Functional Impairment of Cytomegalovirus Specific CD8 T Cells Predicts High-Level Replication After Renal Transplantation
2008; Elsevier BV; Volume: 8; Issue: 5 Linguagem: Inglês
10.1111/j.1600-6143.2008.02191.x
ISSN1600-6143
AutoresFrank Mattes, Anita Vargas, J Kopycinski, Emma Hainsworth, P. Sweny, Gaia Nebbia, A. Bazeos, Mark W. Lowdell, Paul Klenerman, Rodney E. Phillips, P. D. Griffiths, Vincent C. Emery,
Tópico(s)Renal Transplantation Outcomes and Treatments
ResumoAmerican Journal of TransplantationVolume 8, Issue 5 p. 990-999 Free Access Functional Impairment of Cytomegalovirus Specific CD8 T Cells Predicts High-Level Replication After Renal Transplantation Correction(s) for this article Erratum Volume 11Issue 5American Journal of Transplantation pages: 1107-1107 First Published online: April 26, 2011 F. M. Mattes, F. M. Mattes InfectionSearch for more papers by this authorA. Vargas, A. Vargas Sir Peter Medawar Pathogen Research Centre, University of Oxford, Oxford, UKSearch for more papers by this authorJ. Kopycinski, J. Kopycinski InfectionSearch for more papers by this authorE. G. Hainsworth, E. G. Hainsworth InfectionSearch for more papers by this authorP. Sweny, P. Sweny Renal Transplant Unit, Royal Free and University College Medical School (Hampstead campus) and Royal Free Hampstead NHS Trust, London, UKSearch for more papers by this authorG. Nebbia, G. Nebbia InfectionSearch for more papers by this authorA. Bazeos, A. Bazeos Renal Transplant Unit, Royal Free and University College Medical School (Hampstead campus) and Royal Free Hampstead NHS Trust, London, UKSearch for more papers by this authorM. Lowdell, M. Lowdell HaematologySearch for more papers by this authorP. Klenerman, P. Klenerman Sir Peter Medawar Pathogen Research Centre, University of Oxford, Oxford, UKSearch for more papers by this authorR. E. Phillips, R. E. Phillips Sir Peter Medawar Pathogen Research Centre, University of Oxford, Oxford, UKSearch for more papers by this authorP. D. Griffiths, P. D. Griffiths InfectionSearch for more papers by this authorV. C. Emery, Corresponding Author V. C. Emery Infection * Corresponding author: Vincent C. Emery, v.emery@ucl.ac.uk v.emery@ucl.ac.ukSearch for more papers by this author F. M. Mattes, F. M. Mattes InfectionSearch for more papers by this authorA. Vargas, A. Vargas Sir Peter Medawar Pathogen Research Centre, University of Oxford, Oxford, UKSearch for more papers by this authorJ. Kopycinski, J. Kopycinski InfectionSearch for more papers by this authorE. G. Hainsworth, E. G. Hainsworth InfectionSearch for more papers by this authorP. Sweny, P. Sweny Renal Transplant Unit, Royal Free and University College Medical School (Hampstead campus) and Royal Free Hampstead NHS Trust, London, UKSearch for more papers by this authorG. Nebbia, G. Nebbia InfectionSearch for more papers by this authorA. Bazeos, A. Bazeos Renal Transplant Unit, Royal Free and University College Medical School (Hampstead campus) and Royal Free Hampstead NHS Trust, London, UKSearch for more papers by this authorM. Lowdell, M. Lowdell HaematologySearch for more papers by this authorP. Klenerman, P. Klenerman Sir Peter Medawar Pathogen Research Centre, University of Oxford, Oxford, UKSearch for more papers by this authorR. E. Phillips, R. E. Phillips Sir Peter Medawar Pathogen Research Centre, University of Oxford, Oxford, UKSearch for more papers by this authorP. D. Griffiths, P. D. Griffiths InfectionSearch for more papers by this authorV. C. Emery, Corresponding Author V. C. Emery Infection * Corresponding author: Vincent C. Emery, v.emery@ucl.ac.uk v.emery@ucl.ac.ukSearch for more papers by this author First published: 14 April 2008 https://doi.org/10.1111/j.1600-6143.2008.02191.xCitations: 78 Present address for F. M. Mattes: Department of Virology, Barts and the London Hospital NHS Trust, Whitechapel London E1 1BB. AboutSectionsPDF ToolsRequest permissionExport citationAdd to favoritesTrack citation ShareShare Give accessShare full text accessShare full-text accessPlease review our Terms and Conditions of Use and check box