Daniel, Volker MD*; Naujokat, Cord MD*; Sadeghi, Mahmoud MD*; Zimmermann, Rainer MD†; Huth-Kühne, Angela MD†; Opelz, Gerhard MD*
Decreased numbers and dysfunctions of precursor dendritic cell (pDC) subsets, T regulatory (Treg) cells, T suppressor (Ts) cells, and interleukin (IL)-7 receptor/CD127-expressing CD4+ and CD8+ lymphocytes were reported in HIV-infected individuals, and it was speculated that abnormalities of these cell populations affect disease progression.1-26
Numeric and functional dendritic cell (DC) deficiencies were associated with high HIV-1 viral load and low CD4+ peripheral blood lymphocyte (PBL) counts and could be restored in vivo by highly active antiretroviral therapy (HAART) and in vitro by neutralization of IL-10 and depletion of CD4+CD25+ T lymphocytes.1-12 In a previous study, we investigated IL-10- and IL-12-producing monocyte-derived CD11c+CD83+CD40+ DCs in the blood of long-term HIV-infected patients with hemophilia. Similar DC subsets had been studied by Chang et al.27 Our investigation revealed that IL-12+ DCs were associated with activation of CD8+DR+ lymphocytes and that IL-10+ DCs were associated with formation of IgG against autologous CD4+ blood lymphocytes.28
Forkehead/winged helix transcription factor (Foxp) 3 expression was decreased in circulating T cells from untreated patients but normalized after initiation of antiretroviral treatment.13 A fraction of HIV-positive individuals exhibiting a low percentage of CD4+ cells and increased levels of activated T cells showed greatly reduced levels of CD4+CD25+Foxp3+ T cells, suggesting disruption of Treg cells during HIV infection.14-16 The Treg cell number was strongly correlated with CD4+ and CD8+ T-cell activation.15 Targeting and disruption of the T-cell regulatory system by HIV may contribute to hyperactivation of conventional T cells, a characteristic of HIV disease progression. In multivariate modeling, the relation between Treg cell depletion and CD4+ T-cell activation was stronger than any other factor examined, including viral load and absolute CD4+ lymphocyte count.15
During HIV infection, CD8+ T cells lack the costimulatory molecule CD28 increase in number and proportion. This accumulation was associated with disease activity and CD8+ T-cell dysfunction.18 CD8+ Ts cells inhibited antigen-specific CD4+ T-cell proliferation and cellular cytotoxicity through secretion of cytokines such as interferon (IFN)-γ, IL-6, and IL-10. CD8+CD28− T lymphocytes from HIV-1-infected patients secreted factors that induced endothelial cell proliferation and acquisition of cell features typical for Kaposi sarcoma.20
The IL-7 receptor (CD127) on CD4+ and CD8+ lymphocytes of HIV-infected patients was reported to be dysfunctional, and it was speculated that this may be attributable to abnormal activation of the immune system and contribute to the reduced CD4 cell count and the altered function of the CD8 compartment in HIV-infected patients.21-23 IL-7Rα expression on CD4+ T lymphocytes decreased with HIV disease progression and was inversely correlated with immune activation.24 Loss of IL-7Rα was associated with CD4+ T-cell depletion, high IL-7 levels, and CD28 downregulation.25 Others reported that loss of CD127 expression defines an expansion of CD8+ T cells that show phenotypic and functional features of effector T cells.26
Although abnormal in patients with advanced disease, the contribution of these immune parameters to HIV disease progression is unclear. This study provides information concerning regulatory components of the immune system in patients who developed severe immunologic abnormalities during the pre-HAART era29-32 and currently show stable CD4+ and CD8+ PBL counts and a stable HIV-1 viral load more than 25 years after HIV infection and 10 years after initiation of HAART. In contrast to other studies, we analyzed, in addition to absolute cell counts, the proportion of cytokine- and/or Foxp3-expressing cells of a particular DC or T-cell PBL subset with regulatory phenotype to assess the activation status of that particular cell subset. We studied (1) whether stable long-term HIV-infected patients with hemophilia showed differences as compared with healthy controls; (2) whether HIV-positive patients with a detectable HIV-1 viral load showed differences as compared with HIV-positive patients with an undetectable HIV-1 viral load; and (3) whether clinically relevant markers of disease progression, such as HIV-1 viral load and CD4+ and CD8+ blood lymphocyte counts, were associated with other immune parameters.
