Plasmacytoid dendritic cells (pDC) expressing the cell surface markers BDCA-2 and BDCA-4 are the major producers of interferon alpha (IFNα) in humans.1-3 IFNα expression is rapidly upregulated in response to viral infections,4-6 and it orchestrates the innate immune response through the induction of several effector molecules, such as 2′-5′ oligoadenylate synthetase, double-stranded protein kinase, and MxA.7-9 As MxA is only induced by IFNα, it is a useful indicator of IFNα activity.10,11 Moreover, IFNα is of great importance in regulating the adaptive immune response6,12,13 and has antiproliferative activity.14,15
To date, the role of pDC and IFNα during HIV-1 infection still remains unclear. Indeed, HIV-1 disease progression is associated with a decline of pDC from the peripheral compartment, which correlates with high viral load and reduced CD4 cell counts.16 In addition, pDC of patients with HIV-1 produce lower levels of IFNα when challenged in vitro with reference viruses.16-22 However, several studies have documented the chronic production of IFNα in patients HIV-1, which is associated with disease progression.23-26 More recently, Siliciano and colleagues reported the overexpression of type I interferon-regulated genes in activated CD4+ T cells of patients with HIV-1.27 Moreover, these authors found upregulation of cell cycle-associated genes under type I interferon modulation.27
We aimed at clarifying these contrasting results by investigating in greater detail the fate and function of pDC in HIV-1 infection. Thus, we studied the distribution of pDC in paired peripheral blood and lymph node samples from HIV-1-infected individuals at different stages of disease. In addition, we analyzed IFNα and MxA expression levels at steady state, that is, in the absence of any in vitro stimulation or in vitro culture. Our results indicate that pDC of patients with HIV-1 markedly decline in the peripheral compartment but constitutively overexpress IFNα, which represents a hallmark of immune dysfunction in late-stage HIV-1 infection.
All samples were obtained with signed informed consent according to ethical guidelines. Lymph nodes (cervical, axillary, and inguinal) were excised, and peripheral blood was drawn at the University of Hamburg-Eppendorf from 13 HIV-1-positive individuals at different stages of disease. Additionally, peripheral blood of 17 HIV-1-infected individuals at different stages of disease [Centers for Disease Control (CDC) stage A n = 7, CDC stage C n = 10] was collected at the University of Cologne. As control, peripheral blood and tonsils or lymph nodes were collected at the University of Cologne from 10 uninfected individuals. Moreover, peripheral blood samples only were obtained at the University of Cologne from 10 additional healthy HIV-1-negative individuals. In all cases, 50 mL of blood was obtained.
Isolation of Peripheral Blood Mononuclear Cells (PBMC) and Lymphoid Tissue Mononuclear Cells
Lymph nodes and tonsils were placed in normal saline immediately after surgery, and mononuclear cells were dissociated mechanically. Peripheral blood and lymphoid tissue mononuclear cells were isolated by Ficoll centrifugation and stored at −180°C.
Viral Load Measurement
HIV-1 plasma viral load was quantified using Roche Amplicor kits (Roche Diagnostics, Manheim, Germany; limit of detection of 50 copies/mL). For viral load determination in lymph nodes, total RNA was extracted with Trizol from 1 × 106 lymphoid tissue mononuclear cells (LTMC), and HIV-1 RNA copy number was determined with the Roche Amplicor kit.
For surface staining, cells were incubated in the dark for 30 minutes at 4°C with anti-CD123 and anti-BDCA-4 or anti-BDCA-2 antibodies (Miltenyi Biotec, Bergisch Gladbach, Germany). For intracellular IFNα expression, PBMC was first stained with anti-CD123 and anti-BDCA-4 antibodies, fixed and permeabilized with the Cytofix/Cytoperm kit from Becton Dickinson, and then stained with anti-IFNα antibody (PBL; InterferonSource, Piscataway, NJ). As control, we used isotype-matched antibodies labeled with the appropriate fluorochrome (BD Biosciences, Heidelberg, Germany). After staining, the cells were washed with phosphate buffer saline (PBS) and analyzed by flow cytometry (FACSORT) using CellQuest software (both from Becton Dickinson). In all cases, 100,000-300,000 events were acquired corresponding to live mononuclear cells as assessed by forward and side light scatter profile. Interferon alpha expression was determined as mean fluorescence intensity by further gating on the pDC population. Data analysis was performed with the FlowJo software (Tree Star, Ashland, OR).
