Multiprotein signaling complexes that regulate the maturation of the highly proinflammatory cytokine IL-1β were identified by Tschopp et al.[1,2] in the year 2002 as inflammasomes. Since then inflammasomes have been shown to assemble in response to various pathogen-associated molecular patterns (PAMPs) of viral (cytosolic viral DNA or RNA) [3–7] or bacterial origin (flagellin or type III secretion proteins)  as well as endogenous danger-associated molecular patterns (DAMPs) such as uric acid crystals, cholesterol crystals, beta-amyloid and extracellular ATP [9–15]. In macrophages, monocytes and dendritic cells nucleotide oligomerization domain-like receptors containing a Pyrin domain and the HIN-200 family members absent in melanoma 2 (AIM2) and interferon gamma-inducible protein 16 (IFI16) recruit the adaptor protein ASC (apoptosis-associated speck-like protein containing a caspase-recruitment domain) in response to PAMPs or DAMPs into a single ‘ASC speck’ of 0.5–1-μm size [16,17]. The aggregation of ASC is a hallmark of inflammasome activation accompanied by caspase activation, cell swelling, maturation and release of the proinflammatory cytokines IL-1β and IL-18 [18–20]. The subsequent demise of the cell has been termed pyroptosis to reflect the inflammatory nature of this form of cell death [21–23]. In addition to proinflammatory cytokines, ASC specks are themselves released into the extracellular space during pyroptosis . Extracellular ASC specks remain active and can propagate inflammation not only by maturating pro-IL-1β in the extracellular space but also by their phagocytosis by other cells [16,17,24].
Inflammation in the context of HIV-1 infection is a serious condition, which is thought to be linked to a higher morbidity in HIV-1-infected individuals than in the general population . Pyroptotic cell death was recently found to be responsible for the progressive loss of HIV-infected CD4+ T cells in secondary lymphatic tissues such as spleen, tonsils and gut-associated-lymphoid tissue [26,27]. For pyroptotic cell death to occur in tissue resident CD4+ T cells, the virus has to be transmitted to quiescent CD4+ T cells through virological synapses . In resting CD4+ T cells, the viral life cycle is arrested during the elongation step of reverse transcription and viral DNA fragments accumulating in the cytosol are sensed by the DNA sensor IFI16 [27,29]. In macrophages, HIV reverse transcriptase products accumulate in infected macrophages and also stimulate IFI16 . However, if IFI16 sensing of HIV in macrophages primarily controls HIV-1 replication or induces pyroptotic cell death has not been answered. Further evidence suggests, that HIV-1 can also prime for the NOD-like receptor protein 3 (NLRP3) inflammasome in macrophages . In this context, it is of interest to note, that an elevated level of IL-1β has been observed in tissues of HIV-infected patients (such as cerebrospinal fluid, lymph nodes, skin and bronchoalveolar epithelium) [27,32,33]. Moreover, IL-1β can induce replication of HIV in a promonocytic cell line . These results suggest that inflammasome activation may occur commonly in HIV-1 infection and IL-1β might be an important cytokine to promote inflammation in HIV-infected individuals. Moreover, why ‘non-AIDS’ related complications such as atherosclerosis , heart disease, chronic obstructive pulmonary disease and HIV-associated dementia are found with a greater prevalence in HIV-1-infected individuals is still largely unknown .
Surprisingly however, if inflammasome activation occurs in the peripheral blood of HIV-1-infected patients has not been studied. We speculated that ASC specks could be a marker of inflammation and inflammasome activation in HIV-infected individuals. We therefore investigated if circulating peripheral blood mononuclear cells (PBMCs) could be a source of ASC specks in HIV-1-infected individuals, and if ASC specks are detectable in the circulation of HIV-1 patients.