MATERIAL AND METHODS
Patients and Healthy Controls
Forty-one consecutive long-term HIV-infected patients with hemophilia and 110 consecutive healthy controls were studied. The patients were infected with HIV in the early 1980s by virus-contaminated clotting factor products and are part of an ongoing long-term study. Since the end of the year 1996, the patients have been treated with HAART consisting of combinations of nucleoside and nonnucleoside analogues and protease inhibitors. We have studied CD4+ PBL counts in our patients with hemophilia since the year 1982. During the last 25 years, all patients showed periods of disease progression. During disease progression, 7 of the 41 patients had CD4+ PBL counts of 0 to 5 cells/μL, 1 had a CD4+ PBL count of 6 to 10 cells/μL, 13 had CD4+ PBL counts of 11 to 50 cells/μL, 5 had CD4+ PBL counts of 51 to 100 cells/μL, 7 had CD4+ PBL counts of 101 to 150 cells/μL, 5 had CD4+ PBL counts of 151 to 200 cells/μL, and 3 had CD4+ PBL counts >200 cells/μL (Table 1). Twenty-one patients exhibited the lowest CD4+ PBL counts during the pre-HAART era, and 20 patients exhibited the lowest CD4+ PBL counts during the HAART era. CD4+ PBL counts increased in all patients after initiation of HAART or after alterations in the HAART regimen. The control group consisted of healthy blood donors and laboratory staff. Fifty-four of the 110 controls were female, whereas all 41 patients were male. The patients had a mean age of 39.8 ± 8.6 years (range: 23 to 63 years), and the controls had a mean age range of 28.4 ± 9.0 years (range: 19 to 64 years). All patients and controls gave informed consent for the tests performed within this study. The study was approved by the local ethical committee and conducted in adherence with the Declaration of Helsinki.
Determination of IL-10+ or IL-12+ CD11c+CD83+CD40+ DCs
Preparation of Assays
IL-10+ DCs and IL-12+ DCs were determined in freshly obtained heparinized whole blood using 4-color fluorescence flow cytometry as described previously.33 Three tubes were prepared for each test. Tube A served as an isotype control, tube B for the determination of IL-12+ DCs, and tube C for the determination of IL-10+ DCs. Twenty microliters of mouse-IgG1/fluorescein isothiocyanate (FITC; BD Biosciences, Heidelberg, Germany), 20 μL of rat-IgG1/phycoerythrin (PE; BD Biosciences), 10 μL of mouse-IgG2b/allophycocyanin (APC; BD Biosciences), and 5 μL of mouse-IgG1/Cy-Chrome (BD Biosciences) were pipetted into tube A. Tubes B and C contained 20 μL of CD83/Cy-Chrome (BD Biosciences), 5 μL of CD11c/APCs (BD Biosciences), and 20 μL of CD40/FITC (BD Biosciences). One hundred microliters of freshly obtained heparinized whole blood was added to each tube, and the tubes were vortexed and incubated for 30 minutes at 22°C in the dark. Thereafter, 2 mL of fluorescent-activated cell sorting (FACS) lysing solution (BD Biosciences; diluted 1:10 with aqua destillata [aqua dest]) was pipetted into each tube, and the tubes were vortexed, incubated for 10 minutes at room temperature in the dark, and centrifuged at 200g for 8 minutes The supernatant was removed, 2 mL of phosphate-buffered saline (PBS) (GIBCO, Grand Island, NY) containing 0.1% NaN3 and 1% fetal calf serum was added, and the tubes were centrifuged at 200g for 8 minutes The supernatant was removed, and the cell membranes were permeabilized by adding 500 μL of Perm2 solution (BD Biosciences; diluted 1:10 with aqua dest.). The tubes were incubated for 10 minutes at 22°C in the dark, 2 mL of PBS was added, and the tubes were centrifuged at 200 g for 8 minutes. The supernatant was removed, and 20 μL of mouse-antihuman-IL-12/PE (BD Biosciences) was added to tube B and 5 μL of rat-antihuman-IL-10/PE (BD) was added to tube C. After incubation for another 30 minutes at 22°C in the dark, 2 mL of PBS was added to tubes B and C, the tubes were centrifuged at 200 g for 8 minutes, and the supernatant was removed. Five hundred microliters of formaldehyde solution was added to each tube, and the assays were analyzed immediately using a FACSCalibur flow cytometer (BD Biosciences) (Figs. 1A, B).
