Approaches to expand treatment of HIV infection in sub-Saharan Africa have been complicated by the high prevalence of Mycobacterium tuberculosis (MTB) infection.1,2 According to World Health Organization, 1.37 million people worldwide have HIV/tuberculosis (TB), accounting for 456,000 deaths every year, almost 25% of all HIV-related deaths. In addition to the challenge of combined drug toxicity, simultaneous treatment of HIV and MTB coinfection often results in immune reconstitution inflammatory syndrome (IRIS), an unanticipated aggravation of TB symptoms in a previously controlled subject (paradoxical IRIS) or in unmasking of a previously latent infection [unmasking IRIS or antiretroviral treatment (ART)–associated TB].3,4 Recent studies support initiating HIV treatment in TB-infected persons 2 weeks before initiation of anti-TB treatment (CAMELIA study5), and in November 2009, World Health Organization recommended treating patients with antiretrovirals (ARVs) “as soon as possible” within the first 2 months after the start of TB treatment.1 As the risk of IRIS has been shown to be increased when ART is initiated together or shortly after anti-MTB treatment, these recommendations are likely to result in an even higher incidence of TB-IRIS in sub-Saharan Africa.
The immunopathogenesis of IRIS remains unclear; in addition to the wide variety of conditions associated with IRIS (reviewed in Price et al6), growing evidence indicates that even in MTB infection, the pathogenesis of paradoxical and unmasking IRIS may differ significantly: Elliot et al7 showed that interferon gamma (IFN-γ) responses to region of difference 1 antigen were greater in unmasking IRIS, whereas skin test response to purified protein derivative was greater in paradoxical IRIS. Haddow et al8 recently reported that although C-reactive protein (CRP) serum levels are typically elevated in all forms of IRIS, levels of membrane cofactor protein 1 (MCP-1) and interleukin (IL)-10 are reduced in paradoxical TB-IRIS, whereas cases of unmasking tuberculosis-related immune reconstitution inflammatory syndrome (uTB-IRIS) have higher pre-ART IFN-γ and higher tumor necrosis factor α (TNF-α) at the time of event. Finally, ex vivo stimulation with mycobacterial antigens was shown to induce higher levels of IL-18 and CXCL10 and lower CCL2 in subjects with paradoxical TB-IRIS, whereas in uTB-IRIS, only IL-18 was markedly elevated,9 suggesting a prevalence of adaptive responses in uTB-IRIS.
Subjects with active MTB infection have been shown to have increased T-cell activation (CD3+/HLA-DR+ cells) and decreased production of IL-2, IL-4, IL-5, and IL-10 IFN-γ and TNF-α10 and higher serum IL-6 and interferon-γ-induced protein 10 (IP-10).11 Importantly, serum TNF-α is increased in HIV/MTB coinfection,12 which also results in higher serum levels of IL-12 and neopterin compared with MTB alone.13 Based on this knowledge, a number of studies have focused on the detection of serum factors associated with IRIS presentation, demonstrating increased serum levels of IFNg IL-6 and TNF-α in association with IRIS (14,15and also reviewed in the study by Sereti et al16).
A number of studies have assessed the contribution of various cellular subsets to the pathogenesis of IRIS: increased antigen-specific responses in CD4+ TH1 cells and KIR− T-cell receptor gd+ T cells17 and production of type 1 and proinflammatory cytokines18 have been reported in paradoxical TB-IRIS. Evidence for a pathogenetic role of regulatory T cells remains inconclusive.19,20
In addition to triggering adaptive responses via T-cell receptor αβ or γδ,21 mycobacteria interact with cell surface receptors (eg, in macrophages toll-like receptor 2,22,23 mannose receptor, and complement receptor24), increasing production of expression of nitric oxide25 and TNF-α.26 Markers associated with monocyte/macrophage function have been associated with IRIS presentation (eg, IL-6 and related CRP production27), indicating that acute activation of innate effectors may be a distinct feature of IRIS.28
In this article, we present immune correlation of uTB-IRIS, identifying innate immune parameters that are uniquely associated, which may help diagnose and differentiate unmasking IRIS in HIV/TB-coinfected control subjects.