below to share full-text version of article.I have read and accept the Wiley Online Library Terms and Conditions of UseShareable LinkUse the link below to share a full-text version of this article with your friends and colleagues. Learn more.Copy URL Share a linkShare onFacebookTwitterLinked InRedditWechat Abstract Human cytomegalovirus (HCMV) remains an important cause of morbidity after allotransplantation, causing a range of direct effects including hepatitis, pneumonitis, enteritis and retinitis. A dominant risk factor for HCMV disease is high level viral replication in blood but it remains unexplained why only a subset of patients develop such diseases. In this detailed study of 25 renal transplant recipients, we show that functional impairment of HCMV specific CD8 T cells in the production of interferon gamma was associated with a 14-fold increased risk of progression to high level replication. The CD8 T-cell impairment persisted during the period of high level replication and was more prominent in patients above 40 years of age (odds ratio = 1.37, p = 0.01) and was also evident in dialysis patients. Threshold levels of functional impairment were associated with an increased risk of future HCMV replication and there was a direct relationship between the functional capacity of HCMV ppUL83 CD8 T cells and HCMV load (R2= 0.83). These results help to explain why a subset of seropositive individuals develop HCMV replication and are at risk of end-organ disease and may facilitate the early identification of individuals who would benefit from targeted anti-HCMV therapy after renal transplantation. Abbreviations: HCMV human cytomegalovirus LCMV lymphocytic choriomeningitis virus Introduction Historically, human cytomegalovirus (HCMV) infection has been a significant clinical problem following organ transplantation. The virus can lead to a range of overt clinical symptoms such as prolonged fever, hepatitis, enteritis, pneumonitis and retinitis (termed the 'direct effects') as well as a number of 'indirect effects' including acute and chronic organ rejection, accelerated cardiovascular disease, new onset of posttransplant diabetes mellitus and bronchiolittis obliterans (1, 2). In recent years, the development of prophylactic and pre-emptive therapeutic patient management strategies has significantly reduced the mortality attributed to HCMV and impacted positively on morbidity (3-6). A variety of anti-HCMV agents (valaciclovir, ganciclovir [intravenous and oral] and most recently valganciclovir) have been shown in controlled clinical trials to be effective for prophylaxis of solid organ transplant recipients at high risk of HCMV disease, such as HCMV seronegative patients receiving an organ from a HCMV seropositive donor (7-10). However, HCMV disease still occurs in a subset of patients after prophylaxis is stopped, and disease at this stage can still be clinically severe (4). Pre-emptive therapy initiated on the basis of detection of high levels of virus in blood by PCR or antigenemia assays has also been shown to minimize disease and provide rapid control of HCMV replication (10-12), although it may not impact on the longer term consequences of HCMV infection highlighted above. In addition to the high-risk patients mentioned above, seropositive recipients are also at risk of HCMV disease, especially if they receive an organ from a seropositive donor and become reinfected (13). As these seropositive patients form the majority of patients in many areas of the world, the absolute numbers experiencing HCMV disease cannot be ignored. In vivo, HCMV replication is highly dynamic with doubling times between 0.4 and 2 days (14). The basic reproductive number (Ro; the number of newly infected cells in the host arising from one infected cell) for HCMV in immune liver transplant recipients is ∼15 but is reduced to 2.4 in patients who are already HCMV seropositive (15). Several studies, including many from our group, have shown that high level HCMV replication, as detected by high viral loads in blood by quantitative virological assays, and