RNA Extraction and Quantitative Real-Time Polymerase Chain Reaction
Total RNA was extracted using the High Pure RNA Isolation Kit (Roche Diagnostics) and subjected to DNase I treatment. RNA integrity was assessed by denaturing gel electrophoresis (presence of 18S and 28S bands). The same method was also used to isolate RNA from pDC-enriched and pDC-depleted fractions obtained by separation of 0.5-1 × 108 PBMC by positive selection with anti-BDCA-4 microbeads (Miltenyi Biotec, Bergislch Gladbach, Germany). RNAs were reverse transcribed into cDNA by using random hexamers and Superscript II Plus RNase H−Reverse Transcriptase (Invitrogen, Carlsbad, CA). The expression of IFNα and MxA mRNAs was determined from 10 ng samples of cDNA (as estimated from ethidium bromide spot tests) by quantitative real-time polymerase chain reaction (PCR) relative to glyceraldehyde-3-phosphate dehydrogenase using LightCycler FastStart DNA MasterPLUS SYBR Green I (Roche Diagnostics). Relative quantifications were performed by pairwise fixed reallocation randomization tests and corrected for amplification efficiency as evaluated from serial dilutions of cDNA. Repeated runs of the same samples gave a maximal 2%-4% interassay variation, allowing detection of a 2-fold difference in RNA levels at 95% confidence. Linearity of determinations was confirmed for the entire working range of mRNA expression (4-5 log values); failing expression was considered after more than 45 cycles of amplification without increase in fluorescence intensity. The LightCycler protocol consisted of denaturation (95°C for 15 seconds), annealing (60°C for 10 second), elongation (72°C for 20 seconds), and additional melting (85°C for 5 seconds) for 45 cycles. The fourth PCR segment with fluorescence acquisition at elevated temperature was added to eliminate nonspecific fluorescence signals derived by primer dimers or unspecific products. Primer design was done using Primer-3 software. The oligonucleotide primers for IFNα were 5′-TGA AGG ACA GAC ATG ACT TTG G-3′ (sense) and 5′-TCC TTT GTG CTG AAG AGA TTG A-3′ (antisense), yielding a product of 123 bp; the primers for MxA were 5′-ATT TCG GAT GCT TCA GAG GTA G-3′ (sense) and 5′-TAG AGT CAG ATC CGG GAC ATC T-3′ (antisense), yielding a product of 131 bp; the primers for glyceraldehyde-3-phosphate dehydrogenase were 5′-AAA GGG TCA TCA TCT CTG CC-3′ (sense) and 5′-TGA CAA AGT GGT CGT TGA GG-3′ (antisense), yielding a product of 575 bp. Specificity of LightCycler PCR products was assessed by melting curve analysis. For exact length verification, PCR products were separated by agarose gel electrophoresis.
Data were analyzed with Wilcoxon rank-sum test, Wilcoxon matched pairs test, or 1-way analysis of variance test with Holm-Sidak corrections as applicable. All statistical analyses assumed a 2-sided significance level of 0.05. Pearson r was used to describe correlations. Data were summarized using median and interquartile range or mean and standard error of mean (SEM), as indicated in the figure legends. Data analyses were performed with SPSS v12 software (Chicago, IL).
BDCA-4+ CD123+ pDC Are Lost in Peripheral Blood With Disease Progression
Thirty HIV-1-infected individuals at different stages of the disease (CDC stage A n = 13; CDC stage C n = 17; Table 1) were compared with 20 HIV-1-negative individuals for the frequency of circulating pDC. The A, B, and C staging of the CDC classification for HIV-1 infection is based on clinical parameters (opportunistic infections and other manifestations), explaining the high variability of HIV-1 viral load within the 2 patient groups. In addition, only 1 of 13 stage A patients and only 2 of 17 stage C patients included in our study were undergoing antiretroviral therapy at the time of sample collection (Table 1).
