HIV infection deregulates innate immunity to malaria despite combination antiretroviral therapy
Finney, Constance A.M.a; Ayi, Kodjoa; Wasmuth, James D.b; Sheth, Prameet M.a; Kaul, Ruperta,c; Loutfy, Mona R.d; Kain, Kevin C.a,*; Serghides, Lenaa,*
aSandra Rotman Centre for Global Health, University Health Network, University of Toronto, Toronto, Ontario
bDepartment of Ecosystem and Public Health, Faculty of Veterinary Medicine, University of Calgary, Calgary
cDepartment of Immunology
dWomen's College Research Institute, Women's College Hospital, University of Toronto, Toronto, Canada.
*Kevin C. Kain and Lena Serghides have contributed equally as senior authors.
Correspondence to Lena Serghides, Sandra Rotman Centre for Global Health, TMDT, Suite 10-359, 101 College Street, Toronto, ON M5G 1L7, Canada. E-mail: email@example.com
Received 13 April, 2012
Revised 28 September, 2012
Accepted 9 October, 2012
A portion of these data was presented previously at the following meetings: the American Society of Tropical Medicine and Hygiene (2011), the Ontario HIV Treatment Network (2011), and the Canadian Association of HIV Research (2012).
Supplemental digital content is available for this article. Direct URL citations appear in the printed text and are provided in the HTML and PDF versions of this article on the journal's Website (http://www.AIDSonline.com).
Objective: Malaria and HIV-1 adversely interact, with HIV-positive individuals suffering higher parasite burdens and worse clinical outcomes. However, the mechanisms underlying these disease interactions are unclear. We hypothesized that HIV coinfection impairs the innate immune response to malaria, and that combination antiretroviral therapy (cART) may restore this response. Our aim was to examine the innate inflammatory response of natural killer (NK), natural killer T (NKT), and γδ T-cells isolated from the peripheral blood of HIV-infected therapy-naive donors to malaria parasites, and determine the effect of cART on these responses.
Methods: Freshly isolated peripheral blood mononuclear cells from 25 HIV-infected individuals pre-cART (month 0) and post-cART (months 3 and 6), and HIV-negative individuals at matched time-points, were cultured in the presence of Plasmodium falciparum parasitized erythrocytes. Supernatants and cells were collected to assess cytokine production and phenotypic changes.
Results: Compared to HIV-negative participants, NKT, NK, and γδ T-cell subsets from participants with chronic HIV infection showed marked differences, including decreased production of interferon γ (IFNγ) and tumor necrosis factor (TNF) in response to malaria parasites. IFNγ production was linked to interleukin-18 receptor (IL-18R) expression in all three cell types studied. Six months of cART provided partial cellular reconstitution but had no effect on IL-18R expression, or IFNγ and TNF production.
Conclusion: These data suggest that HIV infection impairs the inflammatory response of innate effector cells to malaria, and that the response is not fully restored within 6 months of cART. This may contribute to higher parasite burdens and ineffective immune responses, and have implications for vaccination initiatives in coinfected individuals.
Malaria and HIV-1 (HIV) are major global health priorities. Coinfection with these pathogens results in accelerated disease progression and more severe clinical manifestations [1–8]. Malaria increases HIV plasma viral load [1,3,7] and causes a temporary decrease in CD4+ T-cell counts; conversely, HIV infection is associated with higher parasite burdens and an increased risk of clinical and severe malaria [1,7,9–16]. The mechanisms by which HIV increases susceptibility to malaria are not fully understood. Furthermore, most studies examining the effect of HIV on malaria do so in the absence of antiretroviral therapy.
Innate immune responses are the first line of defence in a malaria infection and can dictate not only the outcome of infection but also the development of adaptive immunity , which is of particular importance to vaccine initiatives. A balanced inflammatory response characterized by an early pro-inflammatory and a secondary immunoregulatory response is required to adequately control parasitemia while limiting host pathology [17,18]. Natural killer (NK) T cells, NK cells, and γδ T cells have all been implicated in the innate response to malaria [19–21]. HIV disease progression is associated with the destruction of CD4+ T cells, but other cell types including γδ T cells, NK cells, and NKT cells have been reported to decrease in number and alter in function in HIV-infected individuals [22–24]. Typically, HIV infection leads to a dysregulation of inflammatory responses, but both enhanced and reduced responses have been reported in the context of malaria coinfection [25–29].
Our aim was to elucidate the impact of HIV infection (both untreated and treated) on innate immune responses to malaria, which are more relevant in the nonimmune individuals (travelers and young children) who are more likely to suffer from severe disease. We hypothesized that coinfection with HIV compromises host innate inflammatory responses to malaria and that this could be responsible for the higher malaria parasite burdens and worse disease outcome observed in coinfected individuals. We further hypothesized that innate immune responses can be restored with cART (combination antiretroviral therapy). Here, we show that HIV-infected participants exhibit alteration in subsets of NK, NKT, and γδ T cells, and a loss of malaria-specific cytokine-producing abilities, that is only partially corrected by cART. The cytokine production defect may contribute to increased parasite burdens and ineffective immune responses observed in coinfected individuals.
Participants and methods
Study population and ethics statement
The University Health Network and University of Toronto Ethics Review Boards approved this study. Study participants gave written informed consent. HIV-1-infected [HIV(+)] treatment-naive participants were recruited at the Maple Leaf Medical Clinic, Toronto, Canada. HIV(+) participants were chronic progressors (HIV-infected for >1 year, with CD4+ T-cell count decline of >50 cells/μl/year). Venous blood samples were collected just prior to the initiation of cART therapy (M0), at 3 months (M3), and at 6 months (M6) post-cART.
Controls were HIV-negative [HIV(−)], recruited in the same demographic area with a similar, but not identical, age and sex profile [female participants were more numerous in the HIV(−) group]. Each pair of HIV(−) and HIV(+) samples was collected at the same time and processed identically. When possible, each HIV(+) donor was matched to the same HIV(−) individual for all three time-points.
Viral loads and CD4+ T-cell counts for all patients can be found in Table 1.