persistant viral load with the equivalent viral turnover are a dominant risk factors for HCMV disease when examined in multivariable statistical analyses (16-20). T-cell immunity is a fundamental effector process controlling HCMV replication in vivo (21-24). The CD8 T-cell immune response to HCMV appears to be dominated by responses to the tegument protein ppUL83 and to the immediate early protein pUL123 (22, 25) although recent data using peptides representing the entire HCMV proteome reveal that many proteins are targeted by the host CD8 T-cell response (26). Studies using class I HLA tetramer reagents have shown that the human host devotes a high proportion of the total CD8 T-cell response to the control of HCMV. In healthy individuals this response averages ∼1% and can reach levels of 10–50% in acute infection in immunocompromised hosts (27-30). In transplant patients, CD4 T-cell help is a key factor to facilitate CD8 T-cell control of HCMV replication (31, 32). While we have a substantial amount of qualitative data relating to the CD8 T-cell immune control of HCMV after transplantation, there are relatively few studies addressing whether specific quantitative malfunction of these CD8 T cells, in addition to their frequency and absolute number, also contributes to the failure to suppress replication and whether these assessments provide prognostic value. We, therefore, set out to investigate the functional capacity of HCMV specific ppUL83 specific T cells in patients who did, or did not, develop high level HCMV replication after renal transplantation. Our results show that patients who progressed to high level replication had a HCMV ppUL83 CD8 T-cell population that was functionally impaired compared to patients who kept HCMV replication suppressed to undetectable levels. Materials and Methods Study population and design A prospective natural history study, investigating the role of HCMV specific CD8 T cells in controlling HCMV virus replication postrenal transplantation, was carried out in a single center (Royal Free Hospital, London, UK) between October 2000 and December 2001. In total, 25 renal transplant patients entered the study, with the following HLA class I types: HLA-A * 0201 (n = 14), HLA-B * 0702 (n = 5), HLA-B * 0801 (n = 7) and HLA-B * 3501 (n = 6). A further 10 patients were excluded since they did not have class I HLA types with appropriate HLA tetramer reagents available. Preservative free heparin blood samples were collected for isolation of peripheral blood mononuclear cells (PBMC) on a weekly basis. PBMCs were isolated on the same day and cryopreserved for immunological studies. In addition, we analyzed PBMCs taken from 10 healthy HCMV seropositive controls, 10 patients with common variable immunodeficiency to provide a suitable control group with a specific deficit in antibody production and 10 patients who were receiving dialysis but had not been transplanted. Citrated blood was collected for HCMV surveillance three times a week when the patient was hospitalized or whenever attending an outpatient clinic. The study was approved by the Local Research Ethics Committee (Institutional Review Board), and all clinical investigations were conducted according to the principles expressed in the Helsinki Declaration. DNA extraction, qualitative and quantitative HCMV PCR DNA was extracted from 200 μL whole blood using a Qiagen extraction kit (Minden, Germany) according to manufacturer's instructions. Qualitative HCMV PCR was carried out as described elsewhere (16), with a lower sensitivity level of 200 genomes/mL whole blood. Quantification of HCMV was performed with a TaqMan (ABI)–based method (Cheshire, UK) adapted from our previously published method (33). In the current study, high-level replication (henceforth referred to as viremia) was defined as viral loads above 200 genomes/mL whole blood. Antiviral and immunosuppressive therapy None of the patients received antiviral prophylaxis for HCMV. Patients with two consecutive positive HCMV PCR samples (>200 genomes/mL blood) received pre-emptive anti-HCMV