Figure 1 shows representative flow cytometry plots obtained by staining PBMC (top row) and LTMC (bottom row) preparations from an HIV-1-negative, an HIV-1-positive stage A, and an HIV-1-positive stage C individual with antibodies directed to BDCA-4 and CD123. We also used BDCA-2 as a second pDC-specific marker to control for possible BDCA-4 modulation in different compartments. Detection of pDC with these 2 direct methods produced essentially identical results (compare pDC frequencies between paired plots).
We extended the analysis with anti-BDCA-4 and anti-CD123 antibodies to paired PBMC and LTMC preparation from all HIV-1-negative and -positive individuals reported in Table 1. As shown in Figure 2A, the frequency of pDC in peripheral blood of HIV-1-positive stage C individuals was markedly reduced compared with HIV-1-negative controls [median (interquartile range)]: HIV-1 negative 0.33% (0.32-0.72); HIV-1 positive stage A 0.34% (0.20-0.37); HIV-1 positive stage C 0.06% (0.04-0.15); stage C/stage A: P = 0.001; stage C/negative controls: P = 0.001. The analysis of pDC in lymph nodes of HIV-1-positive individuals at different stages of disease (stage A n = 6; stage C n = 7) compared with lymphoid tissues of HIV-1-negative individuals (n = 10) did not show a statistically significant difference (P > 0.05) between any of the groups (Fig. 2B): HIV-1 negative controls 0.39% (0.16-1.23); HIV-1 positive stage A 0.42% (0.31-0.63); HIV-1 positive stage C 0.41% (0.36-0.55). Control lymphoid tissues from HIV-1-negative subjects also included tonsils, which are likely to be more activated than lymph nodes due to continuous antigen exposure. However, no significant difference in pDC frequency was observed when comparing exclusively lymph nodes (not shown).
Increased IFNα and MxA mRNA Expression in Peripheral Blood of Patients With HIV-1 Progressing to Disease
The antiviral activity of IFNα depends-among others-on the effector molecule MxA, which continues to be expressed after IFNα is no longer detectable. Peripheral blood and lymphoid tissue mononuclear cells from all HIV-1-positive and -negative individuals analyzed in Figure 2 were tested for IFNα and MxA mRNA expression. Because a recent report suggested that reduced IFNα expression in response to in vitro stimulation does not accurately reflect IFNα expression in vivo,28 we analyzed IFNα and MxA mRNA levels at steady state, that is, in the absence of any stimulation. We found that IFNα mRNA expression levels were 10- and 100-fold higher in peripheral blood of HIV-1-positive stage A and stage C patients, respectively, compared with uninfected individuals (P < 0.05 and P < 0.001, respectively; Fig. 3A). In the same compartment, MxA mRNA expression was 5-fold higher in HIV-1-positive stage A patients and 10-fold higher in stage C patients compared with uninfected controls (Fig. 3B). In lymphoid tissues, IFNα and MxA mRNA levels did not significantly differ between HIV-1-positive stage C patients and uninfected individuals (P > 0.05). However, stage A patients displayed 10-fold lower levels of IFNα and MxA mRNA compared with the other 2 groups (P < 0.05; Figs. 3C and D). Again, comparing IFNα and MxA mRNA levels exclusively in lymph nodes of HIV-1-negative and -positive donors produced identical results (not shown). Therefore, despite declining pDC counts, PBMC of HIV-1 patients display a marked upregulation in IFNα and MxA expression.