All experiments were performed with freshly isolated peripheral blood mononuclear cells (PBMCs), purified from blood samples using Ficoll gradients. PBMCs were plated at 1 × 106/ml of media (RPMI-1640, 10% fetal calf serum, minimal nonessential amino acids, sodium pyruvate, 2β-mercaptoethanol, gentamycin) in 24-well plates and cultured with 3 × 106 parasitized erythrocytes (∼10% parasitemia) as described by , and equivalent hematocrit levels of uninfected red blood cells (uRBCs), media, or phorbol-12-myristate-13-acetate (PMA; 2.5 pg/ml)/ionomycin (250 pg/ml; Sigma, Oakville, Canada) for 4 days. A 3 parasitized erythrocyte : 1 PBMC ratio represents 15 000 parasites/μl  and was chosen because it recapitulates a number of clinical signs associated with malaria disease [30–32]. Supernatants were collected at 24, 48, and 96 h for cytokine analyses. Cells were collected prior to culture and at 48 h for analysis of cell surface markers, and at 72 h for analysis of cytokine production by intracellular staining. Cells were incubated with brefeldin A (3 μg/ml) 8 h prior to intracellular cytokine staining.
For interleukin-18 receptor (IL-18R) blockade experiments, PBMCs were preincubated with blocking monoclonal antibodies at three concentrations [antihuman IL-18Rα clone B-E43 low = 0.05 μg/ml, medium = 0.5 μg/ml, high concentration = 1 μg/ml (Cell Sciences, Canton, Massachusetts, USA); and antihuman IL-18Rβ clone 132016 low = 0.5 μg/ml, medium = 1 μg/ml, high = 5 μg/ml (R&D Systems, Burlington, Canada)] for 1 h, and then exposed to stimulus for 24 h (PMA/ionomycin) or 48 h (parasitized erythrocytes).
Plasmodium falciparum (ITG strain) was cultured as previously described . Cultures were treated with Mycoplasma-Removal Agent (MP Biochemicals, Solon, Ohio, USA) and routinely tested negative for Mycoplasma by PCR. Cultures were synchronized using alanine lysis and trophozoite-stage cultures were used in all assays. Parasites were used at a ratio of 3 parasitized erythrocyte : 1 PBMC.
Culture supernatants were collected at 24, 48, and 96 h, and frozen at −80oC. Interferon γ (IFNγ), tumor necrosis factor (TNF), IL-2, and IL-6 were determined using the Human Th1/Th2 cytometric bead array (BD Bioscience, Mississauga, Canada). IL-10 was assessed by ELISA (R&D Systems and BD Bioscience). For the IL-18R blockade experiments, IFNγ and TNF were assessed by ELISA.
Multicolor flow cytometry
To determine cellular phenotypes, PBMCs were stained with conjugated monoclonal antibodies to CD14, CD4, CD8, CD3, CD56, γδ, and IL-18R (BioLegend, San Diego, California, USA). BD FACS lysing solution (BD Biosciences) was used to fix cells and lyse contaminating RBC. To assess intracellular cytokine production, PBMCs were stained for surface markers, fixed and permeabilized using the cytofix/cytoperm kit (BD Biosciences), and stained with anti-TNF and/or anti-IFNγ antibodies (BioLegend). Cells were analyzed using a FACS-Canto, or LSRII (BD Biosciences) within 24 h after fixing, and data were analyzed using FlowJo (Tree Star, Ashland, Oregon, USA). A minimum of 100 000 CD3+ cells were collected. NK cells were defined as CD14−CD3−CD56+, NKT cells were defined as CD14−CD3+CD56+, γδ T cells were defined as CD14−CD56−CD3+γδ+, and monocytes were defined as CD3−CD14+. The percentages of NK, NKT, and monocytes were calculated by dividing the number of cells in each cell subtype gate by the total cells (excluding doublets). The percentage of γδ T cells was calculated by dividing the number of cells in the γδ T-cell gate by the total CD3+ cells.
To compare IL-18R expression levels between donors, we calculated the IL-18R mean fluorescence index (MFI). This is defined as the ratio of the IL-18R mean fluorescence intensity and the mean fluorescence intensity value for the fluorescence minus one (FMO) control. This approach allows for comparison of multiple test samples within a group and between different groups.
The Wilcoxon matched pair test was used to compare HIV(+) to HIV(−) values. Kruskal–Wallis test with Dunn's multiple comparisons posttest was used to assess differences between groups in the IL-18R blockade experiment. For malaria-specific cytokine production, production in the presence of uRBCs was subtracted from production in response to parasitized erythrocytes.
HIV infection is associated with a blunted innate inflammatory response to malaria
We investigated the malaria-induced cytokine response in the context of HIV infection and examined the effects of cART on this response. Culture supernatants were collected at 24, 48, and 96 h and tested for levels of IFNγ, TNF, IL-2, IL-6, and IL-10. Cytokine levels measured in wells stimulated with uninfected RBCs were subtracted from the cytokine levels measured in wells stimulated with parasitized erythrocytes to determine malaria-specific cytokine production. The time-points displayed in Table 2A and Supplemental Figure 1, http://links.lww.com/QAD/A270 for each cytokine were selected based on the kinetics of the response and the number of responding HIV(+) and HIV(−) donor pairs. The 48-h time-point was optimal for the comparison of TNF, IFNγ, and IL-10. IL-2 responses were minimal until the 96-h time-point. IL-6 responses peaked early at the 24-h time-point.
After 48 h of culture, malaria-specific levels of IFNγ and TNF were significantly lower in HIV(+) compared to HIV(−) cultures (Table 2A, Supplemental Fig. 1a-b, http://links.lww.com/QAD/A270, IFNγ: P = 0.02, TNF: P = 0.03). cART treatment (M6) did not correct the defect in malaria-specific IFNγ and TNF production in HIV(+) cultures (Table 2A, Supplemental Fig. 1a-b, http://links.lww.com/QAD/A270).