Determination of Lineage-Negative HLA-DR+CD11c+CD123− pDC1 and Lineage-Negative HLA-DR+CD11c−CD123+ pDC2
Preparation of Assays
DC subsets were determined in freshly obtained heparinized whole blood using 4-color fluorescence flow cytometry. Twenty microliters of FITC-, PE-, or PE-Cy5-labeled mouse-IgG1 (BD Biosciences) and 20 μL of PE- or APC-coupled mouse-IgG2b were used as isotype controls and were pipetted into tube A. Tube B contained monoclonal antibodies against lineage (10 μL of CD3/PE, 10 μL of CD14/PE, 10 μL of CD16/PE, and 10 μL of CD19/PE), CD11c (APC, 5 μL; BD Biosciences), human leukocyte antigen D-related (HLA-DR), human leukocyte antigen DP (HLA-DP), human leukocyte antigen DQ (HLA-DQ) (FITC, 20 μL; BD Biosciences), and CD123 (PE-Cy5, 20 μL; BD Biosciences). One hundred microliters of freshly obtained heparinized whole blood was added to each tube and incubated for 30 minutes at 22°C. Two milliliters of FACS lysing solution (BD Biosciences; diluted 1:10 with aqua dest.) was pipetted into each tube, incubated for 10 minutes, and centrifuged at 200 g for 8 minutes The supernatant was removed, 2 mL of PBS (Gibco) containing 0.1% NaN3 and 1% fetal calf serum was added, and the tubes were centrifuged again. Five hundred microliters of PBS was added to each tube, and the assays were analyzed immediately using a FACSCalibur flow cytometer (BD Biosciences) (see Figs. 1A, C).
Determination of IL-12-Producing HLA-DR+CD11c+CD123− DCs and IL-10-Producing HLA-DR+CD11c−CD123+ DCs
To study IL-12 or IL-10 production in DCs with a phenotype similar to pDC1 or pDC2, we determined in a separate assay HLA-(DR, DP, DQ)+CD11c+CD123−IL-12+ (similar to pDC1) and HLA-(DR, DP, DQ)+CD11c−CD123+IL-10+ (similar to pDC2) DC subsets (see Figs. 1A, D).
Determination of PBLs With Th1, Th2, Treg, or Ts Phenotype
PBLs with Th1, Th2, Treg, or Ts phenotype were determined using 4-color fluorescence flow cytometry. Isotype control consisted of each 20 μL of FITC-, PE-, PerCP-, or APC-coupled mouse- and rat-IgG1 (BD Biosciences). For the determination of CD3+CD4+CD25+ and CD3+CD8+CD28− lymphocytes, the monoclonal antibodies CD3/FITC (20 μL; BD Biosciences), CD4/PerCP (20 μL; BD Biosciences), CD8/peridinin chlorophyll protein (PerCP; 20 μL; BD Biosciences), CD25/APC (5 μL; BD Biosciences), and CD28/APC (20 μL; BD Biosciences) were used. Intracellular cytokine production was measured using the monoclonal antibodies IL-2/PE (20 μL; BD Biosciences), IL-12/PE (20 μL; BD Biosciences), IFNγ/PE (20 μL; BD Biosciences), IL-4/PE (5 μL of a 1:100 dilution; BD Biosciences), and IL-10/PE (5 μL of a 1:100 dilution; BD Biosciences). Intracellular Foxp3 expression was determined using Foxp3/PE (20 μL; NatuTec, Frankfurt, Germany) monoclonal antibody. In addition, IL-7R expression on CD3+CD4+ and CD3+CD8+ lymphocytes was investigated using anti-CD127/PE (20 μL; BD Biosciences). Dot plots of the isotype controls were adjusted to approximately 0.1% single- or double-stained cells, respectively. This standard gate setting was used for subsequent measurement of all patient and control samples. A total of 100,000 events were analyzed.
