations in CMV seropositive donors have been described in many studies, but none is based in a clinical setting where control of CMV is being progressively lost. We suggest that lower proportions of CD4⁺ T cells and accelerated T cell differentiation in the presence of active CMV replication may have modulated innate immunity, as increased proportions of adaptive NK cells with potential to control the virus were observed in P1. Understanding crosstalk between immune cells driven by CMV may reveal novel biomarkers to identify individuals at risk of uncontrolled CMV replication.

Footnotes

Acknowledgments

The authors thank Anne Warger and Bronwyn Reid for assistance with patient recruitment and the patients and controls who donated blood.

Author Disclosure Statement

The authors have no financial or commercial conflicts of interest to declare.

Funding Information

The study was funded by the Medical Research Foundation of Royal Perth Hospital and the National Health and Medical Research Council of Australia (Grant No. 1068652). N.M. was supported by an India SIRF scholarship from University of Western Australia.

Supplementary Material

Supplementary Figure S1

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