PBMCs from HIV(+) donors also produced significantly lower levels of IL-2 compared to PBMCs from HIV(−) donors at 48 h (data not shown) and 96 h of culture with parasitized erythrocytes (Table 2A, Supplemental Fig. 1c, http://links.lww.com/QAD/A270, P = 0.001). A recovery in malaria-specific IL-2 production was observed with HIV(+) PBMCs collected at M6 (Table 2A, Supplemental Fig. 1c, http://links.lww.com/QAD/A270).
Levels of malaria-specific IL-6 were similar in both HIV(−) and HIV(+) cultures at all time-points (Table 2A, P > 0.05).
Unlike the pro-inflammatory cytokines above, levels of the anti-inflammatory cytokine IL-10 were significantly higher in HIV(+) cultures following 48 h of culture with parasitized erythrocytes (Table 2A, Supplemental Fig. 1d, http://links.lww.com/QAD/A270, P = 0.002). This difference disappeared at M6 post-cART (Table 2A, Supplemental Fig. 1d, http://links.lww.com/QAD/A270).
In summary, malaria-induced cytokine production was perturbed in HIV(+) donors as shown by a defect in TNF, IFNγ, and IL-2 production, and an increased IL-10 response. cART corrected IL-2 and IL-10 responses to levels similar to those seen in HIV(−) controls, but did not correct the defects in TNF or IFNγ production.
HIV infection is associated with altered populations of natural killer, natural killer T, and γδ T cells, with selective restoration following combination antiretroviral therapy
To further understand the defect in malaria-induced IFNγ and TNF responses observed with HIV(+) PBMCs, we investigated innate immune cells that produce these cytokines and that have been implicated in early malaria responses, specifically NK, NKT and γδ T-cells.
Treatment-naive HIV(+) donors had significantly lower levels of NK cells (%CD14−CD56+CD3− of total cells) compared to HIV(−) controls; however, at M6 post-cART, this difference was diminished (Table 3, Supplemental Fig. 2a, http://links.lww.com/QAD/A270). The recovery was primarily due to an increase in the CD56lo NK cells, a principally cytotoxic subset (Table 3, Supplemental Fig. 2b, http://links.lww.com/QAD/A270). Although recovery in the less abundant, largely cytokine producing, CD56hi NK cells did not reach control levels (Table 3, Supplemental Fig. 2c, http://links.lww.com/QAD/A270), the subset did respond to cART treatment, as percentages increased by M6 (Supplemental Fig. 2d, http://links.lww.com/QAD/A270).
As with NK cells, the percentage of NKT cells (%CD14−CD56+CD3+ of total cells) was lower in HIV(+) treatment-naive donors compared to HIV(−) controls, but increased with cART to reach levels similar to HIV(−) controls at M6 (Table 3, Supplemental Fig. 3a, 3d, http://links.lww.com/QAD/A270). NKT cells from HIV(−) donors were mostly CD8−CD4−, whereas those from HIV(+) donors were primarily CD8+ (Table 3, Supplemental Fig. 3b-c, http://links.lww.com/QAD/A270). This difference remained at M6. The levels of CD4+ NKT cells were low and did not differ between HIV(−) and HIV(+) donors pre-cART or post-cART (data not shown).
The percentage of γδ T cells (%γδ+CD3+CD56−CD14− of CD3+CD56−CD14− cells) did not differ between HIV(+) and HIV(−) donors, and was unaffected by cART (Table 3, Supplemental Fig. 4a, http://links.lww.com/QAD/A270). γδ T cells from HIV(+) donors contained a smaller percentage of CD4+ cells than those of HIV(−) donors (Table 3, Supplemental Fig. 4b, http://links.lww.com/QAD/A270); this defect was still evident at M6 (Supplemental Fig. 4c, http://links.lww.com/QAD/A270).
The ability of NK, NKT, and γδ T cells to produce IFNγ and TNF following stimulation with parasitized erythrocytes was examined using intracellular cytokine staining. Although both NK and NKT cell percentages appeared to recover with cART and γδ T-cell percentages were similar between HIV(+) and HIV(−) donors, malaria-specific IFNγ and TNF production from NK, NKT, γδ T cells from HIV(+) donors was minimal compared to matched controls, and remained minimal at M6 (Table 2B, Supplemental Fig. 5a-c, http://links.lww.com/QAD/A270).
To determine whether PBMCs from HIV(+) donors were able to produce IFNγ and TNF to a more general stimulus, PBMCs were cultured in the presence of PMA/ionomycin prior to intracellular cytokine staining. The percentage of IFNγ+ and TNF+ lymphocytes was similar between HIV(+) and HIV(−) donors, and cART had no effect on the cytokine production (Supplemental Fig. 5e, http://links.lww.com/QAD/A270). Cell subset analysis was not performed because PMA/ionomycin stimulation strongly impacts certain cell surface markers .
Monocytes are also an important component of the innate immune response to malaria. Similarly to NK, NKT, and γδ T cells, IFNγ and TNF production from monocytes was lower in HIV(+) donors compared to controls, pre-cART and post-cART (Supplemental Fig. 5d, http://links.lww.com/QAD/A270). However, due to limited sample volume, a single time-point was selected for the analysis of cell-specific cytokine responses that correlated with the highest levels of IFNγ in the culture supernatants. This time-point was not optimal for monocyte-specific responses; thus, we have limited the interpretation of the monocyte-specific data.
Interleukin-18 receptor expression is decreased in innate cells from HIV(+) donors
The human IL-18 receptor (IL-18R) is expressed on NK, NKT, and γδ T cells and has been identified as a marker of a pro-inflammatory phenotype . IL-18R is upregulated in inflammatory environments, leading to increased IFNγ production .
Both the percentage of IL-18R+ and the level of IL-18R expression were significantly lower in resting NK, NKT, and γδ T cells from HIV(+) donors compared to HIV(−) controls prior to culture (Table 4, Supplemental Fig. 6a-c, http://links.lww.com/QAD/A270). cART treatment did not improve the percentage or the expression levels of IL-18R on any cell type (Table 4, Supplemental Fig. 6a-c, http://links.lww.com/QAD/A270). Similar observations were made for cells cultured with parasitized erythrocytes for 48 h (Supplemental Fig. 7, http://links.lww.com/QAD/A270). Monocyte IL-18R expression did not differ between resting HIV(+) and HIV(−) donors (Supplemental Fig. 6d, http://links.lww.com/QAD/A270).