Determination of Lymphocyte Subpopulations
CD3+CD16−CD19−, CD3−CD16+CD19−, CD3−CD16−CD19+, CD3+CD4+CD8−, and CD3+CD4−CD8+ lymphocyte subsets were defined in whole blood using triple-fluorescence flow cytometry and antibodies from Becton Dickinson/Pharmingen (BD Biosciences, Heidelberg, Germany).
Determination of Plasma Cytokines, Soluble Cytokine Receptors, and Soluble Cytokine Receptor Antagonists
Plasma soluble IL-1 receptor antagonist (sIL-1RA), IL-2, sIL-2R, IL-3, IL-4, IL-6, sIL-6R, IL-10, transforming-growth-factor-β2 (TGF-β2), IFNγ, and tumor necrosis factor-α (TNFα) were determined by enzyme-linked immunosorbent assay (ELISA). sIL-1RA, IL-2, sIL-2R, IL-3, IL-4, IL-6, sIL-6R, IL-10, TGF-β2, and TNFα were measured using Quantikine kits (R&D Systems, Wiesbaden, Germany), and IFNγ was tested using HBT kits (Holland biotechnology BV, Biermann, Bad Nauheim, Germany). Plasma was snap-frozen within 2 hours after the blood was drawn and stored at −30°C until testing.
Plasma neopterin was measured with the Neopterin-ELItest assay (Brahms Diagnostics, Berlin, Germany). Based on control measurements in 70 healthy individuals, a plasma neopterin level >15 nmol/L was considered abnormally high.
Determination of HIV-1 RNA Copies in Plasma
HIV-1 RNA was measured using the NucliSens HIV-1 QT kit (bio Merieux, Nürtingen, Germany) according to the manufacturer's instructions. Sensitivity of the assay is 20 copies using samples of 1 mL of plasma.
The Wilcoxon rank sum test and Spearman rank correlation test were used for statistical analysis applying the Statistical Program for the Social Sciences (SPSS, Chicago, IL). Adjustment for multiple testing was done according to the method of Bonferroni. P values <0.01 were considered significant and are printed in bold in the tables.
Current and Previous CD4+/CD8+ PBL Counts and HIV-1 Viral Load
Most patients had stable CD4+ and CD8+ PBL counts and a stable HIV-1 viral load for the past 12 months. Only 6 (15%) of the 41 patients had a current CD4+ lymphocyte count <155 cells/μL (mean − 2 SDs of healthy controls). Twelve patients (29%) had CD8+ PBL counts >1024 cells/μL (mean + 2 SDs of healthy controls). Eighteen patients (44%) had a viral load <20 (not detectable) HIV-1 RNA copies/mL in the plasma, 13 (32%) had a viral load of 20 to 1000 HIV-1 RNA copies/mL, and 10 (24%) had a viral load of 1200 to 120,000 HIV-1 RNA copies/mL (see Table 1). Patients were called “clinically stable long-term HIV-infected patients with hemophilia on HAART” because 25 patients showed an increasing CD4+ PBL count and 15 patients showed a decreasing CD4+ PBL count during the past 12 months of the observation period. Only 5 of the 15 patients with decreasing CD4+ PBL counts had a CD4+ PBL reduction of >30%. In 1 patient, there was no CD4+ PBL count available 12 months before the current CD4+ PBL count. The data indicate that most of our patients with hemophilia had stable CD4+ PBL counts 25 years after infection with HIV and 10 years after initiation of HAART.
