HIV Interferes with the Dendritic Cell–T Cell Axis of Macrophage Activation by Shifting Mycobacterium tuberculosis–Specific CD4 T Cells into a Dysfunctional Phenotype

It is estimated that one third of the world’s population is latently infected with Mycobacterium tuberculosis (1). Over 90% of these infected individuals will never develop active tuberculosis (TB), confirming that the human immune system is capable of controlling M. tuberculosis. However, a failure to control infection occurs during periods of immunosuppression, and it is clear that HIV infection induces immunosuppression and that HIV–M. tuberculosis coinfection is the strongest risk factor for transition of latent infection into active TB (2). In HIV-uninfected individuals, the risk of transition from latent to active TB is 5–10% in a lifetime, which increases up to a 10% annual risk in HIV-infected individuals (3). The most evident immune defect caused by HIV is a progressive reduction in CD4 T cells, which correlates with an increased risk of developing active TB (4), asserting the critical role of CD4 T cells for an efficient M. tuberculosis immune response. Furthermore, even HIV-infected individuals with restored CD4 counts due to antiretroviral therapy (ART) still have an increased risk of developing TB (5). This implies that not only the number of CD4 T cells but also functional aspects of the M. tuberculosis–HIV interaction are of importance. Epidemiological data from HIV–M. tuberculosis–coinfected individuals show both depletion of and impairments in the M. tuberculosis Ag-specific CD4 T cell responses (6–8), although the precise effect on M. tuberculosis Ag-specific CD4 T cells and the molecular mechanism leading up to this dysfunction in the TB protective immunity have not been thoroughly investigated. We recently showed that HIV interferes with dendritic cells’ (DCs) capacity to process and present M. tuberculosis Ags to M. tuberculosis Ag-specific CD4 T cells (9). The functional aspects of this effect have not yet been investigated (i.e., impairment of M. tuberculosis growth restriction within macrophages), including if it is this suboptimal activation by DCs that is the key, or whether the resulting DC–T cell interaction is involved also in triggering a dysfunctional and polarized CD4 T cell response, contributing to the increased risk for reactivating TB in HIV coinfection.

M. tuberculosis Ag-specific CD4 T cells have a central role in the adaptive immune responses and are important for protective immunity against TB (10). Upon activation, M. tuberculosis Ag-specific CD4 T cells secrete IFN-γ and TNF-α, which in turn activate antimycobacterial mechanisms in macrophages (11). DCs are not only capable of activating T cells but can also induce responses not beneficial in the host response against M. tuberculosis, such as immune tolerance (12). The nature and intensity of an immune response are determined by the quality of MHC peptide TCR interaction and subsequent signaling and also by the balance between coinhibitory and costimulatory molecules and receptors (collectively referred to as cosignaling molecules in this article) activated on the DCs and T cells by the cellular cross-talk. The expression and functions of cosignaling molecules are context dependent, and they have a crucial role in T cell biology, in which they regulate pivotal functions like T cell activation and differentiation (13–15).

To avoid immune-mediated pathology and to preserve homeostasis, regulatory T cells (Tregs) maintain a constant equilibrium between pathogens and the host immune response. Control of host effector mechanisms by Tregs is a balancing act and has been shown to contribute to immunosuppression, thereby providing opportunity for pathogens to survive and persist in the host (16–18). Tregs are expanded during TB (19) and HIV-TB infection (20). We have previously reported that HIV–M. tuberculosis coinfection modulates DC functionality by affecting the activation, cytokine response, and Ag presentation efficiency of DCs (9). These dysfunctional DCs could further regulate the expression levels of cosignaling molecules on T cells and thereby activate Tregs. There is a lack of understanding of how cosignaling molecule levels and functionality of M. tuberculosis Ag-specific CD4 T cells are modified by DCs coinfected with HIV and M. tuberculosis and their role in immune activation or suppression during M. tuberculosis infection.

Hence, this study was undertaken to explore if HIV–M. tuberculosis–coinfected DCs dysregulate the function of M. tuberculosis Ag-specific CD4⁺ T cells. In our model, we could show a loss of control of M. tuberculosis infection in human macrophages by this dysregulation. Further, a clear pattern appeared in which HIV–M. tuberculosis coinfection of DCs impaired the proliferation of M. tuberculosis Ag-specific CD4 T cells, upregulated coinhibitory molecules as well as Treg markers, and downregulated costimulatory molecule expression on M. tuberculosis Ag-specific CD4 T cells. This dysregulated response is of functional relevance as these M. tuberculosis Ag-specific CD4 T cells had reduced capacity to activate M. tuberculosis–infected macrophages.

