Accumulation of DC-SIGN+CD40+ dendritic cells with reduced CD80 and CD86 expression in lymphoid tissue during acute HIV-1 infection : AIDS

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Accumulation of DC-SIGN+CD40+ dendritic cells with reduced CD80 and CD86 expression in lymphoid tissue during acute HIV-1 infection

Loré, Karina,b,*; Sönnerborg, Andersa,b; Broström, Christinab; Goh, Li-Eanc; Perrin, Lucd; McDade, Hughc; Stellbrink, Hans-Jürgene; Gazzard, Brianf; Weber, Rainerg; Napolitano, Laura A.h; van Kooyk, Yvettei; Andersson, Janb

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In vivo, immature dendritic cells (DC) engulf microbes and antigens in the periphery for subsequent intracellular processing into short peptides to be presented on MHC class I and II molecules. Such activated DC migrate to regional lymphoid compartments where they form a network as mature interdigitating DC enabling potent primary activation of naive T cells [1]. Antigen presentation by the mature DC also requires expression of co-stimulatory molecules, including CD40, CD80 and CD86, plus several cytokines in order to allow optimal expansion of antigen-specific T cells.

DC in the mucosa are the primary target cells for HIV-1 after sexual transmission [2]. Capture of HIV-1 is mediated by several receptors expressed on DC including CD4, CCR5 and CXCR4 [3] and the unique DC-SIGN molecule, which binds the HIV-1 envelope protein gp120 [4]. This latter interaction may result in a temporary uptake of receptor-coupled HIV-1 by DC without degradation of the virus and thus allows HIV-1 to be re-expressed on the cell surface.

Consequently, DC have a pivotal role in the pathogenesis of HIV-1 infection through delivery of the virus to lymphoid sites and transmission to neighboring CD4 T cells [5]. Also, DC expressing defects in their antigen-presenting capacity may lead to defective HIV-1-specific CD4 T cell activation [6,7].

The aims of the current study were to characterize the recruitment of interdigitating DC in vivo to lymphoid tissue (LT) during different stages of HIV-1 infection. The phenotypic features of DC required for efficient antigen presentation were also studied.


Patients and controls

LT biopsies [lymph nodes (LN) and tonsils] were collected from four cohorts of HIV-1-infected patients. The patients with acute HIV-1 infection (aHI) were enrolled in the international Quest/Probe3005 study [8]. Untreated patients with slow progressive HIV-1 infection (SLP) or long-term non-progessors (LTNP) [9], aviremic patients on potent antiretroviral treatment, control HIV-1-seronegative healthy individuals and patients with acute Epstein–Barr virus (EBV) infection were recruited from Huddinge University hospital, Stockholm, Sweden. Patients with AIDS came from San Francisco General Hospital at the University of California, San Francisco. Approvals were obtained from the institutional review boards and ethical committees at each participating site. Previous studies have shown no difference in cell distribution in the parafollicular areas of the LT nor in the frequency of cytokine or co-stimulatory expressing cells between LN and tonsilar tissue [10–12]. Furthermore, earlier studies did not find any difference in LT with age using specimens obtained from individuals between 5 and 60 years of age, which justified the use of seronegative control material from younger patients than the HIV-1- infected cohort [10,11,13].

CD4 T cell and viral load estimates

CD4 T cell counts in peripheral blood were determined by routine flow cytometric analysis. Plasma viral loads were determined with the Amplicor HIV Monitor test (Roche Molecular Systems, Sommerville, New Jersey, USA) with a detection limit of < 50 copies/ml. Cell-associated HIV-1 RNA and DNA were measured on cells obtained from LT and peripheral blood mononuclear cells (PBMC) using the reagents of the Amplicor HIV-1 Monitor assay (Roche) [14]. Cell-associated HIV-1 DNA was measured by incubating the nucleic acid preparation with Rnase A Dnase-free (Sigma, Buchs, Switzerland). One million cells from each sample were analyzed. The cut-off values were approximately 3 RNA copies/106 cells and 5 DNA copies/106 cells. The mean coefficient of variation for 1000–10 copies/106 cells was 12% (range, 3–26) for cell-associated RNA and 18% (range, 2–29) for cell-associated DNA.