As shown in Table 1, 21 of the 41 patients exhibited the lowest CD4+ PBL counts during the pre-HAART era, most of them during the years 1995 and 1996. Twenty patients showed disease progression during the HAART era, most of them during the years 2000 and 2003. CD4+ PBL counts increased in all patients after initiation of HAART or after alterations in the HAART regimen. These data indicate that CD4+ PBL increases seem to be associated with HAART. We published similar observations in the year 1999.34
Relation of CD4+, CD8+, and CD19+ PBL Counts to HIV-1 Viral Load
Patients had lower mean CD4+ PBL counts (P < 0.001) and, interestingly, higher mean CD8+ PBL counts (P = 0.006) than healthy controls (Table 2). CD4+ and CD8+ PBL counts were similar in HIV-positive patients with detectable or undetectable viral loads (P = not significant [ns]) and were not associated with retroviral load (P = ns) (data not shown). CD8+ PBL counts were associated with CD4+ PBL counts (r = 0.433, P = 0.005; Table 3) and, in addition, with CD3+CD8+CD28−IL-12+ PBL proportions (r = 0.596, P < 0.001; see Table 3), suggesting a dependence of high CD8+ PBL counts on high CD4+ PBL numbers and on high Th1 cytokine-producing CD8+CD28− T lymphocytes.
CD19+ B-lymphocyte counts were similar in HIV-positive patients and healthy controls (P = ns; see Table 2) but were lower in patients with a detectable viral load than in patients with an undetectable viral load (P = 0.002; Table 4; Fig. 2A) and negatively associated with HIV-1 viral load in the total of 41 HIV-positive patients (r = −0.423, P = 0.006; see Table 3). Apparently, CD19+ B-lymphocyte levels were related to retroviral load; however, 95% of the patients had normal B-cell counts (mean ± 2 SDs in controls; see Fig. 2A).
Absolute Numbers of DC- and T-PBL Subsets With Regulatory Phenotype
Patients showed higher CD11c+CD83+CD40+IL-12+ (P < 0.001), CD11c+CD83+CD40+IL-10+ (P < 0.001), HLA-DR+CD11c+CD123−IL-12+ (P < 0.001), and HLA-DR+CD11c−CD123+IL-10+ (P = 0.013) counts; similar lineage-negative HLA-DR+CD11c+CD123− (P = ns) counts; and higher lineage-negative HLA-DR+CD11c−CD123+ DC counts (P < 0.001) than healthy controls (see Table 2). Furthermore, they had lower CD3+CD4+CD25+ PBL numbers expressing IL-2 (P < 0.001), IL-4 (P = 0.001), IL-10 (P = 0.005), IL-12 (P < 0.001), IFNγ (P < 0.001), or Foxp3 (P < 0.001). Of CD3+CD8+CD28− PBLs, only those expressing IL-4 (P = 0.001) or Foxp3 (P < 0.001) were higher in HIV-positive patients than in healthy controls (see Table 2). The data suggest an absolute increase of DCs with regulatory phenotype; an absolute decrease of CD4+ T PBLs with Treg, Th1, or Th2 phenotype; and an absolute increase of CD8+ T PBLs with Ts phenotype in our patient cohort.
In addition to the absolute numbers of cells (see Table 2), we studied the proportion of cytokine- and/or Foxp3-expressing cells of a particular DC- or T-PBL subset with regulatory phenotype to assess the activation status of that particular cell subset (Table 5). This might provide additional information with respect to whether activation of a certain cell subset is up- or downregulated in HIV-positive patients compared with healthy controls.
