Infection by HIV in the absence of antiretroviral therapy is characterized by chronic T cell activation . Among the various markers of T cell activation, the number of CD8 cells with the CD38+ phenotype is known to be raised in both acute and chronic infection [2–11]. There is some debate over the function of these cells, which could represent HIV-specific or cytokine-activated cells [8,9,12]. Individuals with high levels of CD8+/CD38++ cells during chronic HIV infection tend to experience a faster CD4 cell decline , and develop clinical disease more rapidly than those with lower levels [11,14–19]. This association persists even after adjusting for baseline CD4 cell count and viral load (VL) [14,20], suggesting that additional factors involved in HIV pathogenesis contribute to high levels of CD38 cells. It has also been shown that the initiation of antiretroviral therapy is associated with a reduction in the percentage and absolute number of circulating CD8+/CD38++ cells in both acute and chronic infection [5,6,21–27].
Recent methodological improvements in the techniques used to quantitate CD38 cell surface molecule expression has allowed us to focus on a population of active CD8 T cells that express high levels of this molecule [28–30]. Instead of a bulk assay that measures the mean expression of CD38 molecules among the heterogeneous cohorts of CD8 cells [16,20,28], the utilization of a quantitative gating strategy permits the determination of the absolute numbers of CD8+/CD38++ T cells in whole blood . As CD8 T cells expressing the CD38++ phenotype are rare in normal HIV-negative adults, the test for CD8+/CD38++ T cells can be regarded as a particularly sensitive flow cytometric assay for monitoring levels of immune activation in patients receiving a potent four-drug antiretroviral treatment regimen at the time of primary HIV infection (PHI). The objectives of the present investigation were to assess CD8+/CD38++ T cell levels before starting therapy, to describe their dynamics after the initiation of therapy, with attention to whether the levels of CD8+/CD38++ T cells return to those observed in uninfected individuals, to evaluate the associations between VL and CD8+/CD38++ T cells during therapy, and to assess changes in the levels of CD8+/CD38++ T cell levels in patients with VL below 50 copies/ml.
Materials and methods
Patients were participants in the Quest (GW PROB 3005) Study to evaluate the virological and immunological effects of early treatment intervention with potent quadruple antiretroviral therapy [combivir (zidovudine 300 mg plus lamivudine 150 mg), abacavir 300 mg and amprenavir 1200 mg twice a day] in PHI. The study also includes an added randomized vaccination strategy after at least 72 weeks of highly active antiretroviral therapy (HAART) . Eligible patients had confirmed HIV infection defined by their detectable VL measured by HIV-RNA polymerase chain reaction and either a negative or an evolving antibody response. These were defined as antibody negative measured with the third-generation enzyme-linked immunosorbent assay (ELISA), or if having a positive ELISA, those who had no more than three bands on the HIV Western blot. Patients were enrolled into the study during the period February 1998 to October 1999 from clinical centres in Europe (France, Switzerland, UK, Italy, Sweden, Belgium, Germany, Denmark), Australia and Canada. Baseline (pre-therapy) blood samples were collected for the measurement of VL, T lymphocyte subsets and CD8+/CD38++ T cell counts. Further measurements were made on samples collected at weeks 2, 4, 12, 20, 28, 36, 48 and 96 of treatment.
Viral load measurement
Plasma VL was determined by HIV RNA, either using the current Roche Amplicor assay (Version 1.5; Roche Molecular Systems Inc., Alameida, CA, USA) with a lower limit of detection of 400 copies/ml or the Ultrasensitive method (Version 1.5; Roche Molecular Systems Inc., Alameida, CA, USA) with a lower limit of detection of 50 copies/ml. Assays were performed in one of two central laboratories: Covance GE, Switzerland or Sydpath, Sydney, Australia.