Detection of cytokines and cellular markers by immunohistochemistry

The staining procedure used on cryopreserved LT to identify cytokines and cell surface markers at the single cell level has previously been described [15]. The staining reactions were developed brown using diaminobenzidine tetrahydrochloride (DAB). Anti-CD8 (SK1, Becton Dickinson, San Jose, California, USA) was the CD8 T cell subset-specific antibody. The DC phenotypic antibodies were anti-CD1a (NA 1/34, Dako, Glostrup, Denmark), anti-S-100 β-subunit (S-2532, Sigma, St Louis, Missouri, USA), anti-CD83 (HB15e, PharMingen, San Diego, California, USA), anti-DC-LAMP (provided by Dr S. Lebecq [16]) and anti-DC-SIGN (provided by Dr Y. van Kooyk, [4]). The antibodies against co-stimulatory molecules were anti-CD40 (S2C6, Mabtech, Nacka, Sweden), anti-CD80 (anti-CD80, BB1, PharMingen, and anti-BB-1, L307.4, Camfolio, Becton Dickinson) and anti-CD86 (IT2.2, PharMingen). The cytokine-specific antibodies and the secondary antibodies were previously described [15].

Quantification of cytokines and phenotype of cells by in situ image analysis

Digital images of stained samples were transferred into a DMR-X microscope (Leica, Wetzlar, Germany) and further into a computerized image analysis system (Quantimet 550IW, Leica, Cambridge, UK). The complete LT sections were assessed in each sample in a semiquantitative way by a specialized software program [15,17]. The presence of cells expressing DC-specific antigens, CD40 and CD8 was evaluated as the percentage positive stained cellular area out of the total hematoxylin stained cellular area, which was 3–7 × 105/μm2 representing approximately 0.4–1.1 × 104 cells. Furthermore, the numbers of cytokine-, CD80- and CD86-expressing cells were estimated by counting the positive cells in each digital image manually. The number of positive cells out of the total number of cells was calculated and presented as expressing cells per 10 000 cells. All of the analyses were performed in a blinded fashion. These evaluation methods have an interassay variability of < 10% [17,18].

Two-colour staining for co-localization

The staining experiments with immunofluorescence were carried out in blocking serum BSA-C (Aurion BSA-C, Wagningen, the Netherlands). The immunoglobulin-specific secondary biotinylated antibodies [15] were used together with Alexa 488- or Alexa 546-labeled streptavidin (Molecular Probes Inc., Eugene, Origon, USA) and an avidin–biotin blocking kit (Vector Laboratories, Burlingame, California, USA). The sections were evaluated in a laser scanning confocal microscope (SP102, Leica). Cells stained with two colours were counted manually in a blinded fashion.

Statistical analysis

Statistical significance was assessed by the Mann–Whitney U-test and considered significant at two-tailed P value < 0.05.


Increase of interdigitating dendritic cells in the acute phase of HIV-1 infection

Interdigitating DC are mainly found in the parafollicular T cell-rich areas of LT [1]. DC represent a heterogeneous cell lineage with several phenotypic and functional subsets of cells, not all of which have been fully elucidated [19]. Several antigens expressed on DC were, therefore, used to assess their phenotype and maturation [20]. Interdigitating DC characterized by CD1a, S-100b, CD83, DC-LAMP and DC-SIGN expression had similar localization in LT derived from tonsils or LN (Fig. 1). Furthermore, DC expressing CD1a, S-100b and DC-SIGN and, to lesser extent, more mature DC expressing CD83 and DC-LAMP were also localized in the epithelial layer and crypts in tonsils. However, in order to standardize the assessments, DC localized in the epithelial regions were not included in the assessments.

Fig. 1.:
Dendritic cells (DC: stained brown with diaminobenzidine tetrahydrochloride) in the parafollicular area of lymphoid tissue in a patient with acute HIV-1 infection (a,b), a patient with acute Epstein–Barr infection (c,d) and a HIV-1-seronegative healthy control (e,f). (a,c,e) Show that expression of CD1a on DC was high in the patients with acute HIV-1 infection (a) and acute Epstein–Barr infection (c) and low in the HIV-1-seronegative healthy control (e). (Magnification ×140). (b,d,f) show DC-SIGN expression on DC in the same patients. (Magnification ×200).

The LT from the patients with aHI was characterized by a significantly, up to tenfold, increased frequency of DC expressing CD1a, S-100b, CD83 and DC-SIGN compared with HIV-1-seronegative healthy controls (P < 0.02) (Fig. 2). Fewer cells expressed DC-LAMP. The LT from these patients also showed 6- and 25-fold higher intracellular HIV-1 DNA and HIV-1 RNA loads, respectively, compared with that in PBMC (P < 0.01) (Table 1). In addition, a similar pattern of accumulation of mature DC in LT was seen in the patients with acute EBV infection. The frequency of interdigitating DC in LTNP/SLP was lower than in the patients with aHI or EBV infection but still elevated compared with HIV-seronegative individuals (P < 0.04, for CD1a and CD83DC). A small increase in the frequency of DC was also noticed in the patients successfully treated with antiretroviral drugs, with the exception of S-100b DC, which persisted in elevated numbers (P < 0.03). In contrast, the total frequency of DC was reduced in LT from the patients with AIDS compared with the other HIV-1-infected cohorts. However, the total cellularity in the LT from the patients with AIDS was also severely depleted, resulting in a relative increase in the percentage of S-100b-expressing area compared with LT from seronegative controls (P < 0.02).