Proportion of Cytokine-Positive/Cytokine Receptor-Positive DC Subsets
HIV-positive patients with hemophilia and healthy controls had similar proportions of IL-12- (P = ns) and IL-10-producing CD11c+CD83+CD40+ DCs (P = ns), IL-12-producing HLA-DR+CD11c+CD123− DCs (P = ns), IL-10-producing HLA-DR+CD11c−CD123+ DCs (P = ns), lineage-negative HLA-DR+CD11c+CD123− pDC1 (P = ns), and lineage-negative HLA-DR+CD11c−CD123+ pDC2 (P = ns) (see Table 5). There was a trend toward higher proportions of DCs with DC1 quality in patients with a detectable viral load than in patients with an undetectable viral load (CD11c+CD83+CD40+IL-12+ DCs: P = 0.016, and HLA-DR+CD11c+CD123−IL-12+ DCs: P = 0.043, respectively), whereas DCs with DC2 quality were similar in both patient groups (P = ns) (Fig. 3). Strong HIV-1 replication activity was associated with high proportions of DC exhibiting DC1 quality (CD11c+CD83+CD40+IL-12+ DCs: r = 0.482, P = 0.001, and HLA-DR+CD11c+CD123−IL-12+ DCs: r = 0.423, P = 0.006, respectively; see Table 3), whereas high CD8+ lymphocyte counts were associated with low proportions of DC showing DC2 quality (HLA-DR+CD11c−CD123+IL-10+ DCs: r = −0.616, P < 0.001, and lineage-negative HLA-DR+CD11c−CD123+ pDC2: r = −0.572, P < 0.001, respectively; see Table 3). CD4+ lymphocyte counts were not associated with DC subsets (P = ns). These findings suggest (1) normal levels of activated DC subsets determined by cytokine production and/or cytokine receptor expression and (2) association of high DC1 with strong retroviral replication and low DC2 with strong CD8+ lymphocyte responses in long-term HIV-infected patients with hemophilia.
Proportion of Th1/Th2 Cytokine-Producing CD3+CD4+CD25+ and CD3+CD8+CD28− PBLs
HIV-positive patients had higher proportions of IL-4- (P = 0.005) and IL-10- (P = 0.004) producing CD3+CD4+CD25+ blood lymphocytes but lower proportions of IL-2- (P < 0.001), IL-12- (P < 0.001) and IFNγ- (P < 0.001) producing CD3+CD4+CD25+ T lymphocytes than healthy controls (see Table 5). Proportions of these T lymphocyte subsets were similar in HIV-positive patients with detectable and undetectable viral loads (P = ns) and were neither related to retroviral load nor to CD4+ or CD8+ lymphocyte counts (P = ns). Of the Th1/Th2 cytokine-producing CD3+CD8+CD28− PBL subsets, only CD3+CD8+CD28−IL-4+ PBLs were higher in patients than in healthy controls (P = 0.012; see Table 5). Stable long-term HIV-infected patients with hemophilia seem to exhibit upregulation of Th2 and downregulation of Th1 PBLs that are independent of HIV replication activity and do not affect CD4+ or CD8+ lymphocyte counts. In addition, they show a relative and, as already shown in Table 2, absolute increase of CD3+CD8+CD28−IL-4+ PBLs (P = 0.001).
Proportion of Foxp3-Expressing CD3+CD4+CD25+ and CD3+CD8+CD28− PBLs
Proportions of circulating Foxp3-expressing CD3+CD4+CD25+ lymphocytes were similar in HIV-positive patients and healthy controls (P = ns; see Table 5) but were significantly higher in patients with a detectable HIV-1 viral load than in patients with an undetectable HIV-1 viral load (P = 0.012; see Table 3; Fig. 4A) and were positively associated with the degree of retroviral load (r = 0.518, P = 0.003; see Table 3). Although this PBL subset with Treg phenotype increases in parallel to retroviral load, 77% of the patients had normal (mean ± 2 SDs in controls) proportions of CD3+CD4+CD25+Foxp3+ PBLs (see Fig. 4A). High CD3+CD4+CD25+Foxp3+ PBLs were borderline associated with low CD8+ T-lymphocyte counts (P = 0.029; see Table 3).