T cell subset measurements
Peripheral blood T cell subset measurements were routinely performed at two co-ordinated trial centres (Royal Free Hospital, London, UK and Sydpath, Sydney, Australia). Absolute CD4 and CD8 T cell counts were determined on 100 μl ethylenediamine tetraacetic acid blood, using a three-colour direct immunofluorescence method, involving a ‘lyse-no-wash’ technique . In one sample tube, CD3+ T cells were autogated and analysed for CD4 and CD8 cell expression using CD4/CD8/CD3 cell triple staining. In the second tube, lymphocyte counts were obtained as the sum of CD3+ T cells, CD19+ B cells and CD16+ natural killer cells using CD16/CD19/CD3 cell triple staining. In a third tube, a titrated CD45RA/CD38/CD8 cell triple antibody cocktail was used. Here, CD8++T lymphocytes and lymphoblasts were gated in a CD8 cell/side scatter histogram and analysed for CD38 and CD45RA cell expression in a two-parameter histogram. Absolute counts were measured directly using a volumetric flow cytometer (Ortho Cytoron Absolute; Ortho Diagnostics, Amersham, UK) or by fluorescence-activated cell sorting (FACS; Becton Dickinson, Mountain View, CA, USA) . The reagents used in the first two tubes were supplied by Ortho Diagnostics. For the third tube, a CD8 cell antibody conjugated to Tricolor was supplied by TCS Biological, Buckingham, UK, and both CD38 cells conjugated to phycoerythrin and CD45RA cells conjugated to fluorescein-isothiocyanate were supplied by Becton Dickinson, (Becton Dickinson, Oxford, UK). Percentages and absolute counts of lymphocyte subpopulations were determined using the Immunocount II software (Ortho Diagnostics). The median CD4 cell count in healthy HIV-negative individuals was 784 cells/mm3 (interquartile ranges; IQR 629, 1200) and the CD8 cell count was 307 cells/mm3 (IQR 225, 479).
Absolute CD8+/CD38++ cell measurement
The CD38++ cell numbers were counted on the Ortho Cytoron Absolute in London and on FACS in Sydney; both laboratories using a similar semi-quantitative gating strategy (Fig. 1). The flow cytometry gates were standardized in both centres on normal adult and fetal cord blood samples. In healthy adults, CD38 cell expression on CD8+ T cells of the ‘memory’ CD 45RO+ cell phenotype is low (CD38−, < 1500 CD38 molecules/cell) whereas 20–40% of CD8+ T cells of the CD45RA+ cell phenotype as well as granulocytes/neutrophils have an intermediate CD38 cell expression (CD38+; 1000–5000 CD38 molecules/cell). Monocytes (but virtually no CD8 T cells, see below) show a high CD38 cell display (CD38++; > 5000 CD38 molecules/cell) . In fetal cord blood, over 75% of immature T lymphocytes are also CD38++ cells. These populations may thus serve as positive controls when setting the gates for the CD38++ cell populations : in fresh adult blood the standard gate was set between CD38+ neutrophils and CD38++ monocytes, and was further adjusted with cord blood for optimal reproducibility (Fig. 1). As a result, only CD8 T cells with high CD38 cell expression (CD38++, > 5000 CD38 molecules/cell) were scored as ‘positive’ and referred to as ‘CD8+/CD38++'. In the blood of healthy HIV-negative individuals these cells are only present in very low numbers (< 20 cells/mm3 median, 11.0; IQR 7.5, 19.8 cells/mm3;Fig. 1), whereas in cord blood over 75% of T lymphocytes are CD38++ cells.
Other gating strategies, including CD38+/CD8 T cells (1000–5000 CD38 molecules/cell, present mostly on a CD45RA+ cell subset of CD8 T cells) would include normal CD8 T cells in the ‘activated’ gate. This would diminish the powerful discrimination between two inherently different CD8 T cell populations that are the activated CD38++, mostly CD45R0+, CD8 T cells and the normal CD38+, mostly CD45RA+, CD8 T cells.
Associations between pre-therapy (baseline) variables were assessed using Spearman's rank correlations. Virological and immunological parameters were summarized by medians and IQR. Changes in immunological parameters between time-points were assessed using the Wilcoxon signed-rank test for paired data. Median changes in VL were estimated using the Kaplan–Meier survival analysis technique in order to account for the limit of detection of the assay. The association of baseline CD8+/CD38++ T cells with time to reach VL of less than 50 copes/ml was assessed using a Cox proportional hazards regression model. The associations of: (i) CD8+/CD38++ T cell count during follow-up with the time to first VL of less than 50 copes/ml; and (ii) VL during follow-up with the time to first CD8+/CD38++ T cell count of less than 20 cells/mm3 were assessed using Cox regression models with time-updated covariates, taking the most recent value (usually measured on the same day). Both VL and the CD8+/CD38++ T cell count were logged before inclusion as covariates in Cox regression models. Results from Cox regression models are presented as hazard ratios with 95% confidence intervals (95% CI). All patients who remained under follow-up were included in analyses, irrespective of whether they had stopped or changed antiretroviral therapy. Patients who withdrew from follow-up prematurely were included up to the point of withdrawal.