Purified protein derivative (PPD; culture filtrates from M. tuberculosis strain H37Rv) was obtained from the Statens Serum Institut (Copenhagen, Denmark). Recombinant human GM-CSF and IL-4 were from PeproTech. Recombinant human IL-2 and CD1c (BDCA-1) PE were from Miltenyi Biotec (Bergish Gladbach, Germany). CD3 Pacific Orange was from ExBio (Vestec u Prahy, Czech Republic). CD86 Alexa Fluor 700, CD28 PE, CD40L FITC, HLA-DR PerCP, CD1a PE, FOXP3 Alexa Fluor 488, and Alexa Fluor 488 Mouse IgG1 κ isotype control were from BD Pharmingen. Blimp1 (PRDM1) PerCP was from Nonvus Biologicals (Abingdon, U.K.). CD4 Pacific Blue, CTLA-4 PerCP/Cy5.5, PD-1 Alexa Fluor 700, CD25 PE, ICOS Alexa Fluor 700, PD-L1 PerCP/Cy5.5, and CD40 PE were from BioLegend (San Diego, CA).

Non-inactivated human serum (pooled from five donors, used for opsonization) and buffy coats were obtained from healthy individuals (from Linköping Blood Bank, Linköping, Sweden) who had given written consent for research use of the donated blood in accordance with the Declaration of Helsinki and paragraph 4 of the Swedish law (2003:460) on Ethical Conduct in Human Research. All human samples intended for research were anonymized by Linköping Blood Bank.

HIV-1BaL (lot p4238) was produced using chronically infected cultures of the ACVP/BCP cell line (no. 204). Virus was purified and concentrated as previously described (21), and aliquots were frozen in liquid nitrogen vapor. HIV was opsonized by incubating equal volumes of HIV and non-inactivated human serum for 1 h at 37°C (hereafter referred to as HIV) before infection. M. tuberculosis H37Rv was grown at 37°C in Middlebrook 7H9 broth supplemented with 0.05% Tween 80 and 10% ADC enrichment (Becton Dickinson), with the addition of 100 μg/ml hygromycin for luciferase-expressing M. tuberculosis used for macrophage infection. Log-phase M. tuberculosis was pelleted (5000 × g, 5 min) and passed through a 26-gauge needle 10 times in PBS. M. tuberculosis was opsonized by incubating equal volumes of bacterial suspension in PBS and non-inactivated human serum for 30 min at 37°C. The bacterial suspension was vortexed three times during this incubation. The bacteria were pelleted, suspended in RPMI medium, and again needle-sheared to achieve single-cell suspensions. The OD values were measured to determine the bacterial concentration, followed by calculation of multiplicity of infection (MOI), and verified by serial dilution and plating on 7H10 agar. Initial titrations of the dose of M. tuberculosis/DC ratio was done using GFP-expressing H37Rv. M. tuberculosis MOI 1 gave ∼60% infection of DCs at 24 h independent of HIV exposure and was chosen to avoid the impaired viability of the DCs observed at higher MOIs. For further experiments, the unlabeled M. tuberculosis was used for DC infection.

PBMCs were separated from buffy coats of healthy blood donors by density-gradient centrifugation on Lymphoprep (Axis-Shield, Oslo, Norway). Human monocyte-derived macrophages (hMDMs) were derived from buffy coats as previously described (22). To achieve an autologous in vitro priming system for generating M. tuberculosis Ag-specific CD4 T cells, PBMCs from the same donors used for DC generation were used to purify CD4⁺ naive T cells through negative selection using the Human Naive CD4⁺ T Cell Isolation Kit according to manufacturer instructions (STEMCELL Technologies, Grenoble, France). The sorted cells were >95% pure naive CD4⁺ T cells (CD3⁺CD4⁺CD45RA⁺CD45RO⁻). The DC progenitors were selected by plastic adherence of PBMCs in tissue culture dishes (Corning Falcon, Durham, NC) for 1–2 h at 37°C in a 5% CO₂ incubator. The plates were washed gently thrice with RPMI 1640 to remove nonadherent cells. The DC progenitors were cultured in complete tissue culture medium (RPMI 1640, 2 mM l-glutamine, 10 mM HEPES, 13 mM NaHCO3, 100 μg/ml penicillin, 100 μg/ml streptomycin) and 5% heat-inactivated human AB serum (Sigma-Aldrich) supplemented with 10 ng/ml recombinant human GM-CSF and 10 ng/ml recombinant human IL-4 for 4 d. The DCs generated were CD1a/c⁺.

As a precaution, the generated DCs were gamma-irradiated (25 Gy) to negate the effect of any residual T cells originating from the DC culture before their coincubation with naive CD4⁺ T cells for priming. CD4⁺ T cells were cultured with autologous gamma-irradiated DCs at a 4:1 ratio and PPD (10 μg/ml) to generate M. tuberculosis Ag-specific CD4⁺ T cells. Cells were supplemented with IL-2 (20 IU/ml; confirmed by titration, the lowest dose to sustain T cell viability) at the onset of culture and replenished once a week. The Ag specificity was evaluated 3 wk later as previously described (9) using medium control, OVA (unspecific Ag background control), PPD, or staphylococcal enterotoxin B (SEB; unspecific Ag activation, positive control). Prior to experiments, M. tuberculosis Ag-specific CD4⁺ T cells were rested overnight without IL-2, and IL-2 was continuously absent in all experiments.