Blocking interleukin-18 receptor in HIV(−) donors reduces parasitized erythrocyte-specific interferon γ but not tumor necrosis factor production
Parasitized erythrocyte-induced IFNγ, but not PMA/ionomycin-induced IFNγ, was enriched in IL-18R-expressing NK, NKT, and γδ T cells (Fig. 1a). The majority of parasitized erythrocyte-induced IFNγ+ cells were IL-18R+, whereas the majority of PMA/ionomycin-induced IFNγ+ cells were IL-18R−. Blocking the IL-18R using two different antibodies prior to stimulation resulted in significantly lower levels of parasitized erythrocyte-specific IFNγ (Fig. 1b-top left panel), but not TNF (Fig. 1b-top right panel) production. IL-18R blockade did not affect cytokine production from cells exposed to PMA/ionomycin (Fig. 1b-lower panels).
In malaria infection, the innate immune system plays an important role in controlling parasite replication and determining disease outcome, in part via the production of cytokines like IFNγ . Here we show that malaria-induced innate immune responses are altered in chronic HIV infection.
PBMCs from treatment-naive, viremic, HIV(+) donors showed a defect in parasitized erythrocyte-induced IFNγ, TNF, and IL-2 production, and an enhanced IL-10 response compared to HIV(−) controls. Following 6 months of cART, which rendered almost all donors aviremic, parasitized erythrocyte-induced IL-2 and IL-10 responses recovered to levels similar to those seen in controls; however, the defect in IFNγ and TNF production was still present. Parasitized erythrocyte stimulation induced IFNγ and TNF production in NK, NKT, and γδ T cells from HIV(−) but not from HIV(+) donors, and this defect persisted despite cART. NK, NKT, and γδ T cells from HIV(+) donors could produce IFNγ and TNF in response to PMA/ionomycin, suggesting that the cells were not anergic and were able to respond to this stimulus but not to parasitized erythrocytes.
Data from models of malaria and experimental infections in human volunteers suggest that early IFNγ responses are associated with rapid control of parasitemia but at the cost of developing clinical symptoms, whereas high IL-10 levels are associated with high parasitemia but reduced clinical symptoms [17,37]. Chronic HIV infection has been associated with perturbations in cytokine levels, including decreased IFNγ, IL-2, and IL-12 production, and increased IL-10, IL-4, and TNF production [38–40]. Investigation of cytokine responses in the context of HIV-malaria coinfection is limited but, as in our study, impaired malaria-induced IFNγ responses have been reported [25–27]. In response to parasitized erythrocytes, we have now also observed reduced IL-2, enhanced IL-10, and reduced TNF production in PBMCs from HIV(+) donors. In addition, we are the first to report on the effect of cART on cytokine production in the context of HIV-malaria coinfection. Six months of cART corrected the IL-2 and IL-10 differences between HIV(+) and HIV(−) PBMCs, but not the defects in IFNγ and TNF responses.
Several mechanisms could contribute to defective IFNγ and TNF responses. High levels of IL-10 can inhibit TNF and IFNγ induction. However, levels of IFNγ and TNF were not restored at M6, despite normalization of IL-10 responses, suggesting that IL-10 is unlikely to be a major contributor.
Reduced IL-18R levels can also limit IFNγ production. In HIV(−) donors, IL-18R blockade reduced parasitized erythrocyte-induced IFNγ production. It has been previously reported that expression of IL-18R is required for NK cells to produce IFNγ in response to malaria . A correlation between decreased expression of IL-18R and defective IFNγ responses has also been reported in obese diabetic and in cancer patients . In addition, NKT and NK cells from IL-18R-deficient mice showed reduced IFNγ responses to bacterial infection . Therefore, lower IL-18R expression on NK, NKT, and γδ T cells may reduce their sensitivity to IL-18, and thus, limit their ability to produce IFNγ in response to malaria.
This is the first study to report lower IL-18R expression on NK, NKT, and γδ T cells, but not monocytes, in HIV infection, and complements the reported shift in the ratio of IL-18R to ST2L on lymphocytes from HIV(+) donors [23,35]. Our results indicate that changes in IL-18R expression could account for defects in IFNγ production and may relate to the defective IFNγ responses we observed in HIV(+) individuals.
Several baseline cell-type differences were detectable between HIV(+) and uninfected controls that may have contributed to altered cytokine responses to parasitized erythrocytes. NK cells are early responders to malaria infection and serve to initiate the inflammatory response via the production of IFNγ [19,43]. In agreement with published observations, treatment-naive HIV(+) donors had lower levels of NK cells [44,45], particularly in the CD56hi subset that serves as an early source of cytokines (including IFNγ and TNF) . Differences in NK cell numbers were less obvious at M6; however, the CD56hi subset did not fully recover to HIV(−) levels. Studies have reported recovery of NK subsets and cytotoxicity function following cART [47,48], but also, as we have observed, sustained impairment of innate immune function (persistent defect in IFNγ production) even after viremic suppression and NK-cell reconstitution [49,50].
NKT cells can modulate immune responses by rapid production of an array of cytokines, influencing innate and adaptive immune responses . As previously reported [14,52], we observed reduced baseline numbers of NKT cells in our HIV(+) donors that increased with cART, though rapid increases in NKT numbers have not been reported in all studies [12,15,53]. In HIV(−) participants, NKT cells were primarily CD4−CD8−, whereas in HIV(+) donors they were predominantly CD8+, a phenotype observed during viral infections . CD8+ NKT cells were still predominant at M6. Both CD8+ and CD4−CD8− NKT cells have been shown to largely produce Th1 cytokines [55–57]; however, NKT cells from HIV(+) individuals display severe functional impairment .