Material and methods
Blood sample processing
PBMCs were isolated from fresh blood using Ficoll (Biochrom, Berlin, Germany) density gradient centrifugation as described before [37,38]. Cells were washed three times with PBS and aliquots of 107 PBMCs were cryopreserved in heat-inactivated fetal calf serum (FCS) supplemented with 10% dimethyl sulfoxide (Merck, Darmstadt, Germany). All study patients were recruited between the years 2007 and 2015 at the HIV outpatient clinic of the Medizinische Hochschule Hannover (MHH) and gave written, informed consent prior to their participation. The study was approved by the local ethics committee (Votum der Ethikkommission der MHH no. 3150). Plasma HIV-1 RNA levels was measured using the COBAS TaqMan HIV-1 test (Roche Diagnostics, Mannheim, Germany). CD4+ and CD8+ T-cell counts were routinely determined by a flow cytometry-based assay using CYTO-STAT tetra CHROME (Beckman Coulter, München, Germany).
Frozen PBMCs were thawed from 10 healthy controls and 27 untreated HIV patients (Table 1) and washed in PBS containing 10% FCS before staining. Staining was done as described previously . Briefly, cells were fixed with 1% paraformaldehyde (Sigma-Aldrich, Schnelldorf, Germany) on ice for 5 min and then washed in staining medium [PBS, 0.1% sodium azide, 0.1% bovine serum albumin (BSA), 1% FCS]. Cell pellets were resuspended in permeabilization medium (PBS, 0.1% sodium azide, 0.1% BSA, 3% FCS, 0.1% saponin) and rabbit anti-ASC (clone N-15; cat. no. sc-22514-R, Santa Cruz, Heidelberg, Germany) was added to the samples and incubated for 90 min on ice. After centrifugation, supernatant was removed, and cells were incubated for 45 min on ice in permeabilization medium containing AlexaFluor 488 or AlexaFluor 647 goat anti-rabbit IgG (Life Technology, Darmstadt, Germany). Lineage-specific antibodies CD3 PerCP, CD19 PerCP (BD Biosciences, Heidelberg, Germany), CD14 APC-Cy7, CD16 PE-Cy7, CD56 V510 (BioLegend, Koblenz, Germany) and K57-PE (Beckman Coulter) were incubated together with the secondary antibody. Cells were washed in staining medium and flow cytometry was performed on a BD FACS Canto II (BD Biosciences). ACH-2 cells served as a positive control for the presence of Gag protein. ACH-2 cells are a HIV-1 latent T-cell clone with one integrated proviral copy and can be used as a positive control for intracellular p24 protein (Gag protein) (kindly provided by Christine Goffinet, TwinCore).
Freshly isolated PBMC were seeded at a density of 1 × 106 cells/ml in a 12-well plate and primed with 100-ng/ml LPS for 4 h at 37 °C. Cells were further stimulated either with 5-mmol/l ATP (Sigma-Aldrich) or with 10-μmol/l nigericin (Sigma-Aldrich) for another 20 min at 37 °C. ATP or nigericin act as a secondary stimulus to activate the inflammasome after LPS stimulation . The culture plate was spun at 1600 rpm for 5 min before harvesting cells for staining. Cells were fixed with 1% paraformaldehyde (PFA; Sigma-Aldrich) on ice for 5 min and stained for ASC as described above.
For the detection of human ASC specks in the plasma from 120 HIV-1-infected patients and 32 healthy controls (Table 2), an ASC ELISA (Lifespan Biosciences, Eching, Germany) was used according to the manufacturer's instructions. EDTA-plasma was collected and stored at −80 °C until the assay was done. A volume of 100 μl of plasma sample was used from each individual for the ELISA experiment.
Detection of ASC multimers in peripheral blood mononuclear cells from HIV-positive patients
To detect multimeric ASC, frozen PBMC samples from seven HIV-1-infected individuals and two healthy controls were thawed and lysed by mechanical disruption with 20-ga syringe (30 times) in lysis buffer (20-mmol/l 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid-potassium hydroxide, pH 7.5, 10-mmol/l KCl, 1.5-mmol/l MgCl2, 1-mmol/l EDTA, 1-mmol/l EGTA, 320-mmol/l sucrose). The cell lysate was centrifuged at 14 000 rpm, 15 min at 4 °C. Further, cell lysates were resuspended in CHAPS buffer (20-mmol/l HEPES–KOH, pH 7.5, 5-mmol/l MgCl2, 0.5-mmol/l EGTA, 0.1-mmol/l PMSF, 0.1% CHAPS) and then cross-linked with disccinimidyl suberate 2 mmol/l (Pierce, Darmstadt, Germany) prior to immunoblot analysis. Samples were kept on ice or at 4 °C throughout the procedure.