Increased IFNα mRNA and Protein Expression in Circulating pDC of Patients With HIV-1 Progressing to Disease
Next, we sought to reconcile the apparent paradox of declining pDC counts in the face of increasing IFNα expression in peripheral blood of patients with HIV-1 progressing to disease. We obtained pDC-enriched and pDC-depleted fractions by positive selection from PBMC of HIV-1-infected and uninfected individuals. When the 2 fractions were compared for IFNα and MxA mRNA expression at steady state, the pDC-enriched fraction from PBMC of HIV-1-positive and -negative individuals displayed ∼65-fold higher IFNα mRNA levels compared with their matched pDC-depleted fraction (P < 0.01; Fig. 4A). In addition, both pDC-enriched fractions displayed ∼25-fold higher MxA mRNA levels compared with the matched depleted fraction (P < 0.01; Fig. 4B). These data indicate that pDC account for >95% of the IFNα and MxA mRNA expression in PBMC of HIV-1-positive and -negative individuals. Moreover, the pDC-enriched fraction obtained from HIV-1-positive stage C individuals expressed ∼90-fold higher IFNα mRNA levels compared with the pDC-enriched fraction from HIV-1-negative individuals (P < 0.001; Fig. 4A), which closely approximates the difference observed when comparing total PBMC (Fig. 3A). Altogether, these results indicate that pDC-and not other cell types29,30-are fully responsible for the increased expression of IFNα in PBMC of patients with HIV-1. Because of the limited amount of starting cells and the low frequency of circulating pDC particularly in patients with HIV-1 (≤0.3%), the yield of enriched pDC was insufficient to allow both mRNA extraction and assessment of purity by flow cytometry. However, flow cytometry analyses showed less than 0.02% residual pDC in all pDC-depleted fraction (not shown), supporting the conclusion drawn from reverse transcriptase-PCR analyses that increased IFNα mRNA levels in PBMC of patients with HIV-1 are fully accounted for by pDC.
To confirm the results of mRNA expression also at protein level, we performed intracellular flow cytometry analyses. As shown in Figure 4D, IFNα was detectable at higher levels within pDC of patients with HIV-1 in stage C disease compared with uninfected controls and patients in early stage of the infection. The flow cytometry histogram plots in Figure 4C show intracellular staining with anti-IFNα antibodies obtained with cells from a representative HIV-1 negative and an HIV-1-positive stage C individual. These plots display a single-peak histogram-shifted to the right in the latter compared with the former individual-indicating that the entire pDC population as a whole rather than a subset of cells in the HIV-1-positive subject expresses higher levels of IFNα. Therefore, despite their marked decline, pDC of HIV-1-stage C patients display extremely high levels of IFNα expression in the absence of exogenous stimulation with reference viruses.
Direct Correlation Between Peripheral pDC and CD4+ T-Cell Counts
Increased viral replication may be the cause of declining pDC counts and rising IFNα expression. To address this hypothesis, we conducted statistical analyses seeking a correlation between various laboratory and clinical parameters. There was a weak inverse correlation between plasma HIV-1 mRNA and peripheral pDC counts (r = −0.47, P < 0.05, Pearson r test; Fig. 5A), suggesting that HIV-1 may cause loss of pDC by direct infection and cell killing. However, HIV-1 mRNA plasma levels did not correlate with IFNα and MxA mRNA levels (Figs. 5B and C), indicating that higher IFNα levels are not a direct consequence of increased viral replication. On the contrary, a strong correlation existed between CD4+ T-cell and pDC counts (r = 0.68, P < 0.01, Pearson r test; Fig. 5D) but not between CD4+ T-cell counts and IFNα or MxA mRNA levels (P > 0.05; Figs. 5E and F). Also, no statistically significant correlation emerged when analyses between CD4+ T-cell counts or HIV-1 RNA (on one side) and IFNα or MxA mRNA levels (on the other side) were carried out within the 2 patient groups included in our study (not shown). We found no correlation between IFNα or MxA mRNA levels and frequency of circulating pDC (not shown), which was expected in that IFNα expression on a per-cell basis is not a function of pDC frequency. Finally, we did not find any association between HIV-1 mRNA, pDC counts, and IFNα and MxA mRNA levels in lymphoid tissues (not shown).