Fig. 2.:
The frequency of dendritic cells expressing various markers given as the range and median values of the percentage of specific protein-expressing area within the total cellular area as quantified by in situ imaging. *P < 0.05 compared with HIV-1-seronegative healthy controls.
Table 1:
Patient cohort characteristics.

Dissociation between CD40 and CD80/86 expression

The typical pattern of CD40 expression consisted of a weak staining in the follicles (CD40lowcells, presumably B cells) and bright staining in the parafollicular areas (CD40highcells), as reported earlier [21] (Fig. 3d). Higher intensity values for CD40 staining measured by computerized imaging were obtained in parafollicular areas than in follicular areas. Double labeling showed that the bright CD40high expression was to a great extent (> 80%) co-localized with CD1a DC in the parafollicular areas. A significantly increased frequency of CD40high cells was evident in patients with aHI and in patients with acute EBV infection compared with that in healthy controls (P < 0.03) (Fig. 3g). The healthy controls, the LTNP/SLP and the treated aviremic cohort showed comparable levels (P = 0.06). Patients with AIDS showed a significantly reduced expression of CD40high in the LT (P < 0.02).

Fig. 3.:
Expression of co-stimulatory molecules by dendritic cells (DC) in lymphoid tissue. (a) Co-localization of CD1a DC (green) and CD80 DC (red) (white arrow) in a patient with acute HIV-1 infection. (b) CD80 DC (brown) occurred as single individual cells or small clusters in patients with acute HIV-1 infection. (c) Higher numbers of CD80 DC were forming a complete network in a patient with acute Epstein–Barr virus (EBV) infection. (d) Brightly CD40high expressing cells were found in the parafollicular areas in the LT in LTNP. (e–h) Graphs showing the range and median values of incidences of marker-expressing cells at different stage of HIV-1 infection and in EBV infection; seroneg, HIV-1 seronegative; LTNP/SLP, long-term non-progessors/slow progressive of HIV-1 infection. (e,f) Data represent the numbers of CD80 cells and CD86 cells out of 10 000 cells. (g,h) Percentage of CD40- and CD8-expressing area out of the total cellular area is shown. *P < 0.05 and **P < 0.01 compared with values for lymphoid tissue in seronegative healthy controls. [Magnification (a) ×360, (b,c) ×180 and (d) ×30.]

The majority (> 60%) of bright CD80 and CD86 cells were also found to be co-localized with CD1a DC in the parafollicular areas in HIV-1 infection, EBV infection and seronegative healthy tissue (Fig. 3a). LT from patients with aHI showed upregulation of the number of both CD80 and CD86 cells compared with uninfected healthy controls (Fig. 3e,f) (P < 0.02). However, patients with acute EBV infection showed significantly higher numbers of CD80 and CD86 DC in the parafollicular area compared with patients with aHI (P < 0.01). In acute EBV infection, CD80 and CD86 cells generated a complete network (Fig. 3c), which co-localized with DC expression of CD1a, S-100b, DC-LAMP and DC-SIGN. This was in contrast to the patients with aHI, in whom no intact network for CD80 and CD86 was seen. CD80 and CD86 expression was found in scattered clusters of cells in LT from HIV-1-infected individuals (Fig. 3b). The CD80 and CD86 cells were not significantly increased in the LT from LTNP/SLP, patients with AIDS or in aviremic treated patients compared with seronegative controls, again leading to an incomplete network of CD80 and CD86 expression compared with that in acute EBV infection (Fig. 3e,f). No difference in distribution or frequencies of CD40, CD80 and CD86 cells was found to correlate with the age of the patients or the type of biopsy donated.

Increased CD8 expression

An increased frequency of CD8 T cells in LT was observed in the patients with aHI, in LTNP/SLP and in treated chronically infected asymptomatic HIV-1 patients compared with the controls (Fig. 3h) (P < 0.02). In contrast, the patients with AIDS had a severe depletion in the absolute number of cells, resulting in low total levels of CD8 T cells.