DCs were thawed and seeded (2 × 10⁴ per well) in 96-well plates 1 d before infection, and GM-CSF and IL-4 were excluded from this point forward. DCs were infected with opsonized HIV-1, 1 μg/ml (p24 equivalent/ml), for 2 h prior to infection with opsonized M. tuberculosis (MOI 1). The rationale for opsonizing HIV and M. tuberculosis is that in the host pathogens are naturally opsonized by bodily fluids, and in tissues, during infection (23–25). Furthermore, HIV uses complement opsonization to enhance infection of immature DCs (26), that is, complement opsonization functioning as an immune evasion mechanism to establish infection in a silent manner (27). After overnight infection (16 h), supernatants were removed, and infected DCs were cocultured with autologous M. tuberculosis Ag-specific CD4⁺ T cells (1 × 10⁵ cells per well). At the time of coculturing of DCs and CD4⁺ T cells, the fresh medium was supplemented with 10 μM azidothymidine to prevent HIV replication. Cell-free supernatants were examined for cytokine production after 48 h of coculture using cytometric bead array (as described below).

M. tuberculosis Ag-specific CD4⁺ T cells were resuspended in PBS and labeled with 1 μM CFSE for 5 min at 37°C and 5% CO₂. The staining was quenched by adding equal volumes of FBS followed by washing three times with RPMI medium.

DCs, after overnight infection, were cocultured with CFSE-labeled CD4⁺ T cells as described in the previous section. After 72 h of coculture, CD4⁺ T cells were harvested, stained with T cell markers, fixed with 4% paraformaldehyde (PFA), and acquired on a Gallios flow cytometer (Beckman Coulter). The proliferation of CD4⁺ T cells was measured by CFSE dilution by gating on CD3⁺CD4⁺ cells. The data analysis for all flow cytometric evaluations was performed by Kaluza 1.3 software (Beckman Coulter), and representative images were produced by FlowJo version 10.1r5 (OR).

After 72 h of coculture, T cells were collected by gentle pipetting. For harvesting of DCs, partially adherent cells were incubated with 5 mM EDTA for 20 min followed by flushing two times with PBS. Collected T cells were stained to analyze the surface expression of cosignaling molecules using Abs directed against CD3 Pacific Orange, CD4 Pacific Blue, CTLA-4 PerCP/Cy5.5, PD-1 Alexa Fluor 700, CD25 PE, ICOS Alexa Fluor 700, CD28 PE, CD40L FITC, and Blimp1/PRDM1 PerCP for immunophenotyping of T cells. To appreciate the magnitude of cosignaling molecule expression of CD4 T cells activated by infected DCs, SEB-loaded DCs were used as an unspecific activation control (merely used as a reference for T cell activation in the absence of infection and not included in the statistical comparisons).

The expression of Treg marker FOXP3 required intracellular staining. Briefly, after surface staining against CD3, CD4, and CD25, the cells were fixed and permeabilized using the human FOXP3 buffer set (BD Biosciences) according to the manufacturer’s protocol. The cells were subsequently stained intracellularly with FOXP3 Alexa Fluor 488 or the Alexa Fluor 488 Mouse IgG1 κ isotype control.

To analyze the expression of surface molecules on DCs, cells were surface stained with Abs directed against CD3 Pacific Orange, CD4 Pacific Blue, PD-L1 PerCP/Cy5.5, CD40 PE, and CD86 Alexa Fluor 700. Compared to the instrument setting for T cell analysis, flow cytometry analysis of DCs required reducing the instrumental gain from 10 to 1 for side scatter and from 5 to 1 for forward scatter, and therefore the forward/side scatter of the two cell types cannot be compared. Cells within the high-scatter DC gate were CD3⁻CD4⁻ and verified to be of HLA-DR^high and CD1a/c⁺ DC phenotype. All stained cells were fixed with 4% PFA prior to acquisition on the Gallios flow cytometer.

M. tuberculosis Ag-specific CD4⁺ T cells from cocultures of noninfected and infected DCs were washed, stained with CD3 Pacific Orange, and thereafter stained with FITC–annexin V according to the manufacturer’s protocol using propidium iodide to counterstain necrotic cells. As a positive control, CD4⁺ T cells cocultured with DCs were either exposed to UV light for 10 min (240 J/s/m²) and further incubated for 24 h or were treated with staurosporine (2.5 μM) for 24 h prior to staining. Stained T cells were fixed with 4% PFA in 1× binding buffer (to preserve annexin V binding) and detected on a Gallios flow cytometer using the CD3⁺ gate and analyzed by Kaluza 1.3.