Although the physiological role of NKT cells in blood-stage malaria is unclear , we observed a strong malaria-specific NKT cytokine response from HIV(−) participants. In contrast, we observed very little IFNγ or TNF production by NKT cells from HIV(+) participants, a defect that did not improve at M6. Functional impairment of NKT cells persisting despite cART has been reported , although a return of the ability to produce IFNγ after 12 months of cART has also been observed .
γδ T cells represent an early defense mechanism against invading pathogens by recognizing nonpeptidic microbial antigens without antigen processing and major histocompatibility complex restriction [20,59,60]. Phosphoantigens from P. falciparum can stimulate γδ T cells , and the percentage and absolute number of γδ T cells increase in P. falciparum infection [62,63]. Along with NK cells, γδ T cells are part of the early stage innate immune response to malaria and are a major source of IFNγ in response to parasitized erythrocytes [64–66]. We observed similar levels of γδ T-cells in HIV(+) and HIV(−) donors. Reports on the effect of HIV infection on γδ T cells are conflicting, with various groups showing similar [67,68], lower, or higher  numbers. These discrepancies may be explained by differences in the stage of HIV disease progression and the γδ T-cell subsets studied. Although we observed that absolute γδ T-cell numbers were similar between the two groups, CD4+ γδ T cells were significantly lower in the HIV(+) participants and remained lower at M6. CD4+ γδ T cells have been shown to produce higher levels of IFNγ  and IL-2  than CD4− γδ T cells in response to stimulation. A subset shift toward greater CD4− γδ T-cell numbers may, thus, partially contribute to the decreased malaria-specific cytokine responses we observed in our HIV(+) donors. Indeed, γδ T cells from HIV(+) patients have been reported to be less responsive . In our study, γδ T cells from HIV(+) participants remained unresponsive to parasitized erythrocyte stimulation even at M6. However, in response to mycobacterial antigens, γδ T-cell-TNF production was rescued after 12 weeks of cART .
Our study has some limitations. Our sample size was limited to 25 donors in each group. There was a greater number of female participants in the HIV(−) group; however, we did not observe a significant difference in cell subsets or cytokine responses between HIV(−) female and male participants (data not shown). Further, we followed our HIV(+) participants for 6 months following initiation of cART; restoration of defects in cell subsets or cytokine responses may have improved following a longer course of cART. Finally, we used only PMA/ionomycin as a general stimulant to induce cytokine production. PMA/ionomycin is a strong stimulus and its activity is not limited to innate immune cells. Utilization of several stimulants would have been preferred but was not feasible, given limited sample availability. The strengths of our study include its prospective nature, and the fact that we used only freshly isolated PBMCs, thus avoiding cell loss and functional changes that can occur with frozen samples.
In conclusion, our data suggest that chronic HIV infection impairs early innate immune responses to P. falciparum-parasitized erythrocytes mediated by NK, NKT, and γδ T cells. We observed both shifts in cell subset populations and altered cell function in our HIV-infected donors, which may contribute to the impaired innate responses. In particular, we observed a marked defect in parasitized erythrocyte-induced IFNγ production in these cell types that persisted following initiation of cART. This defect may be partially attributed to lower levels of IL-18R expression on NK, NKT, and γδ T cells in our HIV(+) participants, because blocking IL-18R in HIV(−) cells led to a partial reduction in parasitized erythrocyte-induced IFNγ levels. It remains to be determined whether the lower levels of IL-18R expression we observed in our HIV(+) donors can be extended to other HIV populations, and whether correcting the IL-18R expression defect will rescue the parasitized erythrocyte-induced IFNγ response. Given the importance of IFNγ in priming the phagocytic function of monocytic cells, and the link between IFNγ responses and control of parasite replication, defective IFNγ responses in HIV infection may contribute to the higher parasite burdens observed in coinfected patients. Finally, our data may have implications for vaccination initiatives in HIV-infected individuals, as defective innate responses can negatively affect the development of strong, long-lasting adaptive immunity. Indeed, it has been demonstrated that IFNγ production by PBMCs is a valuable surrogate marker of P. falciparum protective immunity .
C.A.M.F. and L.S. participated in study conception and design, acquisition and analysis of data, and drafting of the article. K.A. was involved in data acquisition. J.W. was involved in data analysis and article drafting. M.R.L. was responsible for patient recruitment. P.M.S. and R.K. were involved in study conception and design as well as experiment optimization. K.C.K. participated in the study conception and design as well as article drafting. All authors read and revised the article.
The authors wish to thank all the participants who took part in the study, as well as the staff at the Maple Leaf Medical Clinic, in particular Roberta Halpenny and Tigist Kidane. The authors also wish to thank Dr Erdman and Dr Hawkes for their help with data acquisition.
The present work was supported by a Canadian Institutes of Health Research (CIHR) operating grant (MOP-13721 and 115160) to K.C.K. and L.S., and a CIHR New Investigator Catalyst grant to L.S. The CIHR Canadian HIV Trials Network (CTN) provided funding for patient enrolment. C.A.M.F. was supported by a CTN/Ontario HIV Treatment Network (OHTN) postdoctoral fellowship. L.S. is supported by an OHTN Junior Investigator Development Award. K.C.K. is supported by a Tier 1 Canada Research Chair. M.R.L. is supported by a CIHR New Investigator Award.
Conflicts of interest
There are no conflicts of interest.
1. French N, Nakiyingi J, Lugada E, Watera C, Whitworth JA, Gilks CF. Increasing rates of malarial fever with deteriorating immune status in HIV-1-infected Ugandan adults. AIDS 2001; 15:899–906.
2. Hoffman IF, Jere CS, Taylor TE, Munthali P, Dyer JR, Wirima JJ, et al. The effect of Plasmodium falciparum malaria on HIV-1 RNA blood plasma concentration. AIDS 1999; 13:487–494.
3. Kamya MR, Gasasira AF, Yeka A, Bakyaita N, Nsobya SL, Francis D, et al. Effect of HIV-1 infection on antimalarial treatment outcomes in Uganda: a population-based study. J Infect Dis 2006; 193:9–15.