Correlations at baseline
Of the 148 patients recruited to Quest, full baseline virological and immunological data, including CD8+/CD38++ T cell values, were available for 131 patients recruited from clinics in Europe and Australia. Of these 131 patients (Table 1), 110 were followed-up for at least 28 weeks after HAART initiation; the remaining 21 patients withdrew from the study before week 28. The median pre-therapy CD4 cell count was 450 cells/mm3 (IQR 309, 601; range 113–1295). The median CD8 cell count was 917 cells/mm3 (IQR 581, 1471; range 115–7218) and CD8+/CD38++ T cell count was 461 cells/mm3 (IQR; 216, 974; range, 14–6708). Median VL was 5.45 log copies/ml (IQR 4.8, 5.9; range 2.1–7.6).
When the correlation between virological and immunological parameters was studied at baseline (Table 2) no correlation was found between VL and CD8+/CD38++ T cell counts (r = 0.14;P = 0.11), nor between the CD8+/CD38++ T cell count and the CD4 T cell numbers. However, VL did show a significant inverse correlation with the CD4 T cell count, linking viral replication with CD4 T cell depletion at PHI (r = 0.29;P < 0.001).
Changes in lymphocyte subsets
After the initiation of therapy, CD8+/CD38++ T cell counts declined markedly in peripheral blood (Fig. 2). Within the first 2 weeks of starting therapy, the median level fell from 461 cells/mm3 to 219 cells/mm3 (n = 115) [median change from baseline −261 cells (IQR −668, −44;P < 0.001)], by week 4 (n = 118) the median CD8+/CD38++ cell count was 125 CD8+/CD38++ cells (IQR 78, 308) [median change of −298 cells (IQR −738, −73)]. The median level had fallen to 78 cells/mm3 (n = 105) by week 12 [median change from baseline −420 cells/mm3 (IQR −858, −113)], and to 47 cells/mm3 (IQR 26, 83; n = 92) by week 28 [median change from baseline −436 cells/mm3 (IQR −876, −150)]. The CD8+/CD38++ cell curve showed a two-phase decline: an exponential fall during the first 4 weeks of therapy [median of individual rates of decline 9.9 cells/day (IQR 2.6, 26.8)] followed by a more gradual decline from week 4 to week 28 [median rate of decline 0.40 cells/day (IQR 0.0, 0.97)]. The numbers of patients with a CD8+/CD38++ T cell count within the normal limit (< 20 cells/mm3) at baseline and at weeks 2, 4, 12 and 28 were two out of 131 (1.5%), none, four out of 118 (3.4%), seven out of 105 (6.7%) and 15 out of 92 (16.3%), respectively.
The drop in CD8+/CD38++ T cell counts during the first 4 weeks of treatment coincided with a less marked change in total CD8 T cells by a median of −251 cells (IQR −785, 79) to 660 cells/mm3 (IQR 492, 880; n = 118). However, at week 28 the median CD8 cell counts were relatively unchanged at 695 cells/mm3 (n = 92) [median change from baseline −212 (IQR −746, 102);P < 0.001], indicating that CD8 T cell counts did not continue to decrease between weeks 4 and 28, in contrast to the CD8+/CD38++ T cell counts (Fig. 2). A relatively rapid increase in CD4 T cell counts occurred during treatment in the first 4 weeks of therapy, with a median increase of 121 cells/mm3 (IQR −11, 241;P < 0.001) that reached 211 cells/mm3 (IQR 61, 375) and a median value of 693 cells/mm3 (IQR 521, 868) by week 28.
Correlations between the decline in viral load and other parameters
During the first 4 weeks, VL levels fell by a median of −2.22 (IQR −2.88, −1.77) log copies/ml (adjusted for assay limit < 50 copies/ml) to 3.14 log copies/ml; n = 123. By week 28, VL levels had decreased further to a median of 1.70 copies/ml (n = 93), a median 5.57 log decline from baseline. The continuing marked changes in VL thus mirrored the decline of CD8+/CD38++ cells but not the alterations of total CD8 cell counts (Fig. 2).
Of all the 131 patients who started HAART, 89 patients recorded at least one VL measurement of less than 50 copes/ml during 28 weeks of follow-up. Baseline VL was strongly predictive of time to first VL of less than 50 copes/ml (viral suppression); hazard ratio 1.52 (95% CI 1.18, 1.96, P = 0.001) for every 1 log lower VL. There was also evidence that the baseline CD8+/CD38++ cell count was an independent predictor of time to viral suppression; hazard ratio 1.52 (95% CI 0.96, 2.38), for every 1 log lower baseline CD8+/CD38++ cell count, adjusted for baseline VL, P = 0.072.