Cell-free culture supernatants were stored at −80°C until assayed. The levels of IFN-γ, TNF-α, IL-10, and TGF-β cytokine were determined in culture supernatants by cytometric bead array analysis, performed according to the manufacturer’s protocol (BD Biosciences). Detection of cytokines was performed on a Gallios flow cytometer, and cytokine concentrations were analyzed using the Kaluza 1.3.

Based on published data from our laboratory using different MOIs for M. tuberculosis infection of hMDMs, with low MOIs (1, 2) being controlled by hMDMs for 1–2 wk in contrast to higher MOIs (>5), in which MDMs lose control over M. tuberculosis replication (28, 29), in this study we chose MOI 2 for a controlled infection mimicking a scenario of latent TB. hMDMs were infected with luciferase-expressing M. tuberculosis, and infection was stopped after 1.5 h by removing extracellular bacteria by washing as previously described (22). The next day (18 h postinfection) T cells were collected from 72-h DC–T cell cocultures, and T cells were washed, counted, and then added to the infected hMDMs at a T cell/hMDM ratio that by titration was seen as best for controlling M. tuberculosis in hMDMs (1:6). Prior to adding T cells and 72 h after adding T cells, the extracellular and intra-hMDM M. tuberculosis content was measured. The bacterial luminescence in both supernatants and lysates were measured using a Modulus microplate (Turner BioSystems, Sunnyvale, CA) to present the total bacterial burden of M. tuberculosis–infected hMDMs that had been incubated with M. tuberculosis Ag-specific CD4 T cells that had been previously activated by uninfected, HIV-infected, M. tuberculosis-infected, or coinfected DCs.

Statistical analyses were performed with GraphPad Prism software (version 5.0f), and results are expressed as mean ± SD. The Shapiro–Wilk normality test was used, and for data that were too small to perform the normality test (n ≤ 6) and for data that were not normally distributed, a paired two-tailed nonparametric test was performed (either a Wilcoxon matched-pairs signed rank test for comparing the mean differences between two groups, or the Friedman test with Dunn post hoc test for multiple comparison). For normally distributed data, a paired two-tailed Student t test between two groups or a one-way ANOVA with Dunnett post hoc test for multiple comparison was used. A p value < 0.05 was considered significant. The comparison between the M. tuberculosis and HIV plus M. tuberculosis (coinfected) group using either the t test or the Wilcoxon rank test was performed to reveal if HIV can modify the response to M. tuberculosis, which was the main aim with the study. As uninfected DCs and HIV single-infected DCs were included as technical controls but do not directly answer the primary aim, the indicated multiple comparison test comparing all infected groups against the uninfected DC control is also shown.

To fully evaluate the change in effector function of M. tuberculosis Ag-specific CD4⁺ T cells in a scenario of HIV–M. tuberculosis coinfection, we assessed the full route of DC–T cell–mediated macrophage activation leading to control of intracellular M. tuberculosis. For that, M. tuberculosis Ag-specific CD4⁺ T cells were activated with uninfected or infected DCs, after which T cells were collected and cocultured with M. tuberculosis–luciferase-infected hMDM to determine their effect on M. tuberculosis growth inhibition in macrophages. We found that the CD4⁺ T cells from HIV–M. tuberculosis–coinfected DC cultures had a decreased capacity to activate hMDM, as indicated by the 1.86-fold increase in bacterial growth in those hMDMs compared with those exposed to T cells from M. tuberculosis single-infected DC cocultures (p = 0.0003, n = 10; Fig. 1). This loss in growth inhibition by HIV–M. tuberculosis–coinfected DC was also significant when compared with the uninfected DC control (p < 0.05), whereas CD4⁺ T cells from HIV single-infected DC cocultures were not impaired in their capacity to activate hMDM, indicating that only during HIV–M. tuberculosis coinfection of DCs is the TB protective function of M. tuberculosis Ag-specific CD4⁺ T cells impaired.

Our earlier study demonstrated that HIV–M. tuberculosis–coinfected DCs reduced the activation of M. tuberculosis Ag-specific CD4⁺ T cells (e.g., their INF-γ release response was reduced) (9). To investigate if HIV–M. tuberculosis coinfection of DCs also affects the proliferation of M. tuberculosis Ag-specific CD4⁺ T cells, a CFSE dilution assay was used. Coculture with M. tuberculosis single-infected DCs enhanced the M. tuberculosis Ag-specific CD4⁺ T cell proliferation compared with uninfected DCs (p < 0.01, n = 5; Fig. 2A, 2B), and HIV–M. tuberculosis–coinfected DCs lost the ability to induce a significant T cell proliferation. Again, the HIV single-infected DC group did not induce a response in the M. tuberculosis Ag-specific CD4⁺ T cells that differed from the response induced by the uninfected DC group and will henceforth not be focused on. M. tuberculosis Ag-specific CD4⁺ T cell proliferation was further compared with the IFN-γ release in a setup using the same factors as used for Ag-specificity testing of the CD4 T cells clones to reconcile with the level of proliferation in the uninfected controls. This shows that although there was a low level of unspecific background proliferation of the M. tuberculosis–specific CD4⁺ T cells, their specific activation viewed as the IFN-γ release response was highly specific for M. tuberculosis Ags (PPD) (mean ± SD was 3156 ± 963) and showed no activation in presence of the unspecific Ag control OVA or in the unchallenged group (Fig. 2C). The viability of DC-activated M. tuberculosis Ag-specific CD4⁺ T cells was high (≥90%) and comparable in the M. tuberculosis single-infected and HIV–M. tuberculosis–coinfected condition as shown by annexin V/propidium iodide staining (Fig. 2D).