MATERIALS AND METHODS
Subjects were recruited at the Themba Lethu Clinic, Johannesburg, South Africa. All subjects received ART. Written informed consent was obtained for all participants; consent forms and procedures, and study protocol, were approved by the University of the Witwatersrand's Ethics Committee and the Wistar Institute's Institutional Review Board.
One hundred seven subjects were divided as follows.
Group 1: HIV Control
Fifty-eight HIV+ subjects: inclusion criteria, documented HIV infection and receiving first-line ARV treatment for up to 3 months.
Group 2: HIV/TB Coinfection
Thirty-one HIV+ subjects with confirmed MTB infection and receiving first-line ARV treatment for up to 3 months. These subjects were referred from surrounding primary TB clinics (site of first diagnosis) for initiation of ART and according to local guidelines after initiating anti-TB treatment.
Group 3: uTB-IRIS
Eighteen HIV+ subjects receiving first-line ARV treatment for up to 3 months and presenting with unmasking MTB-related IRIS at scheduled or unscheduled follow-up visits or referred from surrounding hospitals for treatment.
Putative uTB-IRIS cases were classified based on a revision of the definition by Shelburne et al.29,30 At protocol closure, uTB-IRIS cases were reviewed according to the AIDS Clinical Trial Group's IRIS definition [2005 revision, summarized in Supplemental Digital Content 1 (see Table, http://links.lww.com/QAI/A210)]; all IRIS cases were observed in subjects with no prior known diagnosis of MTB infection (“unmasking” IRIS). Subjects matching definitions 1–5 were classified as confirmed IRIS; subjects matching definitions 1–4 or 1, 2, 3, and 5 were classified as probable IRIS. Based on this assessment, uTB-IRIS diagnosis was classified as confirmed in 16 subjects and probable in 2 subjects; 5 subjects were not confirmed and were not included in this analysis.
A total of 96 variables were assessed.
Treatment history, age, sex, ethnicity, and weight were collected during the first visit. Full blood differential counts, CD4 and CD8 counts, and blood chemistry were performed at the Department of Molecular Medicine and Hematology, University of the Witwatersrand, Johannesburg, South Africa.
Whole blood–based 4-color flow cytometry was performed at the same location using the following antibody panels: (1) T-cell memory subsets: CD45RA, CD62L, CD3, and CD4; (2) T-cell activation: CD38, CD28, CD3, and CD4; (3) natural killer (NK) cell subsets: CD56, CD161, CD3, and CD16; (4) NK cell activation: CD56, CD69, CD3, and HLA-DR; (5) plasmacytoid dendritic cells: CD86, CD303/304, HLA-DR, and CD197; and (6) myeloid dendritic cells: CD1c, CD11c, CD19, and CD197.
Soluble cytokines and acute-phase reactants (APRs) were assessed in cryopreserved plasma using the following Invitrogen multiplex beads (Invitrogen, Carlsbad, CA) and analyzed on an Affimetrix Luminex platform (Center for AIDS Research Immunology Core, University of Pennsylvania). Multiplex 1: IL-1β; IL-1R α; IL-2; IL-2R; IL-4; IL-5; IL-6; IL-7; IL-8; IL-10; IL-12p40/p70; IL-13; IL-15; IL-17; TNF-α; IFN- α; IFN-γ; granulocyte macrophage colony-stimulating factor; macrophage inflammatory; protein 1 α; macrophage inflammatory protein 1 β; IP-10; monokine induced by Gamma-Interferon; Eotaxin; regulated upon activation, normal T-cell-expressed, and secreted; MCP-1; vascular endothelial growth factor; granulocyte colony-stimulating factor; endothelial growth factor; fibroblast growth factor basic; and hepatocyte growth factor. Multiplex 2: human β2 microglobulin, haptoglobin, CRP, and Gc globulin.
We assessed differences across groups for the 96 variables tested using the Kruskal–Wallis test; adjustment for multiple testing was performed using the Benjamini–Yekutieli false discovery rate method31 on the complete set of variables; variables with false discovery rate adjusted P values <0.1 were considered significant; for these variables, differences between each group were assessed using an unadjusted Wilcoxon rank sum test.