HIV-1 disease is associated with a loss of pDC in peripheral blood and with their reduced ability to produce IFNα after in vitro challenge with reference virus.16-18 However, previous studies did not assess whether the decline of pDC in peripheral blood was the consequence of HIV-dependent cytotoxicity, defects in bone marrow output, or redistribution to lymphoid tissues (the major sites of HIV-1 replication during the chronic phase of infection,31-33 as observed in other viral infections.34
Our study represents the first comparative analysis of pDC frequency and function in peripheral blood and lymphoid tissues of HIV-1-infected individuals at different stages of disease. We show that pDC decline in peripheral blood but remain unchanged in lymph nodes of patients with HIV-1 progressing to disease. The total number of pDC in the body includes cells present in peripheral blood, secondary lymphoid tissues, mucosal tissues, and peripheral nonlymphoid tissues. As the blood contains less than 10% of the total body immune cells, a decline of pDC limited to the peripheral blood is likely to impact only in small proportion the total pDC body count and cannot be the measure of generalized systemic depletion. Our results indicate that the frequency of pDC in lymph nodes remains unchanged even in stage C patients, without generalized systemic decline of pDC. This result seems to exclude a deficiency in bone marrow output-which is expected to produce systemic effects-and to place more emphasis on alternative hypotheses. First, we observed a strong correlation between CD4+ T-cell and pDC counts, suggesting that the same HIV-induced mechanism(s) may lead to depletion of both CD4+ T cells and pDC. Second, the decline of circulating pDC may be associated with homing to lymphoid tissues to fight HIV-1 replication.34 However, because we failed to observe increased pDC numbers in lymph node samples of patients with HIV-1, for this hypothesis to be true, the influx of pDC into lymphoid tissues should be balanced by increased pDC death rates due to a homeostatic control mechanism or an HIV-1-mediated killing. Finally, pDC might redistribute to peripheral nonlymphoid tissues responding to opportunistic infections or inflammatory responses. The pronounced decline of circulating pDC in stage C patients (that displays opportunistic infections and other clinical manifestations) but not in stage A patients (that shows higher lymphoid tissue-associated viral replication) seems to put more emphasis on the last hypothesis.
Our results are in contrast with a recent study that found reduced pDC numbers in lymph nodes of HIV-1-infected patients.35 This discrepancy may be the consequence of several methodological differences. In our study, pDC frequency in lymph nodes of patients with HIV-1 was analyzed by BDCA-4 and CD123 staining in comparison to PBMC samples from the same individual and lymphoid tissue samples from control individuals. Moreover, a second direct detection method (BDCA-2 and CD123) allowed to exclude the possibility of modulated BDCA-4 expression in lymph nodes, thus supporting and validating the results obtained in lymph nodes. On the other hand, Biancotto et al35 analyzed pDC numbers in lymph nodes but not paired PBMC samples using an indirect approach (Lin−/CD11c−/HLA-DR+/CD123+) involving a complex series of gating steps and without taking into account the possible effect of cell surface marker modulation. This is particularly relevant considering that Biancotto et al35 studied lymph node samples after in vitro culture, which might lead to higher pDC death rates in HIV-1-positive samples due to insufficient IL-3 expression36 and to altered cell surface marker expression. Although our study employed archived samples, cryopreservation does not affect cellular phenotype.37,38 Finally, the different antiretroviral therapy status of the patient populations in these 2 studies might account for the discrepancy in lymph node-homed pDC frequency. Indeed, there is still much to be learned about the dynamics of pDC in HIV-1 disease and in response to antiretroviral therapy.