As HIV–M. tuberculosis–coinfected DCs triggered a reduced proliferation of M. tuberculosis Ag-specific CD4⁺ T cells, this effect on total CD4 effector T cells could itself can have a detrimental effect on TB control. However, because the number of DC-activated M. tuberculosis Ag-specific CD4⁺ T cells used in the M. tuberculosis growth inhibition assay was adjusted to the number of macrophages in the wells (Fig. 1), thereby eliminating the effect of the varying T cell proliferation and T cell numbers caused by the preceding coculture with infected or coinfected DC, this indicates that not only the absolute number but also the functional properties of effector T cells is of importance. To investigate the factors that might be involved and to better understand the molecular mechanism of T cell activation or immune suppression, the expression of established coinhibitory and costimulatory molecules expressed by T cells were examined. Expression of multiple and different combinations of inhibitory receptors is associated with an immunological check point leading to T cell suppression (30). We found that M. tuberculosis single-infected DCs increased the expression and that HIV–M. tuberculosis–coinfected DCs further upregulated the expression of the coinhibitory molecules PD-1, CTLA-4, and Blimp-1 on M. tuberculosis Ag-specific CD4 T cells after 72 h of coculture, compared with the expression by uninfected and HIV single-infected DCs. For all coinhibitory molecules, both the protein expression levels (i.e., median fluorescence intensity [MFI]), and percentage of marker-positive CD4 T cells was significantly higher in the HIV–M. tuberculosis–coinfected compared with the M. tuberculosis single-infected group (Fig. 3A, 3B).

To delineate the relationship between HIV–M. tuberculosis coinfection in DCs and their regulation of costimulatory molecules on T cells, the expression of these surface molecules was examined. CD40L, CD28, and ICOS have been implicated as important costimulatory molecules (13). Compared with activation with M. tuberculosis single-infected DCs, the CD40L expression on M. tuberculosis Ag-specific CD4⁺ T cells activated with HIV–M. tuberculosis–coinfected DCs showed a decreased expression both in term of percentage of positive cells (p = 0.0078) and total MFI (p = 0.024; n = 10) (Fig. 4A, 4B). M. tuberculosis Ag-specific CD4⁺ T cells were all positive for CD28 and ICOS, but the analysis of changes in their total MFI revealed that activation by HIV–M. tuberculosis–coinfected DCs decreased the protein expression levels of CD28 (p = 0.0098 compared with activation by M. tuberculosis single-infected DC; n = 10) and that only activation with M. tuberculosis single-infected DCs showed a significant increase in ICOS expression (p < 0.05 compared with activation with uninfected DCs; n = 6) (Fig. 4A, 4B). Although the shift in expression by HIV–M. tuberculosis coinfection versus M. tuberculosis single infection of many costimulatory molecules was small, the overall pattern makes out a clear trend in which M. tuberculosis Ag-specific CD4⁺ T cells activated with HIV–M. tuberculosis–coinfected DCs show upregulation of several coinhibitory molecules and downregulation of several costimulatory molecules. This indicates a modulation and polarization of the T cells that could have a substantial effect on their functionality. Furthermore, our initial experiments reveal the same shift in expression of CD28 and PD-1 by HIV–M. tuberculosis coinfection when specifically analyzing the proliferating (CFSE-diluted) portion of the M. tuberculosis Ag-specific CD4⁺ T cells (Supplemental Fig. 1). This indicates that the altered expression of cosignaling molecules during HIV–M. tuberculosis coinfection is not restored in the proliferated portion of the T cells and that the variation of cosignaling molecule expression could play an important role in the immune response to TB.

TB is associated with immune modulation in which DCs play a pivotal role by either activating or suppressing the immune system. PD-L1 is an important molecule involved in the generation of tolerogenic DCs (31, 32). HIV–M. tuberculosis–coinfected DCs exhibited an increased expression of PD-L1, and more importantly, for assessing the immunological breaking point to tolerance, the ratio of coinhibitory PD-L1 to costimulatory CD40 molecule expression (MFI ratio) was increased compared with uninfected DCs (p < 0.05 for PD-L1/CD40; n = 4), and so was the PD-L1/CD86 MFI ratio in HIV–M. tuberculosis–coinfected compared with M. tuberculosis-single infected DCs (p = 0.019; n = 10) (Fig. 5B).