4. Kublin JG, Patnaik P, Jere CS, Miller WC, Hoffman IF, Chimbiya N, et al. Effect of Plasmodium falciparum malaria on concentration of HIV-1-RNA in the blood of adults in rural Malawi: a prospective cohort study. Lancet 2005; 365:233–240.
5. Mermin J, Ekwaru JP, Liechty CA, Were W, Downing R, Ransom R, et al. Effect of co-trimoxazole prophylaxis, antiretroviral therapy, and insecticide-treated bednets on the frequency of malaria in HIV-1-infected adults in Uganda: a prospective cohort study. Lancet 2006; 367:1256–1261.
6. Patnaik P, Jere CS, Miller WC, Hoffman IF, Wirima J, Pendame R, et al. Effects of HIV-1 serostatus, HIV-1 RNA concentration, and CD4 cell count on the incidence of malaria infection in a cohort of adults in rural Malawi. J Infect Dis 2005; 192:984–991.
7. Whitworth J, Morgan D, Quigley M, Smith A, Mayanja B, Eotu H, et al. Effect of HIV-1 and increasing immunosuppression on malaria parasitaemia and clinical episodes in adults in rural Uganda: a cohort study. Lancet 2000; 356:1051–1056.
8. Xiao L, Owen SM, Rudolph DL, Lal RB, Lal AA. Plasmodium falciparum antigen-induced human immunodeficiency virus type 1 replication is mediated through induction of tumor necrosis factor-alpha. J Infect Dis 1998; 177:437–445.
9. Francesconi P, Fabiani M, Dente MG, Lukwiya M, Okwey R, Ouma J, et al. HIV, malaria parasites, and acute febrile episodes in Ugandan adults: a case-control study. AIDS 2001; 15:2445–2450.
10. Baratin M, Roetynck S, Lepolard C, Falk C, Sawadogo S, Uematsu S, et al. Natural killer cell and macrophage cooperation in MyD88-dependent innate responses to Plasmodium falciparum. Proc Natl Acad Sci U S A 2005; 102:14747–14752.
11. Eberl G, Lees R, Smiley ST, Taniguchi M, Grusby MJ, MacDonald HR. Tissue-specific segregation of CD1d-dependent and CD1d-independent NK T cells. J Immunol 1999; 162:6410–6419.
12. Moll M, Snyder-Cappione J, Spotts G, Hecht FM, Sandberg JK, Nixon DF. Expansion of CD1d-restricted NKT cells in patients with primary HIV-1 infection treated with interleukin-2. Blood 2006; 107:3081–3083.
13. Skold M, Faizunnessa NN, Wang CR, Cardell S. CD1d-specific NK1.1+ T cells with a transgenic variant TCR. J Immunol 2000; 165:168–174.
14. van der Vliet HJ, van Vonderen MG, Molling JW, Bontkes HJ, Reijm M, Reiss P, et al. Cutting edge: rapid recovery of NKT cells upon institution of highly active antiretroviral therapy for HIV-1 infection. J Immunol 2006; 177:5775–5778.
15. Vasan S, Poles MA, Horowitz A, Siladji EE, Markowitz M, Tsuji M. Function of NKT cells, potential anti-HIV effector cells, are improved by beginning HAART during acute HIV-1 infection. Int Immunol 2007; 19:943–951.
16. Flateau C, Le Loup G, Pialoux G. Consequences of HIV infection on malaria and therapeutic implications: a systematic review. Lancet Infect Dis 2011; 11:541–556.
17. Stevenson MM, Riley EM. Innate immunity to malaria. Nat Rev Immunol 2004; 4:169–180.
18. Schofield L, Grau GE. Immunological processes in malaria pathogenesis. Nat Rev Immunol 2005; 5:722–735.
19. Artavanis-Tsakonas K, Riley EM. Innate immune response to malaria: rapid induction of IFN-gamma from human NK cells by live Plasmodium falciparum-infected erythrocytes. J Immunol 2002; 169:2956–2963.
20. Rzepczyk CM, Anderson K, Stamatiou S, Townsend E, Allworth A, McCormack J, et al. Gamma delta T cells: their immunobiology and role in malaria infections. Int J Parasitol 1997; 27:191–200.
21. Vasan S, Tsuji M. A double-edged sword: the role of NKT cells in malaria and HIV infection and immunity. Semin Immunol 2010; 22:87–96.
22. Appay V, Sauce D. Immune activation and inflammation in HIV-1 infection: causes and consequences. J Pathol 2008; 214:231–241.
23. Chan WL, Pejnovic N, Liew TV, Lee CA, Groves R, Hamilton H. NKT cell subsets in infection and inflammation. Immunol Lett 2003; 85:159–163.
24. Li H, Peng H, Ma P, Ruan Y, Su B, Ding X, et al. Association between Vgamma2Vdelta2 T cells and disease progression after infection with closely related strains of HIV in China. Clin Infect Dis 2008; 46:1466–1472.
25. Chaisavaneeyakorn S, Moore JM, Otieno J, Chaiyaroj SC, Perkins DJ, Shi YP, et al. Immunity to placental malaria. III. Impairment of interleukin(IL)-12, not IL-18, and interferon-inducible protein-10 responses in the placental intervillous blood of human immunodeficiency virus/malaria-coinfected women. J Infect Dis 2002; 185:127–131.
26. Migot F, Ouedraogo JB, Diallo J, Zampan H, Dubois B, Scott-Finnigan T, et al. Selected P. falciparum specific immune responses are maintained in AIDS adults in Burkina Faso. Parasite Immunol 1996; 18:333–339.
27. Moore JM, Ayisi J, Nahlen BL, Misore A, Lal AA, Udhayakumar V. Immunity to placental malaria. II. Placental antigen-specific cytokine responses are impaired in human immunodeficiency virus-infected women. J Infect Dis 2000; 182:960–964.
28. Moore JM, Chaisavaneeyakorn S, Perkins DJ, Othoro C, Otieno J, Nahlen BL, et al. Hemozoin differentially regulates proinflammatory cytokine production in human immunodeficiency virus-seropositive and -seronegative women with placental malaria. Infect Immun 2004; 72:7022–7029.