Surprisingly, the decline of pDC in peripheral blood of patients with HIV-1 was associated with overexpression of IFNα and MxA in peripheral blood of patients with HIV-1. Previous reports found that pDC of patients with HIV-1 produced suboptimal levels of IFNα in response to in vitro stimulation with CpG and reference viruses and concluded that these cells were functionally impaired.16-22 Here we studied IFNα expression at steady state in the absence of exogenous stimuli, and we demonstrate that circulating pDC of patients with HIV-1 markedly overexpress IFNα and MxA both at the mRNA and protein levels. Our results are in line with-and provide proof to-a recent report suggesting that reduced IFNα expression in response to in vitro stimulation may not reflect impaired function in vivo but rather might represent a negative feedback control mechanism due to persistent exposure to high levels of IFNα in vivo.28 The observation of increased IFNα expression was limited to the peripheral compartment and was not paralleled by similar results in lymph nodes. Previous studies have shown that pDC express IFNα after in vitro exposure to HIV-1,39,40 suggesting that HIV-1 may increase IFNα expression by unstimulated pDC and raise the blood levels of this cytokine. However, we did not find any correlation between HIV-1 RNA levels in blood and IFNα mRNA levels in PBMC. Surprisingly, lymph node samples of HIV-1 stage A patients showed lower IFNα and MxA mRNA levels compared with controls and stage C patients. Although pDC express IFNα after in vitro exposure to HIV-1,39,40 the situation in lymphoid tissues may be different. For instance, HIV-1 gp120 and other proteins trapped in the intercellular space or expressed on cell surface above a critical concentration (such as in the stage A patients in our study) may cause impaired pDC function and decreased IFNα expression. Indeed, Fauci and his colleagues recently showed that monomeric and trimeric gp120 suppress IFNα production by pDC.41
Although IFNα expression is part of the natural immune response, its overproduction in peripheral blood is unusual and may contribute to the progressive immune suppression typical of late-stage HIV-1 disease. Several lines of evidence support this hypothesis. Herbeuval et al42-44 showed that HIV-1 induces IFNα expression and TNF-α-related apoptosis including ligand (TRAIL)-dependent apoptosis, suggesting a role for these cytokines in HIV-1 immunopathogenesis. Abel et al45 showed that during the chronic phase of simian immunodeficiency virus (SIV) infection, higher levels of IFNα mRNA are found in rhesus macaques progressing more rapidly to AIDS. A very recent study analyzed pDC dynamics and IFNα expression in the case of nonpathogenic SIV infection in African green monkeys and showed that: (1) pDC migrate from peripheral blood to lymph nodes during the acute phase of the infection, but then pDC blood levels return to normal without decline during the chronic phase of the infection; (2) before SIV infection, pDC of African green monkeys display moderate expression of IFNα after toll-like receptor (TLR) stimulation in vitro; and (3) SIV-infected African green monkeys produced IFNα during the acute phase but only transiently and at moderate levels.46 Finally, Siliciano and colleagues showed that type I interferons (IFNα and IFNβ) promote a state of immune hyperactivation in patients with HIV-1.27 Altogether, these studies suggest that IFNα might contribute to CD4+ T-cell depletion and immune dysfunction that characterize progression to AIDS.
Previous studies showed that influenza virus induces the expression of MxA and IFNα with similar kinetics; however, MxA mRNA accumulates and persists long after IFNα mRNA levels decline.47 Interestingly, we show that although pDC of patients with HIV-1 expressed 100-fold higher levels of IFNα mRNA compared with uninfected controls, MxA mRNA levels were only 10-fold higher. Thus, our results suggest that IFNα produced in HIV-1-positive individuals may have reduced antiviral properties. This is in line with a previous report showing that in vitro coculture of PBMC with HIV-1-infected monocytes led to the production of IFNα with 20-fold lower antiviral activity compared with equal amounts of recombinant IFNα2b.48 A possible explanation for this observation is that HIV-1 counteracts the antiviral activity of IFNα, as observed for other viruses.49,50 For instance, the HIV-1 Tat protein associates with protein kinase R, an IFNα antiviral effector molecule, and inhibits its activity.51,52 Alternatively, pDC of patients with HIV-1 progressing to disease express IFNα subtypes with weaker antiviral activity.53 Several authors have also described a heavily deglycosylated, acid labile form of IFNα in late-stage HIV-1 disease, which is found in certain forms of autoimmunity such as systemic lupus erythematosus.54
In conclusion, we found markedly lower pDC counts in peripheral blood but not in lymph nodes of patients with HIV-1. Thus, loss of pDC in PBMC of patients with HIV-1 cannot be explained simply by homing in lymphoid tissues. The mechanisms underlying the decline in circulating pDC remain to be identified, but our studies suggest an indirect effect of HIV-1 rather than direct HIV-dependent cytotoxicity. Residual circulating pDC in patients with HIV-1 expressed markedly higher levels of IFNα and MxA mRNA and protein at steady state. Chronic activation of the IFNα system-despite significant reduction of circulating pDC-correlated with declining CD4+ T-cell counts and disease progression, underlining the involvement of IFNα in HIV-1 immunopathogenesis.
We thank Katja Rolshofen for technical assistance, Ingrid Stahmer for the data handling of the patients and shipment of samples, and Tilman Brusis and Hartmut Pallasch for providing the tonsils of the HIV-negative controls.
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