From the same cocultures, M. tuberculosis Ag-specific CD4⁺ T cells activated with HIV–M. tuberculosis–coinfected DCs had higher expression of PD-1 (Fig. 3), and the ratio of coinhibitory PD-1 to costimulatory CD40L molecule expression was higher compared with T cells activated by M. tuberculosis single-infected DCs (p = 0.003; n = 9; Fig. 5C). Moreover, only the PD-1/CD40L MFI ratio of the M. tuberculosis Ag-specific CD4⁺ T cells activated with HIV–M. tuberculosis–coinfected DCs increased compared with activation by uninfected DCs (p < 0.05; n = 9). This suggest that HIV–M. tuberculosis–coinfected DCs induce an immunosuppressive M. tuberculosis Ag-specific CD4⁺ T cell phenotype.

Pathogens actively provoke the generation of Treg, thereby evading the immune response (19). Accordingly, we observed an increased expression of CD25^hiFOXP3⁺ T cells among the M. tuberculosis–specific CD4⁺ T cells that had been activated with HIV–M. tuberculosis–coinfected DCs compared with those T cells activated by M. tuberculosis single-infected DCs (p = 0.04, Fig. 6), suggesting a greater switch to a Treg phenotype during HIV–M. tuberculosis coinfection. M. tuberculosis Ag-specific CD4⁺ T cells activated with HIV–M. tuberculosis–coinfected DCs was also the only group among the infected groups that showed a significant increase in the frequency of CD25^hiFOXP3⁺ T cells compared with activation by uninfected DCs (p < 0.01).

One important effector function mediated by CD4 T cells is the release of activating or immunosuppressive mediators. We therefore analyzed the cytokine levels in cell-free culture supernatant collected after 48 h coculture of DCs and M. tuberculosis Ag-specific T cells. In cocultures from HIV–M. tuberculosis–coinfected DCs, there was a significant reduction in the levels of the Th1-associated cytokines IFN-γ and TNF-α compared with cocultures from M. tuberculosis single-infected DCs (p = 0.006 and p = 0.031, respectively), and for TNF-α, only the M. tuberculosis single-infected DC–T cell coculture induced a significant induction compared with uninfected DCs (p < 0.01) (Fig. 7). In agreement with the increased shift toward a CD25^hiFOXP3⁺ Treg phenotype (Fig. 6), the secretion of the immunosuppressive cytokines TGF-β and IL-10 was higher in cocultures from HIV–M. tuberculosis–coinfected DCs compared with cocultures from M. tuberculosis single-infected DCs (p = 0.007 and p = 0.031, respectively; Fig. 7). When comparing the cytokine levels to uninfected controls, TGF-β was the only cytokine not increasing during M. tuberculosis single infection. Instead, M. tuberculosis infection decreased a basal TGF-β production seen in this DC M. tuberculosis Ag-specific T cell coculture system, and HIV–M. tuberculosis coinfection of the DCs prevented this M. tuberculosis regulation/suppression of this cytokine.

M. tuberculosis–specific CD4 T cells have a crucial role in the development of adaptive immunity and are essential for providing protection against M. tuberculosis infection (10) in which, through the DC–T cell axis, they participate in the activation of macrophages. We recently demonstrated that HIV interferes with DC capacity to process and present M. tuberculosis Ags to M. tuberculosis Ag-specific CD4⁺ T cells (9). Besides being suboptimal for activating M. tuberculosis Ag-specific CD4⁺ T cells, in this article, we show that HIV–M. tuberculosis–coinfected DCs skews these effector T cells into a dysfunctional phenotype that is immune suppressive rather than protective against M. tuberculosis. Although some results by themselves do not show great differences (albeit significant), the results from the functional assay (M. tuberculosis growth inhibition assay) and those from the phenotypic markers make out an overall clear pattern strongly arguing for a functional link. HIV–M. tuberculosis–coinfected DCs was shown to be of a tolerogenic phenotype and stimulated M. tuberculosis Ag-specific CD4⁺ T cells to a shift into enhanced coinhibitory molecule expression and favoring the release of the immunosuppressive cytokines IL-10 and TGF-β over that of TNF-α and IFN-γ. M. tuberculosis Ag-specific CD4⁺ T cells activated with HIV–M. tuberculosis–coinfected DCs also took on more of a CD25^hiFOXP3⁺ regulatory phenotype, and coculture with these T cells enhanced growth of M. tuberculosis in macrophages rather than controlling the intracellular M. tuberculosis growth, as seen in the case when these T cells were activated with M. tuberculosis single-infected DCs. We could not detect that HIV single-infected DCs modulated any of the M. tuberculosis Ag-specific CD4⁺ T cell functions investigated in this DC–T cell autologous system, indicating that HIV only during coinfection with M. tuberculosis has the capacity to impair the TB protective functions of M. tuberculosis Ag-specific CD4⁺ T cells.