29. Nti BK, Slingluff JL, Keller CC, Hittner JB, Ong’echa JM, Murphey-Corb M, et al. Stage-specific effects of Plasmodium falciparum-derived hemozoin on blood mononuclear cell TNF-alpha regulation and viral replication. AIDS 2005; 19:1771–1780.
30. Erhart LM, Yingyuen K, Chuanak N, Buathong N, Laoboonchai A, Miller RS, et al. Hematologic and clinical indices of malaria in a semi-immune population of western Thailand. Am J Trop Med Hyg 2004; 70:8–14.
31. McElroy PD, Beier JC, Oster CN, Beadle C, Sherwood JA, Oloo AJ, et al. Predicting outcome in malaria: correlation between rate of exposure to infected mosquitoes and level of Plasmodium falciparum parasitemia. Am J Trop Med Hyg 1994; 51:523–532.
32. Tchokoteu PF, Bitchong-Ekono C, Tietche F, Tapko JB, Same Ekobo A, Douala-Mouteng V, et al. Severe forms of malaria in children in a general hospital pediatric department in Yaounde, Cameroon. Bull Soc Pathol Exot 1999; 92:153–156.
33. Patel SN, Serghides L, Smith TG, Febbraio M, Silverstein RL, Kurtz TW, et al. CD36 mediates the phagocytosis of Plasmodium falciparum-infected erythrocytes by rodent macrophages. J Infect Dis 2004; 189:204–213.
34. Tanahashi M, Yokoyama T, Kobayashi Y, Yamakawa Y, Maeda M, Fujii Y. Effect of phorbol ester and calcium ionophore on human thymocytes. Hum Immunol 2001; 62:771–781.
35. Chan WL, Pejnovic N, Lee CA, Al-Ali NA. Human IL-18 receptor and ST2L are stable and selective markers for the respective type 1 and type 2 circulating lymphocytes. J Immunol 2001; 167:1238–1244.
36. Kunikata T, Torigoe K, Ushio S, Okura T, Ushio C, Yamauchi H, et al. Constitutive and induced IL-18 receptor expression by various peripheral blood cell subsets as determined by antihIL-18R monoclonal antibody. Cell Immunol 1998; 189:135–143.
37. Walther M, Woodruff J, Edele F, Jeffries D, Tongren JE, King E, et al. Innate immune responses to human malaria: heterogeneous cytokine responses to blood-stage Plasmodium falciparum correlate with parasitological and clinical outcomes. J Immunol 2006; 177:5736–5745.
38. Kedzierska K, Crowe SM. Cytokines and HIV-1: interactions and clinical implications. Antivir Chem Chemother 2001; 12:133–150.
39. Yadav A, Fitzgerald P, Sajadi MM, Gilliam B, Lafferty MK, Redfield R, et al. Increased expression of suppressor of cytokine signaling-1 (SOCS-1): a mechanism for dysregulated T helper-1 responses in HIV-1 disease. Virology 2009; 385:126–133.
40. Yoo J, Chen H, Kraus T, Hirsch D, Polyak S, George I, et al. Altered cytokine production and accessory cell function after HIV-1 infection. J Immunol 1996; 157:1313–1320.
41. Zilverschoon GR, Tack CJ, Joosten LA, Kullberg BJ, van der Meer JW, Netea MG. Interleukin-18 resistance in patients with obesity and type 2 diabetes mellitus. Int J Obes (Lond) 2008; 32:1407–1414.
42. Ghose P, Ali AQ, Fang R, Forbes D, Ballard B, Ismail N. The interaction between IL-18 and IL-18 receptor limits the magnitude of protective immunity and enhances pathogenic responses following infection with intracellular bacteria. J Immunol 2011; 187:1333–1346.
43. Doolan DL, Hoffman SL. IL-12 and NK cells are required for antigen-specific adaptive immunity against malaria initiated by CD8+ T cells in the Plasmodium yoelii model. J Immunol 1999; 163:884–892.
44. Hu PF, Hultin LE, Hultin P, Hausner MA, Hirji K, Jewett A, et al. Natural killer cell immunodeficiency in HIV disease is manifest by profoundly decreased numbers of CD16+CD56+ cells and expansion of a population of CD16dimCD56- cells with low lytic activity. J Acquir Immune Defic Syndr Hum Retrovirol 1995; 10:331–340.
45. Tarazona R, Casado JG, Delarosa O, Torre-Cisneros J, Villanueva JL, Sanchez B, et al. Selective depletion of CD56(dim) NK cell subsets and maintenance of CD56(bright) NK cells in treatment-naive HIV-1-seropositive individuals. J Clin Immunol 2002; 22:176–183.
46. Cooper MA, Fehniger TA, Caligiuri MA. The biology of human natural killer-cell subsets. Trends Immunol 2001; 22:633–640.
47. Weber K, Meyer D, Grosse V, Stoll M, Schmidt RE, Heiken H. Reconstitution of NK cell activity in HIV-1 infected individuals receiving antiretroviral therapy. Immunobiology 2000; 202:172–178.
48. Aladdin H, Ullum H, Katzenstein T, Gerstoft J, Skinhoj P, Klarlund Pedersen B. Immunological and virological changes in antiretroviral naive human immunodeficiency virus infected patients randomized to G-CSF or placebo simultaneously with initiation of HAART. Scand J Immunol 2000; 51:520–525.
49. Azzoni L, Papasavvas E, Chehimi J, Kostman JR, Mounzer K, Ondercin J, et al. Sustained impairment of IFN-gamma secretion in suppressed HIV-infected patients despite mature NK cell recovery: evidence for a defective reconstitution of innate immunity. J Immunol 2002; 168:5764–5770.
50. Goodier MR, Imami N, Moyle G, Gazzard B, Gotch F. Loss of the CD56hiCD16- NK cell subset and NK cell interferon-gamma production during antiretroviral therapy for HIV-1: partial recovery by human growth hormone. Clin Exp Immunol 2003; 134:470–476.