Fifty percent of the patients had CD3+CD8+CD28−Foxp3+ PBLs in the normal range (mean ± 2 SDs in healthy controls). In contrast to CD3+CD4+CD25+Foxp3+ PBLs, the proportion of CD3+CD8+CD28−Foxp3+ PBLs was higher in HIV-positive patients than in healthy controls (P < 0.001; see Table 5). HIV-positive patients with a detectable viral load had higher CD3+CD8+CD28−Foxp3+ PBLs than HIV-positive patients with an undetectable viral load (P = 0.004; see Fig. 4B) or healthy controls (P < 0.001; see Table 4), whereas patients with an undetectable retroviral load had nearly normal CD3+CD8+CD28−Foxp3+ PBL levels (P = 0.055; see Table 4). CD3+CD8+CD28−Foxp3+ PBLs were positively associated with HIV-1 viral load (r = 0.530, P = 0.003; see Table 3) and negatively associated with CD19+ PBL counts (r = −0.494, P = 0.006; see Fig. 2B). Apparently, this PBL fraction with Ts phenotype increases to abnormally high levels in parallel to retroviral load and is associated with lower B lymphocyte counts. Figure 5 shows representative dot plots of Foxp3-expressing CD3+CD4+CD25+ and CD3+CD8+CD28− PBLs in 2 HIV-positive patients with hemophilia.
Proportion of CD127-Expressing CD3+CD4+ and CD3+CD8+ PBLs
HIV-positive patients had lower proportions of CD127+ CD3+CD4+ (P < 0.001) and CD3+CD8+ PBLs than healthy controls (P < 0.001; see Table 5), and the proportions of these 2 PBL subsets were similar in patients with <20 or ≥20 HIV-1 RNA copies/mL of plasma (P = ns, data not shown). Whereas low CD4+ lymphocyte counts were associated with low CD3+CD4+CD127+ PBL proportions (r = 0.382, P = 0.014; see Table 3), low CD8+ lymphocyte counts were not related to low CD127 expression on CD8+ PBLs (Fig. 6). Rather, patients with abnormally high CD8+ PBL counts showed abnormally low CD127 expression on CD8+ PBLs (r = −0.305, P = 0.052).
Compared with healthy controls, HIV-positive patients had higher neopterin (P < 0.001), sIL-1RA (P < 0.001), sIL-2R (P < 0.001), and IL-6 (P = 0.005) plasma levels and similar IL-2, IL-3, IL-4, sIL-6R, IL-10, TNFα, TGF-β2, and IFNγ plasma levels (P = ns; see Table 5). Patients with a detectable viral load showed higher IL-10 plasma levels than patients with an undetectable viral load (P = 0.005; see Table 4), whereas plasma levels of the other cytokines were similar in the 2 patient groups (P = ns, data not shown). HIV-1 viral load was associated with high plasma levels of neopterin (r = 0.417, P = 0.006), sIL-2R (r = 0.424, P = 0.006), and IL-10 (r = 0.395, P = 0.012) (see Table 3). CD4+ and CD8+ lymphocyte counts were not associated with plasma cytokine levels (P = ns). Patients with hemophilia seem to have a strong activation of neopterin- and IL-10-producing cells that is related to retroviral replication and does not affect CD4+ and CD8+ lymphocyte counts (see Fig. 1C).
The goal of this study was to investigate in a group of clinically stable long-term (more than 25 years) HIV-infected patients with hemophilia the regulatory components of the immune system that were previously reported to be associated with disease progression. In difference to the studies cited in the introduction section, our patients were homogeneous with respect to route and duration of infection, antiretroviral treatment, stability of disease, and gender.