During the first 28 weeks after HAART initiation, there was a close relationship between changes in VL and changes in CD8+/CD38++ cell count. When fitting the latest VL assessment as a time-updated covariate in a Cox model of the time to reach a CD8+/CD38++ T cell count of less than 20 cells/mm3 (that occurred in 31 patients), there was a hazard ratio of 2.99 (95% CI 1.35, 6.64) per 1 log lower VL (P = 0.007). Conversely, the latest CD8+/CD38++ T cell count was also strongly associated with the time to a VL of less than 50 copies/ml: a hazard ratio of 3.38 (95% CI 2.01, 5.70) for every 1 log lower CD8+/CD38++ T cell count (P < 0.001).
Changes in CD8+/CD38++ T cell counts were examined in 61 individuals who achieved and maintained a VL below 50 copies/ml, and who had at least two assessments of CD8+/CD38++ T cell counts during the time of virological suppression. Of these 61 patients, 41 (67.2%) experienced a further fall in CD8+/CD38++ T cells, whereas changes in VL were undetectable, with counts remaining below 50 copies/ml. There was a median change in cell numbers of −18 cells/mm3 (IQR −42, 9;P < 0.001) from the first available CD8+/CD38++ T cell count to the last, implying that reductions in CD8+/CD38++ T cells had continued after the plasma VL had become undetectable.
In addition to VL, PHI can be characterized by marked changes in lymphocyte subpopulations and their activation status [7,34]. In the present study, we confirmed previous observations in PHI that have documented marked CD8 lymphocytosis and CD8 activation, determined as an increase in the number of CD8/CD38+ T cells [3,4,7,30]. In other viral infections, CD8 lymphocytosis [35,36], and the upregulation of CD38 expression on CD8 cells are also observed, sometimes even at levels higher than those seen in HIV. With the Epstein–Barr virus (EBV), the average CD8 cell values reached almost 4000 cells/mm3 . Such a response normalizes naturally, with a gradual decline in immune activation once immune control of the acute phase of EBV infection is achieved .
Unlike EBV infection, in which latency is established after immune resolution, the passage from the acute to the chronic phase of HIV infection is characterized by the persistence of immune activation. This occurs despite a decrease in the levels of plasma viraemia [2,7,9–11,38]. In contrast to EBV infection, the persistence of high CD8 cell counts and of activated CD8 T cells in HIV infection is thought to reflect the failure of the immune system to suppress viral replication fully . The degree of immune activation further increases with the progression of HIV infection to the later stages of symptomatic HIV disease [10,11,13,17,20,24].
In our cohort of 131 PHI patients, we found high numbers of circulating, activated CD8+/CD38++ T cells at baseline (median of 461 cells/mm3) in agreement with previous observations [22,39,40]. The assay that we employed was standardized to quantitate CD8 immune activation above normal levels by selectively counting activated CD8+/CD38++ T cells that are virtually absent (< 20 cells/mm3) in the blood of healthy HIV-negative individuals (Fig. 1). This elevation of the absolute numbers of CD8+/CD38++ T cell counts in the blood of patients who present at the time of PHI apparently reflects a vigorous immune response to HIV using an assay that effectively discriminates between activated (CD38++) and normal (CD38− and CD38+) CD8 T cell populations.
Perhaps the most significant observations of the study were that viral replication was effectively inhibited with a potent HAART combination during PHI, and that this decrease in VL was mirrored by a profound and rapid decline in the CD8+/CD38++ T cells. The number of cells decreased to approximately one sixth of the pre-therapy value by week 12 of HAART intervention. In the majority of patients, the VL was no longer detectable (< 50 copies/ml) by week 28, and the CD8+/CD38++ T cell values were fully normalized in 16% of our cohort, and showed only moderate elevation above the levels in healthy HIV-negative individuals. Indeed, we observed a continued decline in median CD8+/CD38++ T cell counts after VL levels had declined below 50 copies/ml. Consequently, a longer follow-up of this cohort is warranted to assess whether a sustained period of antiviral therapy can fully reverse T cell activation, as defined by CD8+/CD38++ cell counts, in patients who respond to treatment.
It is relevant to the lack of correlation between VL and CD8+/CD38++ cell counts at baseline to consider the absence of a steady state of the VL at this very early stage of infection. Changes in VL appeared to precede CD38++ cell alterations. In approximately a fifth of our patients, the CD8+/CD38++ cell counts were still on the increase and peaked after treatment initiation, whereas VL was invariably highest at presentation. At weeks 2 and 4 after treatment initiation, the correlation between VL and CD8+/CD38++ cell counts appeared to strengthen, increasing from r = 0.37 (P < 0.001, n = 111) to r = 0.43 (P < 0.001, n = 117).