Several studies have shown that HIV–M. tuberculosis coinfection contributes to a reduction of M. tuberculosis–specific CD4 T cells (33–35), although it should be considered that what is reported as reduced numbers could also be partly influenced by the effect of T cell impairments rendering certain cells unresponsive to M. tuberculosis Ags and therefore undetectable. Using our in vitro approach of HIV infection of DCs in which the infected DCs received fresh medium with the antiretroviral agent azidothymidine at the time of coculture with the M. tuberculosis Ag-specific CD4⁺ T cells, we could not detect evidence of increased cell death in either DCs (9) or the T cells after coculture (as shown in this article). This approach was used to minimize the effect of direct HIV-infected T cells and so that the dominant effect studied in this study would be the DC–T cell interaction. Although the total numbers of CD4⁺ T cells surely are relevant, our data suggest that it is not only the total numbers of M. tuberculosis Ag-specific CD4⁺ T cells but also their functional quality and their polarization that are important for maintaining protection against TB. The mechanisms underlying the increased risk of developing TB in HIV patients with restored numbers of CD4 T cells after ART are unknown. Based on our findings, we speculate that if DCs are infected by HIV and M. tuberculosis that the subsequent DC–T cell interaction have a detrimental effect on T cell functionality. This effect could be the mechanistic trigger for 1) reactivating (or developing) TB in patients with normal CD4 counts and a recent seropositive HIV, 2) work in concert with the reduced CD4 count in patients with advanced disease (without ART), and 3) also be the reason why patients on ART, who never completely eradicate the HIV virus because of persistent viral reservoirs, still have an increased risk of developing TB.

TCR signaling in the presence of coinhibitory and absence of costimulatory signaling or with weak costimulatory signals during T cell activation can lead to T cell tolerance (14, 15). Ultimately, the balance between the costimulatory and coinhibitory signals decides the fate and the type of T cell response. We show that HIV–M. tuberculosis–coinfected DCs trigger M. tuberculosis Ag-specific CD4 T cell to express higher levels of coinhibitory molecules such as PD-1, CTLA-4, and Blimp-1. PD-1 and CTLA-4 are both important in regulating T cell responses and are effective targets for immunomodulatory treatment against certain forms of cancer, where their expression on T cells dampens the effector responses (31, 36). Mycobacteria can upregulate the expression of PD-1 and its ligands PD-L1/PD-L2 (37), and T cells and NK cells from TB patients have increased PD-1 expression (38). Our study suggests that HIV–M. tuberculosis coinfection enhances the PD-1 and PD-L1/PD-L2 pathways to further dampen the host M. tuberculosis immune responses. The transcriptional repressor Blimp-1 can maintain the homeostasis of effector T cells (39) and was shown to attenuate T cell proliferation and CD4 Treg functions, and its expression is enhanced in Ag-experienced T cells (40). Compared with the effect of M. tuberculosis single-infected DCs, we observed that HIV–M. tuberculosis–coinfected DCs induce more Blimp-1 expression in M. tuberculosis Ag-specific CD4 T cells. In naive T cells primed with HIV-pulsed DCs, high Blimp-1 expression was correlated with increased PD-1 and CTLA-4 expression (41). We now show that M. tuberculosis Ag-specific CD4 T cells are affected by HIV in a similar manner only during coinfection with M. tuberculosis, in which an increased expression of Blimp-1 correlates with an increased expression of PD-1 and CTLA-4, and that these signals are triggered through T cell activation by HIV–M. tuberculosis–coinfected DCs.