A conditional inference tree (CIT) framework32 (R “party” package, http://cran.r-project.org/web/packages/party/index.html, ctree function) was used to characterize combinations of variables associated with HIV infected, TB/HIV coinfected with no IRIS and HIV/TB coinfected with IRIS. Binary splits of all variables were considered and retained if the associated Bonferroni-adjusted P value was less than 0.05. All statistical analyses were performed using R ver. 22.214.171.124
HIV Control Group (Group 1)
Median baseline (pre-ART) CD4 count was 118 [interquartile range (IQR) = 49.5–176]. At the time of the study, blood draw median CD4 count was 221 (IQR = 159–339); 25 subjects had detectable viral load (VL) (median = 1250, IQR = 602–3170). ART was lamivudine (3TC) + stavudine (D4T) (58) and efavirenz (EFV) (52) or nevirapine (NVP) (2) or EFV + NVP (4); one subject also received lopinavir/ritonavir. Median time on ART was 44.5 days (IQR = 28–57 days); one subject was a protocol deviation with treatment >90 days. Median age was 34.5 (IQR = 31.25–42) years, and sex distribution was 76% female.
TB/HIV Control Group (Group 2)
ART was 3TC (31) + D4T (30) or azidothymidine (1) and EFV (30) or EFV + NVP (1); median time on ART was 31 days (IQR = 27.5–56.5). Median baseline (pre-ART) CD4 count was 60.5 (IQR = 11–110). At the time of the study, blood draw median CD4 count was 101 (IQR = 75–220); 13 subjects had detectable VL (median = 4300, IQR = 1950–20,200). Median anti-TB treatment duration was 42.5 (IQR = 27.75–64.25) days; all subjects were treated with a standard 4-drug regimen (rifampin, isoniazid, pyrazinamide, and ethambutol) for 8 weeks from diagnosis, followed by 2-drug (rifampin and isoniazid) maintenance for 16 weeks. Median age was 32 (IQR = 29.5–37.5) years, and sex distribution was 45% female.
uTB-IRIS Group (uTB-IRIS, Group 3)
A detailed documentation of CD4 count, VL, clinical presentation, diagnosis parameters, and follow-up for this group is presented in Table 1; briefly, median baseline (pre-ART) CD4 count was 115 (IQR = 41–145); as many baseline VL assessment could not be retrieved from the referring clinics, we interpreted all VL < 400 as fulfilling the requirement for virological response to ART. At the time of the study visit, the median CD4 count was 154 (IQR = 65.5–231.5) and 6 subjects had detectable VL (median = 605, IQR = 278.5–972.75). ART was 3TC (18) + D4T (17) or azidothymidine (1) and EFV (18). Additionally, subjects received Tenofovir disoproxil fumarate (4), emtricitabine (3), or NVP (3). Median time on ART was 55 (IQR = 31–63.5) days; 2 subjects were protocol deviations with ARV treatment >90 days. Median age was 38 (IQR = 30.5–44) years, and sex distribution was 70% female. Thirteen subjects had been hospitalized at the time of diagnosis; 11 subjects had pulmonary presentations, 2 of which had pleural involvement; and the other 7 presentations were extrapulmonary, 3 of which disseminated and 4 with lymphoadenopathy (2 abdominal, 1 submandibular, and 1 cervical). Six-month clinical follow-up was available for 14 subjects: 11 subjects improved on anti-MTB treatment, whereas 3 subjects died within 1 month of IRIS diagnosis (2 deaths attributed to MTB infection and 1 unclear). Four subjects were lost to follow-up.
Median time on ART was not significantly different between groups [group 1 = 45 (IQR = 28–59), group 2 = 31 (IQR = 27–57), and group 3 = 52 (IQR = 27–64) days, P = 0.3940]; median age was also not significantly different between groups (not shown).
Immunologic Characterization of uTB-IRIS
To determine whether uTB-IRIS is characterized by a defined immunological and hematological profile, subjects classified as confirmed or probable uTB-IRIS (group 3) were compared with both HIV-infected individuals (group 1) and MTB/HIV-coinfected individuals (group 2) undergoing ART for a comparable length of time. A summary of the primary variables evaluated is provided in Table 2, and those variables associated with uTB-IRIS are illustrated in Figure 1 (for complete list of primary and secondary variables assessed, see Table, Supplemental Digital Content 2, http://links.lww.com/QAI/A211).