therapy with either i.v. ganciclovir (5 mg/kg bd adjusted for creatinine clearance, n = 7 patients) or i.v. foscarnet plus i.v. ganciclovir each at half dose (n = 3) as part of a previously published randomized clinical trial which showed that these two treatments were equipotent (12). Antiviral therapy was continued until two consecutive PCR negative samples were obtained. Initial immunosupression consisted of prednisolone plus tacrolimus, alone or with sirolimus, basiliximab, or mycophenolate mofetil, prednisolone plus cyclosporin alone or in combination with either sirolimus or azathioprine. The majority of the patients (72%) received induction therapy with baxiliximab as part of their immunosuppressive regimen. Isolation of PBMCs from whole blood Whole blood (20 mL) was collected using the monovette system, containing preservative free heparin. PBMCs were isolated by centrifugation through Ficoll-Paque (600 g, 20 min at room temperature; Amersham-Pharmacia Biotec, Bucks, UK). The supernatant containing mononuclear cells was transferred to a new 50-mL tube and washed three times in RPMI 1640 supplemented with 2-mM glutamine. Purified PBMCs were counted and cryopreserved in fetal calf serum (FCS) containing 10% dimethyl sulphoxide, at a concentration of 3 × 106 to 6 × 106 per mL, and stored in liquid nitrogen. Tetrameric complex and surface marker staining HCMV specific tetrameric complexes were constructed for HLA-A * 0201, HLA-B * 0702, HLA-B * 0801 and HLA-B * 3501, as described elsewhere (34), using the appropriate ppUL83-derived immunogenic peptides (A2: NLVPMVATV, B7: TPRVTGGGA, B8: DANDIYRIF, B35: IPSINVHHY). All samples from an individual patient were analysed in the same assay to minimize variation. PBMCs were thawed by placing the vial in a 37°C water bath. Lymphocytes were washed in RPMI and R10 (RPMI 1640 containing 10% FCS), resuspended in 2 mL of R10 and incubated for 2 to 3 h at 37°C prior to staining. Lymphocytes were counted and the cell number adjusted to 2 × 106–4 × 106 cells/mL. Fifty microliters of the cell suspension were transferred into a FACS tube (Becton Dickinson) together with 0.3 μg (in PBS) of the HLA-A2 or HLA-B35 tetramer or 1 μg (in PBS) of the HLA-B7 or HLA-B8 ppUL83-specific tetrameric complexes. After incubation for 30 min at 37°C, cells were subsequently stained for the surface marker CD8 by adding 5 μL of antihuman Tri-color labeled CD8+ antibody (Becton Dickinson, Oxford, UK). Surface marker staining was carried out on ice for 30 min and the cells then washed with PBS/0.1% sodium azide and resuspended in 200 μL of 2% freshly prepared paraformaldehyde. All results were acquired on a FACScalibur (Becton Dickinson) and analyzed with the CellQuest software (Becton Dickinson). Enzyme-linked immunospot (ELISPOT) for interferon-gamma producing CD8+ T cells A multiscreen, 96-well filtration plate (Millipore, Watford, UK) was coated with 50 μL of 1:66 dilution of antihuman IFN-γ antibody (D1K, Mabtech, Nacka Strand, Sweden) in filtered PBS and incubated overnight at 4°C. Unbound antibody was removed by washing the plate six times with filtered PBS. Cryopreserved PBMCs were thawed as described above and incubated for at least 3 h at 37°C before peptide stimulation. Either 5 × 104 or 1 × 105 of lymphocytes (in a 90 μL volume) were added to each well together with 10 μL of the ppUL83 peptide (200 μM stock in RPMI) or R10 medium for the control well. All ELISPOT assays were carried out in triplicate. After 16 h incubation at 37°C / 5% CO2, cells were removed by washing the plates four times with PBS containing 5% Tween 20 and twice with PBS. Fifty microliters of biotinylated anti-IFN-γ antibody was added (1:1000 dilution, 7-B6-1-biotin; Mabtech) and incubated for 3 h at room temperature. The ELISPOT plate was washed a further six times with PBS/Tween 20 and incubated for 2 h with streptavidin-ALP substrate (Mabtech, Hamburg, Germany) followed by the addition of an alkaline phosphatase conjugate substrate (50 μL; Bio-Rad, Hemel Hempstead, UK). The resulting spots were counted semi-automatically