Costimulatory molecules are essential for effective T cell activation and differentiation (42). CD40–CD40L interplay is an important step in DC–T cell interaction, mediating efficient protection against mycobacteria (43), and CD40L blocking was shown to induces tolerance in T cells (44). There is evidence indicating that M. tuberculosis manipulates CD40–CD40L expression (45) and that CD40 is suppressed on infected cells of lepromatous patients and in patients with chronic mycobacterial infections (46, 47). CD40L expression on Th1 cells of TB patients has been correlated with the intensity of IFN-γ secretion (45), which supports our findings of reduced surface expression of CD40L on M. tuberculosis Ag-specific T cells correlating with suppressed levels of IFN-γ in HIV–M. tuberculosis coinfected cell culture supernatant. Previously, we showed that CD40 is suppressed on HIV–M. tuberculosis–coinfected DCs (9). We speculate that HIV further hampers CD40 expression and manipulates CD40L signaling during coinfection with M. tuberculosis to evade host defense mechanisms. CD28 on T cells, together with its ligands CD80 and CD86 on APCs, is one of the most pivotal costimulatory molecules for inducing functional T cell responses (48). Stimulation of T cells in the absence of CD28 results in impaired proliferation, reduced cytokine production, and altered Th1/Th2 balance (49). We observed lower expression of CD28 in M. tuberculosis Ag-specific CD4⁺ T cells activated with HIV–M. tuberculosis–coinfected DCs compared with when activated with M. tuberculosis single-infected DCs. Besides mycobacteria, many other pathogens like HIV, Leishmania donovani, and Bacillus anthracis exploit CD80/CD86-CD28 pathways for their persistence (50). CD28 cosignaling in turn triggers ICOS expression, which is an important regulator for CD4 T cell differentiation (51). The finding that ICOS expression in T cells from active TB patients correlated directly with IFN-γ secretion, in which ICOS ligation enhanced IFN-γ and, vice versa, IFN-γ also enhanced ICOS expression in a M. tuberculosis Ag-specific manner (52), suggests that this costimulatory pathway of activation is promoting the induction of a protective Th1 cytokine responses against intracellular pathogens. Our finding of lower expression of the costimulatory molecules CD40L, CD28, and ICOS in M. tuberculosis Ag-specific CD4⁺ T cells activated with HIV–M. tuberculosis–coinfected DCs suggests that this could be one mechanism by which HIV manipulates the M. tuberculosis–specific protective immune response and instead transforms these T cells into a dysfunctional and immune suppressed phenotype.

Our finding that expression of Treg markers was elevated in M. tuberculosis Ag-specific CD4⁺ T cells activated with HIV–M. tuberculosis–coinfected DCs is therefore particularly interesting and indicates a polarization toward suppressive T cells that can antagonize and dampen the protective immunity against M. tuberculosis. We have previously demonstrated that HIV–M. tuberculosis coinfection reduces the DC cytokine response and Ag-presenting capacity (9), and in this article we show that these dysfunctional DCs have a tolerogenic effect, as shown by their increased PD-L1/CD40 and PD-L1/CD86 ratio, which might promote the generation of Tregs. Tregs restrain the proliferation of naive and memory T cells, thereby suppressing pre-existing T cell immunity (53, 54). FOXP3 expression in CD4 T cells has been described as a marker for severity of HIV disease progression (55). The frequency of Tregs also increased in patients with active TB (19) and HIV-TB (20), and Tregs had the capacity to suppress IFN-γ production (19). These observations highlight Tregs as a pivotal player orchestrating the tipping point of control and lost control over intracellular pathogens. Our findings further identify that M. tuberculosis Ag-specific CD4⁺ T cells are also a target and delineate a possible course of interactions in which HIV–M. tuberculosis–coinfected DCs trigger certain numbers of these cells to shift into a regulatory phenotype, explaining the depressed IFN-γ secretion by M. tuberculosis Ag-specific CD4⁺ T cells that we observe. Our study also suggests that HIV takes advantage of tuning cosignaling molecules of M. tuberculosis Ag-specific CD4 T cells in such a way that instead of providing protection for the host, these T cells starts suppressing the host immune response and favoring survival of both HIV and M. tuberculosis.

The protective immunity against both TB and HIV is associated with the release of IFN-γ, TNF-α, and IL-2 by T cells after Ag stimulation, and it has been shown that individuals with advanced HIV infection have a reduced T cell secretion of these effector cytokines (11, 56, 57). We observed that HIV–M. tuberculosis–coinfected DCs induced significantly lower levels of IFN-γ and TNF-α and higher levels of TGF-β and IL-10 in the T cells compared with when the M. tuberculosis Ag-specific CD4⁺ T cells were activated with M. tuberculosis single-infected DCs. This ability of DCs to shift the T cells response from Th1 to immunosuppressive cytokine releasing could be part of the key mechanisms that M. tuberculosis exploits during HIV coinfection. This would explain why the effector capacity of M. tuberculosis Ag-specific CD4⁺ T cells activated with coinfected DCs had a considerably diminished capacity to activate M. tuberculosis–infected hMDM to control M. tuberculosis growth.

Our results suggest that HIV–M. tuberculosis–coinfected DCs can serve as the cellular link that impairs the quality and effector functions of M. tuberculosis Ag-specific CD4⁺ T cells and thereby be the mechanistic trigger for immunosuppression seen during TB-HIV coinfection. HIV–M. tuberculosis–coinfected DCs reduced Th1 effector immunity and skewed M. tuberculosis Ag-specific CD4⁺ T cells into a dysfunctional phenotype that made them less capable of controlling infection with M. tuberculosis (Fig. 8). This could explain why HIV-infected individuals even during ART are more prone to develop active TB.

We thank Julian Bess and the Biological Products Core of the AIDS and Cancer Virus Program, Leidos Biomedical Research, Inc., Frederick National Laboratory, Frederick, MD for providing HIV-1 virus preparations.

The authors have no financial conflicts of interest.