Proteins resolved on a 12% acrylamide SDS–PAGE were transferred on a nitrocellulose membrane (GE Life sciences, Freiburg, Germany) blocked with 5% skim milk in TBST buffer (tris buffer saline with tween-20) for 1 h at room temperature. Membranes were incubated with rabbit anti-ASC (Santa Cruz) at a dilution of 1 : 1000 in 5% skim milk in TBST buffer overnight at 4 °C. Excess antibody was removed by washing five to six times with TBST buffer following incubation with secondary goat anti-rabbit IgG HRP-conjugated antibody (Bio-Rad, München, Germany) at 1 : 5000 dilution in 2.5% skim milk for 1 h at room temperature. The washed membrane was exposed to chemiluminescent substrate (Clarity ECL; Bio-Rad, München, Germany) for signal detection.
Unpaired t test or Spearman correlations were performed as appropriate. Multiple comparisons were analyzed by one way analysis of variance, followed by Kruskal–Wallis test. GraphPad Prism software (GraphPad Software, San Diego, California, USA) was used for statistical evaluation of the data. A P value less than 0.05 was considered significant.
ASC specks are found in peripheral blood CD14++CD16− classical monocytes from HIV-infected patients
Detection of ASC forming inflammasomes in PBMCs from patients was hampered until recently due to the lack of sensitive tools measuring their formation on a cellular level. Under resting stage, the inflammasome adaptor ASC is distributed uniformly over the cell. Upon inflammasome activation, ASC rearranges into a large intracellular ASC speck. Sester et al. recently developed a reliable method to visualize the re-distribution of ASC by flow cytometry on a single cell level . We employed Sester's methods and confirmed them by using LPS-primed PBMCs stimulated with the inflammasome activators ATP or nigericin (Fig. 1a, of the stimulated PBMCs only gated CD14++CD16− monocytes after exclusion of T cells, B cells and natural killer cells are shown). As expected, we observed a marked increase in speck forming CD14+ monocytes under inflammasome activating conditions, as compared with resting conditions (Fig. 1a). The inflammasome forming cells were observed via reduction in the ASC fluorescent pulse width, with a concomitant increase in the ASC pulse area (lower right gate, low W : A ratio). Resting cells had high ASC fluorescent pulse width and low ASC fluorescent pulse area (upper right gate, high W : A ratio).
As we had frozen PBMCs samples of untreated HIV-1-infected patients, we next evaluated whether these frozen samples could be employed without false positive ASC speck formation due to the freeze-thaw cycle via flow cytometry for ASC speck formation. We therefore compared PBMCs isolated freshly from healthy donors to PBMCs that had been subjected to a freeze-thaw cycle. No false positive events were observed in comparison (Supplemental Fig. 1, http://links.lww.com/QAD/B188).
We next tested PBMCs from healthy donors for ASC speck formation and found, as expected, no evidence of ASC-forming inflammasomes. In contrast, in our cohort of HIV+ patients, we found a small, but detectable fraction of cells forming an ASC-containing inflammasome (Fig. 1b). In line with results by the group of Munoz-Arias et al., who reported peripheral circulating CD4+ T-cells resistant to pyroptosis, further analysis excluded that the specking cells were CD4+ T lymphocytes (Supplemental Fig. 2, http://links.lww.com/QAD/B188), but instead fell exclusively within the fraction of CD14++CD16− classical monocytes (Fig. 1c and Supplemental Fig. 3, http://links.lww.com/QAD/B188). Using immunoblotting, we confirmed that ASC specks detected by flow cytometry in HIV-infected patients were indeed multimeric aggregates of ASC (Fig. 1d). Together these data show that circulating classical monocytes in HIV-1-infected individuals form ASC containing inflammasomes.