with an ELISPOT reader. Results were expressed as percentage of cells secreting IFN-γ after subtracting the number of spots due to spontaneous IFN-γ release (measured in the control wells) from the number of spots obtained in the wells incubated with the ppUL83 peptide. Recent data obtained using the ELISPOT assay for IFN-γ and an intracellular cytokine assay have shown high correlation between the two assays and similar functional impairment of HCMV specific CD8 T cells as observed in the current study (data not shown). Statistical analyses The ppUL83 specific HCMV CD8+ T-cell frequency was recorded as a percentage of CD8 cells or total lymphocytes. Multiple samples derived from the same patient were expressed as a median ppUL83 specific HCMV CD8+ T-cell frequency. The relative ppUL83 specific CD8+ T-cell frequency was calculated as a percentage of CD8+ T cells or as a percentage of lymphocytes for comparison with the ELISPOT data. Samples with less than 20 000 total events on the FACS plots were excluded from the analysis. The number of cells secreting IFN-γ was expressed as percentage of cells secreting IFN-γ relative to the total number of cells used in the assay. These percentages were log transformed to provide a normal distribution before performing further statistical tests. Data were stratified according to whether patients experienced high level replication (viremia >200 genomes/ mL blood) with a sub-stratification to samples taken before, during and after the episode of high level replication. Samples from patients who did not experience HCMV viremia ( 200 genomes/mL blood) and were given pre-emptive antiviral therapy. Four patients were HCMV IgG negative at transplantation and received an organ from a HCMV seronegative donor. None of these became HCMV viremic. In contrast, the two seronegative patients who received a kidney from a HCMV seropositive donor became viremic. Eight of the 19 HCMV seropositive patients, who received either a HCMV negative or positive kidney, became viremic. This HCMV immune experienced patient group (n = 19) was studied in more detail to determine factors associated with appearance of HCMV viremia. A more comprehensive summary of patient demographics is shown in Table 1. Recipients who experienced high level HCMV replication (viremic) were significantly older than those who maintained viral replication below detectable levels (viremic patients: 56 [range 22–67] years versus nonviremic patients: 26 [range 19–59] years; p = 0.02). This age effect was also observed if the analysis was restricted to the 19 patients with pre-existing immunity to HCMV. There were no significant differences in the immunosuppressive therapy (baseline or induction) received by patients who did or did not experience high level HCMV replication. In addition, there were no significant differences in the donor/recipient HCMV serostatus and incidence of viremia in the 10 patients excluded from the study (70% R+, 40% viremia) compared to the patients who comprised the study cohort. Table 1. Demographic characteristics of the study population Characteristic High level replication No (n = 15) Yes (n = 10) Sex (male/female) 10/5 7/3 Renal graft (deceased/living donor) 13/2 9/1 Median Follow up (days) 363 397 Number of samples (per patient) 13 13 Immunosuppression Cyclosporin based 13 4 Tacrolimus based 2 4 Other 0 2 Induction therapy 10 8 Donor (D) and recipient (R) HCMV serostatus D–R– 4 0 D–R+ D+R+ 11 8 D+R– 0 2 HCMV specific CD8+ T-cell frequencies in HCMV seropositive patients using class I HLA tetramers In the 11 HCMV seropositive recipients who remained PCR-negative (HCMV load < 200 genomes/mL blood) following transplantation, the median ppUL83 specific CD8+ T-cell frequency was 0.28% (0.02–3.35) prior to day 50 and 0.37% (0.01–6.92) after day 50 (p nonsignificant). In the eight HCMV seropositive patients who experienced high-level replication, the HCMV specific CD8+ T-cell frequency was elevated prior to viremia (0.71%[0.29–3.65]) during viremia (1.19%[0.27–5.2]) and following viremia (0.5%[0.03–6.46]) compared