Although NK cell numbers were similar across groups (not shown), uTB-IRIS subjects had a higher frequency of activated NK (CD69+ and CD69+/HLA-DR+) cells than either control group. In keeping with the clinical hyperinflammatory state, uTB-IRIS subjects also had increased plasma levels of APRs (CRP and β2 microglobulin) and a number of T-cell, macrophage, and mesenchymal derived proinflammatory cytokines (IL-6, IL-8, EGF, and hepatocyte growth factor).
The observed differences in NK cell activation (frequency of circulating CD69+ or CD69+/HLA-DR+ NK cells, Figs. 1A, B), CRP (Fig. 1C), and IL-8 (Fig. 1D) remained significant (adjusted P < 0.1) after adjustment for multiple testing.
Immune Parameters Associated With MTB Infection
In addition to defining parameters specifically associated with uTB-IRIS, some variables were significantly different between groups 2 and 3 vs group 1 and were thus considered associated with MTB infection, independent of presentation (summarized in Fig. 2).
CD4 count was lower in MTB-infected subjects (groups 2 and 3) compared with HIV-monoinfected subjects (group 1, Fig. 2D); importantly, in the same subjects, lymphocyte count (Fig. 2B) and percentage (Fig. 2A) were also lower than in subjects who are HIV monoinfected, suggesting that the CD4 depletion may be due to generalized lymphocytopenia. Neutrophil % (group 1 = median 49.9, IQR 41–60.9; group 2 = median 64.1, IQR 55.75–68; group 3 = median 67, IQR 53.6–76.95; Fig. 2C) was higher in groups 2 and 3, supporting a skewing of the blood differential linked to lymphocytopenia.
MDC levels were observed to be significantly lower in MTB-infected (groups 2 and 3) than in HIV-monoinfected subjects (Fig. 2H); conversely, IP-10 plasma levels trended higher in the same groups (Table 2), in keeping with previous observations.11,34
The frequency of HLA-DR–expressing T cells was significantly higher in group 2 subjects compared with both group 1 and group 2 controls, which had similar lower levels of activated T cells (Fig. 2F). However, T cells coexpressing HLA-DR with the acute activation marker CD69 had similar frequency in groups 2 and 3 (Fig. 2E), which was higher than HIV-monoinfected subjects, suggesting a state of chronic activation. Plasma levels of the T-lymphocyte activation–associated marker soluble IL-2 receptor were also significantly higher in MTB-infected subjects compared with monoinfected controls (Fig. 2G), further supporting the existence of an increased activation of the adaptive compartment in MTB-infected subjects, independent of clinical presentation.
Expression of HLA-DR on CD56+ NK-like T cells was also higher in group 2 compared with control group 1, whereas the basophil count was lower in group 2 than in group 1 or 3; however, these differences were not significant after adjustment for multiple testing.
Use of Hematologic and Immunologic Variables for the Classification of Unmasking MTB Related
The result of applying the CIT framework32 to define combinations of variables characteristic of uTB-IRIS is given in Figure 3: a frequency of CD69+ NK >36.957% was associated with an estimated probability of uTB-IRIS of 90% (node 5). In contrast, subjects with CD69+ NK frequency ≤36.957% had a much lower risk of unmasking IRIS (12.5% and 6% for lymphocyte count ≤ 1.33 and > 1.33, respectively) (node 4). Importantly, subjects in node 4 (CD69+ NK ≤ 36.957% and lymphocyte count > 1.33) had a cumulative estimated probability of TB infection of only 18%, suggesting that this combination of variables could also be potentially useful, in combination with other clinical tests, to assess the likelihood of MTB infection in HIV-infected individuals.
By comparing subjects with acute uTB-IRIS with both HIV-infected and MTB/HIV-coinfected control subjects, we identified for the first time a number of innate response parameters that are uniquely associated with uTB-IRIS.