HIV-infected antiretroviral therapy-naive patients with higher viral load and lower CD4+ T-cell numbers have higher numbers of ASC-forming inflammasomes within the CD14++CD16− classical monocyte subset
We next compared the percentages of the ASC speck containing CD14++CD16− (of the CD14++CD16− monocytes) from healthy donors (n = 10) and HIV-1-infected ART-naive patients (n = 27). We found a significant difference between the groups (P = 0.0121) (Fig. 2a) with a higher percentage of ASC speck positive CD14++CD16− in the HIV-1-infected ART-naive patients. We then asked, if HIV-1 RNA viral load would influence the capacity of monocytes to form ASC specks. For this, we calculated the median viral load (median: 88 800 copies/ml) of our cohort of HIV-infected patients, and compared the percentage of ASC speck+ CD14++CD16− monocytes below and above this median. This analysis revealed a significant difference with more monocytes within the higher viral load group forming ASC specks (P = 0.032) (Fig. 2b). Clinically, HIV-1-infected individuals are commonly grouped based on the number of peripheral CD4+ T-cell counts into the U.S. Centre for Disease Control and Prevention classification (CDC). Based on the lowest documented CD4+ cell counts HIV patients can thereby be classified in three groups: CDC group 1 with at least 500 CD4+ cell counts/μl, CDC group 2 with 200–499 CD4+ cell counts/μl and CDC group 3 with less than 200 CD4+ cell counts/μl . The percentage of ASC speck+ monocytes between different CDC groups did not reveal statistical significant differences. But, HIV patients with CD4+ T-cells counts below 500/μl (i.e. patients of CDC groups 2 and 3 combined) had significantly more pyroptotic CD14++CD16− monocytes when compared with HIV patients with CD4+ T-cell counts greater or equal than 500/μl (CDC group 1) (Fig. 2c). We further found no correlation of ASC speck+ monocytes with C-reactive protein (CRP) as a clinical marker of infection, and/or inflammation (data not shown) nor the cell death marker lactate dehydrogenase (LDH) (Fig. 2d). Together, these data show that lower CD4+ T-cell counts and higher HIV-1 viral load correlate with more pyroptotic ASC speck+ CD14++CD16− monocytes within our cohort of HIV-1-infected individuals.
ASC speck+ monocytes do not productively replicate HIV-1 virus
As we had observed a trend to a greater number of pyroptotic monocytes within HIV-1 patients with a higher viral load, we were next interested if the pyroptotic monocytes were productively replicating HIV. HIV Gag KC57 staining of the specking monocytes revealed only a small fraction of these cells positive for HIV Gag (Fig. 3), which would be indicative of active HIV replication. However, as most of the ASC speck+ cells stained negative for Gag, we conclude, that HIV replication is not the dominant cause of inflammasome activation within the pyroptotic monocytes of HIV-1+ patients.
ASC specks are released into the circulation in HIV-1-infected patients
As multimeric ASC specks released into the extracellular space were shown to propagate inflammation through their uptake by nonpyroptotic cells , we hypothesized that ASC specks within the bloodstream of HIV-infected individuals might convey inflammation to distant sites and thereby contribute to a chronic inflammatory state. We therefore analyzed the plasma of HIV-infected patients with an anti-ASC ELISA. Synovial fluid drawn from the joints of patients with gouty arthritis served as a positive control, as uric acid crystals are a prototypic stimulus of the NLRP3 inflammasome . None of the tested healthy donor's plasma samples were positive in the ASC ELISA, however, a small number of the tested HIV plasma samples were (Fig. 4). We were then interested, if ASC release correlated with clinical parameters of cell death (LDH) or inflammation (CRP). However, of the few HIV plasma samples testing positive in the ASC ELISA only one had elevated CRP and one had elevated LDH levels. In addition, due to the small sample size, we could not find a significant correlation between the release of ASC and CDC status, or viral load (data not shown).
Recent groundbreaking reports have revealed that a major percentage of CD4+ T cells abortively infected with HIV-1 die due to pyroptosis in secondary lymphatic organs, indicating a role for the inflammasomes in HIV-1 infection [26,27,29]. Our study now extends the connection between HIV-1 infection and inflammasome activation beyond CD4+ T cells. We have identified peripheral blood classical CD14++CD16− monocytes as important inflammasome activated cells in HIV infection. Although our study does not define the specific sensor protein upstream of inflammasome activation, this is the first report demonstrating ASC speck formation, and the release of these particles into the circulation of a non-autoinflammatory disease such as HIV-1. These findings might be of importance regarding systemic inflammation in HIV-1 infection [25,41].