51. Bendelac A, Savage PB, Teyton L. The biology of NKT cells. Annu Rev Immunol 2007; 25:297–336.
52. Unutmaz D. NKT cells and HIV infection. Microbes Infect 2003; 5:1041–1047.
53. Yang OO, Wilson SB, Hultin LE, Detels R, Hultin PM, Ibarrondo FJ, et al. Delayed reconstitution of CD4+ iNKT cells after effective HIV type 1 therapy. AIDS Res Hum Retroviruses 2007; 23:913–922.
54. He Y, Xiao R, Ji X, Li L, Chen L, Xiong J, et al. EBV promotes human CD8 NKT cell development. PLoS Pathog 2010; 6:e1000915.
55. Gumperz JE, Miyake S, Yamamura T, Brenner MB. Functionally distinct subsets of CD1d-restricted natural killer T cells revealed by CD1d tetramer staining. J Exp Med 2002; 195:625–636.
56. Kim CH, Johnston B, Butcher EC. Trafficking machinery of NKT cells: shared and differential chemokine receptor expression among V alpha 24(+)V beta 11(+) NKT cell subsets with distinct cytokine-producing capacity. Blood 2002; 100:11–16.
57. Takahashi T, Chiba S, Nieda M, Azuma T, Ishihara S, Shibata Y, et al. Cutting edge: analysis of human V alpha 24+CD8+ NK T cells activated by alpha-galactosylceramide-pulsed monocyte-derived dendritic cells. J Immunol 2002; 168:3140–3144.
58. Moll M, Kuylenstierna C, Gonzalez VD, Andersson SK, Bosnjak L, Sonnerborg A, et al. Severe functional impairment and elevated PD-1 expression in CD1d-restricted NKT cells retained during chronic HIV-1 infection. Eur J Immunol 2009; 39:902–911.
59. Poccia F, Gougeon ML, Bonneville M, Lopez-Botet M, Moretta A, Battistini L, et al. Innate T-cell immunity to nonpeptidic antigens. Immunol Today 1998; 19:253–256.
60. Tanaka Y, Sano S, Nieves E, De Libero G, Rosa D, Modlin RL, et al. Nonpeptide ligands for human gamma delta T cells. Proc Natl Acad Sci U S A 1994; 91:8175–8179.
61. Behr C, Poupot R, Peyrat MA, Poquet Y, Constant P, Dubois P, et al. Plasmodium falciparum stimuli for human gammadelta T cells are related to phosphorylated antigens of mycobacteria. Infect Immun 1996; 64:2892–2896.
62. Ho M, Tongtawe P, Kriangkum J, Wimonwattrawatee T, Pattanapanyasat K, Bryant L, et al. Polyclonal expansion of peripheral gamma delta T cells in human Plasmodium falciparum malaria. Infect Immun 1994; 62:855–862.
63. Roussilhon C, Agrapart M, Guglielmi P, Bensussan A, Brasseur P, Ballet JJ. Human TcR gamma delta+ lymphocyte response on primary exposure to Plasmodium falciparum. Clin Exp Immunol 1994; 95:91–97.
64. Hensmann M, Kwiatkowski D. Cellular basis of early cytokine response to Plasmodium falciparum. Infect Immun 2001; 69:2364–2371.
65. D’Ombrain MC, Robinson LJ, Stanisic DI, Taraika J, Bernard N, Michon P, et al. Association of early interferon-gamma production with immunity to clinical malaria: a longitudinal study among Papua New Guinean children. Clin Infect Dis 2008; 47:1380–1387.
66. McCall MB, Sauerwein RW. Interferon-gamma: central mediator of protective immune responses against the preerythrocytic and blood stage of malaria. J Leukoc Biol 2010; 88:1131–1143.
67. Mathiot ND, Krueger R, French MA, Price P. Percentage of CD3+CD4-CD8-gammadeltaTCR- T cells is increased HIV disease. AIDS Res Hum Retroviruses 2001; 17:977–980.
68. Norazmi MN, Arifin H, Jamaruddin MA. Increased level of activated gamma delta lymphocytes correlates with disease severity in HIV infection. Immunol Cell Biol 1995; 73:245–248.
69. Poccia F, Boullier S, Lecoeur H, Cochet M, Poquet Y, Colizzi V, et al. Peripheral V gamma 9/V delta 2 T cell deletion and anergy to nonpeptidic mycobacterial antigens in asymptomatic HIV-1-infected persons. J Immunol 1996; 157:449–461.
70. Spits H, Paliard X, Vandekerckhove Y, van Vlasselaer P, de Vries JE. Functional and phenotypic differences between CD4+ and CD4- T cell receptor-gamma delta clones from peripheral blood. J Immunol 1991; 147:1180–1188.
71. Morita CT, Verma S, Aparicio P, Martinez C, Spits H, Brenner MB. Functionally distinct subsets of human gamma/delta T cells. Eur J Immunol 1991; 21:2999–3007.
72. Poccia F, Wallace M, Colizzi V, Malkovsky M. Possible protective and pathogenic roles of gamma delta T lymphocytes in HIV-infections (review). Int J Mol Med 1998; 1:409–413.
73. Martini F, Urso R, Gioia C, De Felici A, Narciso P, Amendola A, et al. gammadelta T-cell anergy in human immunodeficiency virus-infected persons with opportunistic infections and recovery after highly active antiretroviral therapy. Immunology 2000; 100:481–486.
74. Perlaza BL, Sauzet JP, Brahimi K, BenMohamed L, Druilhe P. Interferon-gamma, a valuable surrogate marker of Plasmodium falciparum preerythrocytic stages protective immunity. Malar J 2011; 10:27.
This article has been cited 2 time(s).
Immunological ReviewsHIV and co-infectionsImmunological Reviews
Immunological ReviewsImmune activation and HIV persistence: implications for curative approaches to HIV infectionImmunological Reviews
γδ T-cells; antiretroviral therapy; HIV malaria coinfection; inflammation; innate immunity; interleukin-18R; natural killer cells
Supplemental Digital Content
© 2013 Lippincott Williams & Wilkins, Inc.
Highlight selected keywords in the article text.