Expression of cytokines

Interleukin 1 receptor antagonist (IL-1ra) was mainly expressed in CD1aDC localized in the crypts or in epithelial layer of the tonsils. However, the number of IL-1ra-expressing cells within the LT was also found to be upregulated in the aHI cohort, but not other cohorts, compared with the healthy controls (P < 0.01) (Table 2). Interleukin (IL) 1α was predominantly expressed in endothelial cells in high endothelial venules. Approximately 10% of all IL-1α-expressing cells were CD1aDC in the parafollicular area. The total number of IL-1α-expressing cells, compared with healthy controls, was increased during aHI (P < 0.01) and in LTNP/SLP (P < 0.02), but not in those with AIDS or those treated with antiretroviral drugs. IL-1β was expressed mainly in the parafollicular areas. CD1a DC represented a portion (roughly estimated 20–50%) of the total number of IL-1β-expressing cells. The patients with aHI showed a significant rise in IL-1β-expressing cells compared with healthy controls (P < 0.01), which was not observed in the other HIV-1-infected cohorts. In contrast, expression of tumor necrosis factor α was not significantly upregulated in any of the HIV-1 cohorts.

Table 2:
Relative incidence of cytokine-expressing cells in lymphoid tissue during HIV-1 infectiona.

Furthermore, the vast majority of the HIV-1-infected individuals had cells expressing interferon α (IFN-α) in their LT while IFN-α was only found in one of the four HIV-1-seronegative controls. The highest number of IFN-α-expressing cells was found in the patients with aHI, but the LTNP/SLP group also had a significant upregulation of IFN-α (P < 0.03). In addition, the aHI cohort had increase expression of IL-12 p70 (P < 0.02), which was not found in the other HIV-1 cohorts. No difference in the numbers of cytokine-expressing cells was found between LN and tonsil tissue.


Here, we provide evidence that there is a significant migration in vivo of DC to the lymphoid compartments in the early phase of HIV-1 infection to an extent that is comparable to that found in acute EBV infection. However, patients with aHI show a distinct block in co-stimulatory molecule expression by DC, as indicated by a significant induction of CD40 but, to lesser extent, CD80 and CD86. This suggests that functional defects of DC may occur at the very early onset of HIV-1 infection and are likely to persist in the asymptomatic chronic phase of the infection. In addition, DC binding of HIV-1 through CD4, CCR5, CXCR4 or the DC-SIGN receptor can promote transmission of HIV-1 to CD4 T cells [3,4]. Therefore, the massive migration we found of DC-SIGN DC to the LT shortly after HIV-1 transmission may be followed by a subsequent local activation of CD4 T cells, which would facilitate HIV-1 infection and replication in the latter cells [5,22]. Indeed, the intracellular HIV-1 DNA as well RNA levels were significantly higher in cells in the LT compared with PBMC in the acutely infected patients. This activation may contribute to the peak plasma viremia noticed during the initial phase in aHI [23].

The frequencies of immature DC in blood [6,24], skin [6] and mucosa [25] are reduced during HIV-1 infection. Our contrasting finding of an increase of interdigitating DC in LT suggests that there is a redistribution of DC from the periphery to the LT during HIV-1 infection. Several factors, such as persistent production of HIV-1 antigens or continuous proinflammatory cytokine and chemokine expression in LT, may be involved in this recruitment. Furthermore, persistent increased frequency of DC in LT could also reflect reduced elimination of DC by CD8 T cells and natural killer cells [26].

LTNP/SLP is a highly selected group representing a small proportion of HIV-1 patients who have preserved CD4 T cell levels and low viral replication [9]. Our finding of an increase in interdigitating CD1a DC and CD83 DC and CD8 T cells also in these patients indicates that this condition is associated with a persistent activation of cellular immunity. Patients on successful antiretroviral treatment had lower incidences of DC and CD8 T cells in the LT than patients with aHI and LTNP/SLP, which may indicate that pharmacologically suppression of virus production reduces the recruitment and need of DC and CD8 T cells. The fall in the levels of both DC and CD8 T cells in the group with AIDS compared with the other HIV-1-infected cohorts probably reflects the failure of the regenerative capacity of the immune system.