As higher viral loads correlated with a greater percentage of pyroptotic to resting monocytes, we speculated that HIV-1 itself might cause their pyroptotic demise. However, the majority of monocytes positive for ASC specks were negative for Gag, therefore leaving the exact mechanism(s) upstream of ASC speck formation unanswered. Further studies will have to address, if ASC speck positive monocytes in HIV-1 positive patients are abortively or latently infected with HIV-1. As HIV-1 does not produce Vpx to counteract the viral restriction factor SAM domain and HD domain-containing protein1 , monocytes productively infected by HIV-1 are likely rather scarce. Moreover, the Gag protein is only detectable during the late phase of HIV-1 replication, and thus, intracellular Gag staining performed in our study is not sufficient to detect abortively or latently infected cells. Nevertheless, previous small interfering ribonucleic acid experiments targeting the inflammasome sensors NLRP3, retionic acid inducible gene 1 and AIM2 showed, that HIV-1-infected human monocytes were preferentially deficient in IL-18 secretion when NLRP3 was knocked down , suggesting that NLRP3 is the dominant sensor protein for inflammasome formation in HIV-1 infection.
Our finding that ASC speck positive monocytes are significantly increased in HIV patients with lower compared to higher CD4+ T-cell counts may also be explained by the increase of nonphysiological bacterial translocation from the gut in CD4+ T-cell depleted HIV patients . Although we do not have data regarding endotoxin levels in our cohort, it is conceivable that increased amounts of LPS and other bacterial products could engage the alternative inflammasome pathway recently described to operate in human monocytes . As in our cohort we found no correlation of ASC specks to existing markers of inflammation or cell death, such as CRP or LDH, respectively, we currently assume that the pyroptotic cell death of classical CD14++CD16− monocytes in HIV-1-infected patients may be an independent marker of subclinical inflammation.
We observed a surprisingly lower frequency of patients positive for the presence of free ASC in the plasma (10% positive), compared with a greater frequency of HIV-1+ patients positive for ASC specks within monocytes (55% positive). These seemingly contradicting results may be due to the uptake of ASC specks by the reticuloendothelial system, due to the resistance of the monocyte/macrophage lineage to the cytopathic effect of HIV-1  or due to the uncoupling of inflammasome activation and pyroptotic cell death, which was observed in monocytes during alternative inflammasome activation by LPS .
The current study now identifies CD14++CD16− monocytes as the key inflammasome activated cell type within the circulation of a subset of HIV-1-infected individuals and demonstrates, that ASC specks are additionally released into the blood stream of HIV-1-infected patients. The long-term consequences of ASC specks released into the circulation are currently unknown. Further studies are warranted, whether antiretroviral treatment will lower the number of released ASC specks and if degenerative diseases or cardiovascular disease in long-term ART-treated HIV-infected individuals are linked to inflammasome activation.
We thank all healthy donors and HIV patients who participated in this study. We thank the funding agencies, German Research Foundation grants DFG BO 4325/1-1 (to L.B.), the HiLF of the MHH (to L.B.), Priority Programme 1923 (to E.L.) and ERC InflammAct (to E.L.) as well as DZIF TTU HIV 04.810 and 04.811 (to R.E.S.) for supporting this study. We thank Prof Christine Goffinet, TwinCore, MHH, Hannover for providing the ACH-2 cells.
Authorship contributions: L.B., R.E.S., F.A. and N.M. conceived and designed the experiments. F.A., N.M. and L.B. carried out the experiments and data analysis. E.L. and B.S.F. provided reagents and gave experimental advice. F.A. and G.A. selected patient's samples and collected the clinical data of the patients. L.B., F.A., B.S.F. and N.M. wrote the article.
Conflicts of interest
There are no conflicts of interest.
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* Fareed Ahmad and Neha Mishra contributed equally to the article.