Unfortunately, the protocol for the present study did not include the concomitant investigation of an untreated PHI control group. However, in a small historical control cohort of untreated patients (n = 9) at weeks 20 and 28, estimated time-points after seroconversion, the average level of CD8+/CD38++ cells remained 222 and 181, respectively, and we were not able to find a normalization of CD8+/CD38++ cell numbers. in any of these patients. Furthermore, in six QUEST patients who discontinued therapy after achieving viral suppression, we observed a simultaneous increase in CD8+/CD38++ cell counts and VL after treatment cessation. These data suggest that, in the absence of therapy, increased levels of CD8+/CD38++ cells indeed persist during the passage from acute to chronic infection.
The marked rate at which CD8+/CD38++ T cells disappear from the peripheral blood after viral inhibition, with a halving in numbers within the first 2 weeks of treatment, has implications for understanding the role of CD8+/CD38++ T cells. During the first 2 weeks, the rapid decline in VL and CD8+/CD38++ T cell counts could not be wholly attributed to therapy. Such cells undergo rapid apoptosis or exit from the peripheral blood [41,42]. The parallel loss of total CD8 and CD8+/CD38++ T cell numbers during this early phase would also be consistent with CD8+/CD38++ T cell death or exit from the bloodstream to tissues. This agrees with our findings in the historical control cohort, who showed a similar decline to week 4. However, by week 4, the decrease in CD8+/CD38++ T cell counts was already more profound in the treated as opposed to the untreated patients. Furthermore, at 28 weeks, the decline in CD8+/CD38++ T cells was far more pronounced than that of the total CD8 T cells, implying that, in the absence of further viral stimulation, a significant number of CD8+/CD38++ T cells must have differentiated or downregulated their high CD38++ cell phenotype .
Although VL and CD8+/CD38++ T cell levels were not closely related at baseline, an intimate relationship developed between these parameters during therapy. Once treated, those individuals with the lowest VL values at follow-up were more likely to reach a CD8+/CD38++ T cell count of less than 20 cells/mm3. CD8+/CD38++ T cell counts may represent a sensitive measure of residual viral replication during therapy, and may provide a predictive parameter for response to therapy, defined as VL falling below 50 copies/ml.
Further studies are needed to assess the sensitivity of CD8+/CD38++ cell as a marker of low-level viral replication. As the CD8+/CD38++ cell assay is simple but not HIV specific, it may be employed as an early warning signal in three related areas. First, a study would benefit from a coordinated use of this simple flow cytometric assay, and particularly sensitive virological methods of HIV detection in the blood down to a level of 3 copies/ml . Second, this cellular test may also reflect the existence of viral reservoirs in other body compartments, such as the lymph nodes, at a time when VL in the blood remains undetectable as suggested by others . Finally, the early signs of immune activation might provide the clinician with a warning of patient non-compliance or viral rebound after the cessation of HAART or during its intermittent use.
The QUEST study group wishes to thank all the QUEST scientists for their advice and work, and physicians and nurses for patient referral and clinical care, and the patients for their participation, cooperation and dedication to this study.
We would also like to thank the local study monitors in Australia (M Haberl, J Young), Belgium (D Luyts), Canada (S Pratt), Denmark (R Balschmidt), France (JM Vauthier), Germany (M Sikora), Italy (C Gussetti, CM Anghileri, V Piva), Sweden (G Larrson), Switzerland (I Schauwecker, E Gremlich, C Python) and the UK (P Humphreys). Special mention also goes to the lead investigators in Australia (D Baker, M Bloch, D Smith, P Cunningham, D Cooper, R Finlayson), Belgium (N Clumeck), Canada (J Montaner, C Tsoukas), Denmark (L Mathiesen), France (PM Girard, J Modai, F Raffi, Saitmt), Germany (S Staszewski, H Stellbrink), Italy (A Lazzarin, G Tambussi), Sweden (H Gaines), Switzerland (M Battergay, K Wolfe, L Perin, P Vernazza, R Weber) and the UK (MJ Fisher, B Gazzard, D Hawkins, M Tyrer, M Youle, M Kahan and M Johnson), as well as the Quest Team at GlaxoSmithKline, UK (V Mallet, S Turkish, O Fortes, H Maseruka and H McDade) and Roche Molecular Systems (B Dale and A Capt).
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