with patients not experiencing viremia (p = 0.008). In all samples analyzed, patients with viremia had a significantly higher ppUL83 specific CD8+ T-cell frequency by HLA tetramer analysis than patients who remained PCR negative (p = 0.008) (Figure 1). Figure 1Open in figure viewerPowerPoint Frequency of ppUL83 specific tetramer+(tet+) CD8+ T cells in patients with and without HCMV high level HCMV replication according to time posttransplantation or relative to the period of viremia. Data are expressed as a proportion of total CD8 T cells. The median value of each dataset is shown as a horizontal within the box which encompasses the 25th and 75th percentiles. The whiskers extending from either end of the box represent the extent of the data. Using ANOVA, there was a significant difference between the groups stratified according to high level replication (yes or no, p = 0.008). IFN-γ secretory capacity of HCMV specific CD8+ T cells from HCMV seropositive patients In the 11 HCMV seropositive patients who remained PCR-negative, the median frequency of IFN-γ positive ppUL83 epitope specific CD8+ T cells (as a percentage of the total PBMC population) was 0.023% (0.005–1.08) before day 50 and 0.032% (0.005–4.66) after day 50 (Figure 2). For comparison with the HLA tetramer data, the ELISPOT frequency prior to day 50 is equivalent to a frequency of 0.3% in the specific CD8 T-cell compartment. In patients with high-level replication, a frequency of 0.027% (0.005–0.009) (∼0.14% of the CD8 T cells) was observed before viremia, 0.082% (0.005–0.87) during viremia and 0.17% (0.005–0.82) after viremia (Figure 2). In contrast to the results obtained with the ppUL83 HCMV specific tetramers, there was no significant difference in the frequency of IFN-γ producing CD8 T cells after peptide stimulation between patients who did or did not experience high-level replication (p = 0.35). Figure 2Open in figure viewerPowerPoint Frequency of ppUL83 specific PBMCs producing IFN-γ following peptide stimulation in patients with or without high level HCMV replication according to time posttransplantation or relative to the period of viremia. It should be noted that data are expressed as a proportion of total PBMCs. The data are presented as described in the legend to Figure 1. Using ANOVA, there were no significant differences between the groups (p = 0.35). Functional impairment of ppUL83 specific CD8+ T cells and failure to control HCMV replication There was a strong correlation between the frequency of ppUL83 specific IFN-γ secreting CD8 cells and CD8 cells identified using ppUL83 specific tetrameric complexes in patients with or without viremia (R2= 0.49 viremic patients, R2= 0.41 non-viremic patients, p = 0.01; Figure 3). However, patients who experienced high-level replication had significantly fewer cells able to secrete IFN-γ following peptide stimulation. This functional impairment is illustrated in Figure 4, where the ratio of the HCMV CD8+ T cells identified using class I HLA tetramers and using the IFN-γ ELISPOT assay are compared. The difference in peptide responsiveness of the ppUL83 CD8 T cells stratified according to high-level HCMV replication as a binary variable was highly significant (p = 0.016). We also performed similar analyses on a group of healthy HCMV seropositive individuals (n = 10) and also a group of patients with common variable immune deficiency (CVID; n = 10). These data revealed that 65.3% (±26.5%) of the tetramer+ cells from healthy seropositive individuals could produce IFN-γ after peptide stimulation, which was comparable to the HCMV specific CD8 T cells present in patients with CVID (59.4 ± 22.2%, p = 0.6). In a group of dialysis patients (n = 10), a similar distribution was observed with an average of 47.5% (± 38.4%) of the HCMV CD8 tetramer positive T cells being able to secrete IFN-γ although this ranged from 0.71 to 107% for individual patients (p = 0.21 and p = 0.38 when compared with healthy individuals and CVID patients, respectively). Comparison of these data with the average capacity of the HCMV CD8 tetramer+ T cells to produce IFN-γ in the renal transplant patients showed that there were no significant differences in this ratio between renal transplant patients, who remained HCMV PCR negative, and the healthy controls/ CVID patients (p = 0.89) and dialysis patients (p = 0.08); whereas the ratio was significantly lower in renal transplant patients who experienced HCMV viremia and the healthy controls and CVID patients ((p = 0.006) but not against the dialysis patients (p = 0.09). Figure 3Open in figure viewerPowerPoint Scatterplot showing the correlation between ppUL83 CD8+ T cells identified by class I HLA tetramer staining and those secreting IFN-γ following peptide stimulation in patients with and without viremia. Figure 4Open in figure viewerPowerPoint Comparison of the proportion of ppUL83 CD8+ T cells able to produce IFN-γ following peptide stimulation between patients with or without high level HCMV replication. The data are presented as described in the legend to Figure 1. Factors associated with high-level HCMV replication after renal transplantation A series of univariable and multivariable logistic regression models were used to determine the relative importance of the risk factors identified in this study, which predispose renal transplant recipients to high-level HCMV replication. Importantly, these models were based on samples taken before patients became HCMV PCR positive, that is when viral loads exceeded 200 genomes/mL blood. Univariable models showed that each 10-fold increase in ppUL83 specific CD8 T-cell frequency was associated with an odds ratio of 2.15 for high-level replication. In contrast, a lower frequency of IFN-γ secreting ppUL83 CD8 T cells was associated with high-level HCMV replication (odds ratio = 3.03, 96% CI 1.25–7.14). However, it was the functional impairment of these CD8 T cells (the proportion of tetramer positive cells that could secrete IFN-γ following peptide stimulation) that was associated with the greatest risk (odds ratio = 6.7) for the development of high-level replication. Recipient age above 40 years was also associated with an elevated risk of developing high-level replication (odds ratio = 1.07). In bivariable and multivariable additive models, both age and functional impairment of ppUL83 CD8 T cells remained independent risk factors for high level HCMV replication after renal transplantation. A summary of the models is shown in Table 2. Table 2. Univariable and multivariable logistic regression models relating immune parameters and age with future appearance of high level replication Model Variable1 Odds ratio 95% Confidence interval p-Value Univariable Age (per 1 year increase) 1.12 1.07–1.17 0.02 ppUL83 tet+ CD8+ frequency (per 10-fold increase) 2.15 0.99–4.70 0.05 ppUL83 specific IFN-γ secreting cells (per 10-fold decrease) 3.03 1.25–7.14 0.01 Proportion of HCMV tetramer+ CD8 T cells secreting IFN-γ (per 10-fold decrease) 6.7 1.5–33.3 0.01 Multivariable Proportion of HCMV tetramer+ CD8 T cells secreting IFN-γ (per 10-fold decrease) 14.2 2.7–100 0.02 Age (above 40 years) 1.37 1.07–1.75 0.01 1Model based on log10 transformed frequencies. Following these logistic regression analyses, we constructed graphs relating the probability of a patient developing high-level replication based upon the functional impairment of their ppUL83 specific CD8– T cells (Figure 5, top panel). The graphs revealed a rapid increase in the probability of viremia once critical levels of functional impairment became evident. Indeed, the probability can increase substantially over relatively small decreases in functional capacity. For example, the probability of high-level replication is 10% when the functional CD8 impairment is 50% of normal but increases to 60% when only 10% of the HCMV ppUL83 tetramer+ CD8 T cells can secrete IFN-γ. The effect of age both changes the shape of the probability curve and shifts it to the left indicating that a patient above 40 years of age has a significantly increased risk of high-level replication since their HCMV tetramer+ CD8 T cells are already impa
Referência(s)