The maturation process of DC is complex [18]. This was exemplified in the current study by the high frequency of CD1a, S-100b, CD83, DC-SIGNDC without a concomitant increase of DC LAMP expression, a molecule upregulated late in the DC maturation process. In addition, CD40, required for CD40 ligand-mediated DC activation and maturation, was upregulated during the acute stage of both HIV-1 and EBV infection. However, CD80 and CD86 expression occurred only in scattered clustered DC in HIV-1 infection, in contrast to acute EBV infection where their expression was found on higher numbers of cells forming a complete parafollicular network. Consequently, the interdigitating DC in HIV-1 infection may have insufficient CD80 and CD86 expression for the proliferative induction of efficient numbers of naive CD4 T cells in antigen-specific responses. Activation of naive T cells requires both T cell receptor binding to the specific peptide antigen–MHC complex on antigen-presenting cells and ligation of CD80/CD86 and CD28/CTLA-4 on the T cell surface. Studies in knock-out mice have shown that CD80 and CD86 ligation is required for CD4 T cell proliferation upon antigen stimulation [27,28]. Downregulated expression of CD80 and CD86 on monocytes from HIV-1-infected individuals has previously been reported, which has been shown to lead to subsequent impaired activation of CD8 T cells, resulting in inefficient suppression of HIV-1 replication [29–31]. Binding of HIV-1 gp120 to CD4 T cells inhibits upregulation of CD80 on co-cultured antigen-presenting cells [32]. HIV-1 may, therefore, interfere with the regulation of co-stimulatory molecules, which would contribute to defects in the activation of HIV-1-specific T cell clone expansion. The ratio between intracellular HIV-1 RNA levels in LT and PBMC was 25:1 in the aHI patients, indicating much higher production of gp120 in LT where the downregulation of CD80/CD86 expression was found. The functional capacity of the DC-mediated HIV-1-specific T cell response in the patients enrolled in the current study has not been investigated. However, there are data indicating that in vitro cultured blood-derived DC exposed to HIV-1 do not upregulate CD80/CD86 and are unable to induce proliferation of T cells [7,33], although immature blood-derived DC may not completely mimic the interdigitating mature DC in LT.

Another important aspect to consider is that HIV-1 harboring mature CD80/CD86 DC promote virus transmission and replication in clustered CD4 T cells both in vitro and in vivo [5,34]. HIV-1 particles expressing CD80 and CD86 in the viral envelope, derived from the host cells, mediate intracellular signaling cascades that upregulate HIV-1 replication in infected CD4 T cells [35]. Furthermore, the transmission of HIV-1 from DC to CD4 T cells can be inhibited by anti-CD80 or anti-CTLA-4 antibodies [36]. Restricted upregulation of these accessory molecules may, therefore, play a role in the host's prevention of DC-mediated spread of the virus to adjacent T cells.

The cytokine production profile of DC changes with the differentiation pathway [37]. Here, we could demonstrate differences in the cytokine expression pattern in DC depending on the maturity and the locality of the DC in the tissue. Intraepithelial immature CD1a DC expressed IL-1ra, while mature CD1a DC in the parafollicular areas of the LT expressed IL-1β and IL-1α. In addition, there were elevated numbers of cells expressing IL-12 and IFN-α in the parafollicular area in LT from aHI patients. We have previously found that cultured immature DC constitutively expressed IL-1ra and needed stimulation to induce production of other cytokines such as IL-1β, tumor necrosis factor α and IL-12 [33]. The mature DC accumulated in LT during HIV-1 infection found in the current study appeared to be functional in their ability to produce the cytokines inducing T helper type 1 responses. However, high expression of proinflammatory cytokines in DC may also induce HIV-1 replication in CD4 T cells [33].

In conclusion, accumulation of DC in LT was similar in aHI and acute EBV infection. A persistent small increase of interdigitating DC was also observed in LTNP/SLP. However, DC did not seem to undergo full differentiation in HIV-1 infection during the initial adaptive immune response, characterized by incomplete upregulation of the co-stimulatory molecules CD80 and CD86. The failure of DC to differentiate completely is likely a factor limiting their capacity to generate HIV-1-specific CD4 T cell responses. Consequently, lack of proper helper function may restrict CD8 T cell maturation and prevent effective elimination of HIV-1-infected cells. However, one may also argue that this downregulation is important to control the activation of T cells and thereby HIV-1 replication. Overall, reconstitution of CD80/CD86 expression in DC may be important to consider in development of therapeutic HIV-1 vaccines to be administrated in combination with effective antiretroviral therapy.


We would like to thank all the patients involved and the recruiting sites for enrolling patients in the Quest PROB 3005 study and Glaxo Wellcome UK for organizing the network. We are also grateful for the assistance of those who provided additional LT biopsies including Karin Ågren, Huddinge University Hospital, Nancy Abbey, Brian Herndier and Joseph M. McCune, San Francisco General Hospital, University of San Francisco, San Francisco. We also thank PharMingen, San Diego, California for kind supply of cell marker- and cytokine-specific antibodies and Dr S. Lebecq, Laboratory of Immunological Research, Dardilly for kindly providing the anti-DC-LAMP antibody.


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HIV-1; dendritic cell; cytokines; co-stimulatory molecule; DC-SIGN; lymphoid tissue; primary HIV-1 infection

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