Our data do not provide evidence for decreases and/or dysfunctions of DC subsets in clinically stable long-term HIV-infected patients with hemophilia receiving HAART. Only absolute numbers of CD4+ PBLs with Treg, Th1, or Th2 phenotype were strongly decreased in patients, and these decreases can be ascribed to the general CD4+ PBL depletion of the patients. Our data show that there are differences between patients and healthy controls with respect to the activation status of DC and T lymphocytes with regulatory phenotype. The findings presented in this study show that those long-term HIV-infected patients with hemophilia who developed severe immunologic abnormalities during the pre-HAART era29-32 exhibit at least partial reconstitution of immune functions 10 years after initiation of HAART. They have normal levels of activated DC1 and DC2 that seem to react to viral load rises with DC1 increases and seem to support CD8+ PBL responses with DC2 decreases. CD3+CD4+CD25+Foxp3+ and CD3+CD8+CD28−Foxp3+ PBLs show normal levels in patients with undetectable retroviral load. Both subsets increase in parallel to retroviral load and seem to downregulate increased CD8+ T- and CD19+ B-lymphocyte counts in patients with a detectable retroviral load. We speculate that they contribute to the reduction of autoimmunity observed in our patients after initiation of HAART.34 In the pre-HAART era, our patients showed autoimmune phenomena, such as an increased IgG load on circulating CD4+ PBLs that nearly disappeared after initiation of HAART.34 In the present study, only 2 of 41 HIV-positive patients had abnormally increased CD4+IgG+ PBLs of >30%, which was our cutoff for the differentiation of autoantibody-negative and autoantibody-positive patients (data not shown).31 It now seems that the higher CD3+CD4+CD25+Foxp3+ PBLs in patients with a higher viral load might downregulate or even prevent the formation of autoantibodies against CD4+ PBLs. High CD3+CD8+CD28−Foxp3+ PBL proportions were associated with a high viral load and reduction of CD19+ B-lymphocyte counts, and this reduction in B-lymphocyte counts might additionally contribute to downregulation of autoantibody formation.
Interestingly, Th1 and Th2 proportions and the abnormally increased CD8+ PBL counts were not associated with HIV-1 viral load. Continuous administration of pooled clotting factor preparations, in part contaminated with inactivated virus particles, and chronic infections with viruses other than HIV might contribute to CD8+ PBL count increases and Th2 domination. The fact that our HIV-positive patients are able to generate abnormally high CD8+ PBL numbers indicates immunocompetence. We did not measure the HIV-1-specific CD8+ lymphocyte response; however, Betts et al35 reported recently the maintenance of highly functional HIV-specific CD8+ T cells in HIV nonprogressors and an inverse correlation of the HIV-specific T-cell response with retroviral load in progressors. We therefore assume that the increased CD8+ lymphocyte response measured in our HIV-positive patients with hemophilia includes an HIV-specific CD8+ lymphocyte response.
A downregulation of IL-7R on CD4+ and CD8+ lymphocytes in HIV-positive patients with low CD4+ PBL counts was described by several authors.21-26 Signaling by means of IL-7R is essential for T-cell homeostasis and maintenance of T-cell memory. IL-7 plays a major role in T lymphocyte homeostasis and has been proposed as an immune adjuvant for lymphopenic patients.36-38 Whereas in our HIV-positive patients, the proportion of CD3+CD8+CD127+ PBLs was not associated with CD8+ PBL counts, low proportions of CD3+CD4+CD127+ lymphocytes were related to low CD4+ PBL counts, suggesting a contribution of low IL-7R expression on CD4+ lymphocytes to CD4+ lymphopenia. Considering the importance of IL-7 and IL-7R expression for lymphopoiesis, it seems paradoxic that those patients with the highest CD8+ PBL counts showed the lowest proportions of CD127-expressing CD3+CD8+ PBLs (see Fig. 6B). Based on these data, the importance of CD127 expression for CD4+ and CD8+ PBL counts remains unclear.
These results are encouraging because they show that our patients who had developed severe immunologic abnormalities during the pre-HAART era29-32 and currently show stable CD4+ and CD8+ PBL counts and a stable HIV-1 viral load regained immunoregulatory capacity on HAART that might bestow immunocompetence, halt disease progression, and stabilize the clinical condition of these long-term HIV-infected patients.
The authors acknowledge the skillful technical assistance of Roland Seidel, Silja Petersen, Martina-Kutsche-Bauer, Regina Seemuth, Cima Farahmandi, Marion Miltz, and Gabi Schmeckenbecher.
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