NK cell activation, as determined by surface expression of CD69 alone or with HLA-DR, is significantly higher in uTB-IRIS. CD56+ NK cells have been reported to be involved in the host response to MTB, as indicated by the activation (HLA-DR and CD69 expression) and expression of chemokine receptors (including CXCR2, CCR1, 2, and 7 in CD56bright and CCR5 on CD56dim) on pleural NK cells from subjects with TB and their ability to migrate in response to IL-8, IP-10, MCP-1, and secondary lymphoid-tissue Chemokine.35 It is possible that the observed NK cell activation may result in part from their interaction with mycobacterial products (eg, via TLR236) and from enhanced chemokine production by infected/bystander macrophages at the sites of infection. Our observation that IL-8 is significantly increased is uTB-IRIS further supports this model, suggesting that IL-8 might be involved in the activation and chemotaxis of NK cells expressing CXCR137 or CXCR2.35
Importantly, we identified a profile (CD69+ NK and lymphocyte count) that emphasizes the contribution of innate variables in classifying uTB-IRIS subjects in our cohort: further validation in an independent cohort is needed to elucidate the generalizability of this finding.
We only observed a moderate rise in IL-6 in uTB-IRIS, which was significant only before adjustment for multiple testing. This is in contrast with prior studies in paradoxical TB-IRIS,15 showing increased IL-6: this discrepancy may depend on the experimental design (cell culture supernatants of MTB-stimulated peripheral blood mononuclear cells rather than serum/plasma) and IRIS presentation, as the pathogenesis of paradoxical TB IRIS is thought to be at least in part separate from that of uTB-IRIS.3,7 Further studies assessing the cellular subsets' source of the cytokines associated with uTB-IRIS may help further define these differences.
Although we observed an increased T-cell activation (HLA-DR/CD69 expression, plasma IL-2 receptor) in MTB-coinfected subjects, uTB-IRIS subjects had levels of T-cell activation similar to HIV/MTB-coinfected subjects, suggesting that T-cell activation is driven by MTB infection, irrespective of clinical presentation.
In combination with the increased activation of NK cells in uTB-IRIS, these observations suggest that, in addition to the proposed role for adaptive immune responses,7 which have also been shown to underlie paradoxical TB-IRIS,17,18 innate immune effectors play a role in the pathogenesis of uTB-IRIS, by contributing to the TH1 response to MTB (NK cells) or producing proinflammatory mediators, as suggested by prior in vitro observations (reviewed in Korbel et al38). Prior demonstration that NK cell rapidly achieves partial functional recovery upon ART initiation39 further supports their potential contribution to the pathogenesis of uTB-IRIS.
Although we could confirm the association of increased APRs with uTB-IRIS in our cohort by assessing between-group differences, our results indicate that APRs are not the most useful variable to classify uTB-IRIS, as they are outranked by NK cell activation and other hematological parameters (lymphocyte and neutrophil count) in our ctree analysis. It is possible that the chronic activation underlying both advanced HIV and MTB infection may result in some degree of elevation APRs, thus obfuscating their association with IRIS. Given the relatively small size of our cohort, larger studies will be required to confirm this result, particularly in reference to the cutoff values identified here, and to assess the usefulness of these markers in diagnosing other forms of IRIS, particularly in reference to different pathogens.
We have identified a number of immunological parameters associated with MTB infection, independent of IRIS presentation; among these are decreased lymphocyte and increased neutrophil counts. Although prior studies in MTB-infected subjects in the same geographical area did not show alteration of these subsets,40 lymphopenia is a relatively common feature of MTB infection41,42; it is likely that the advanced HIV coinfection, and ongoing anti-MTB treatment, may have contributed to the profile of our subjects. CD4+ T-cell counts were also lower in MTB-coinfected subjects, although MTB-related CD4+ T-cell depletion has been reported.41,43,44
The study has a number of limitations, the first being the relatively small size of the cohort: although our confidence in the significance of the tests conducted is based on a conservative approach to multiple testing adjustment, it is possible that other parameters (eg, IL-6 and IP-10 levels) would have been found to be significantly different using a larger cohort.
Second, our work focused exclusively on subjects with the unmasking form of TB-IRIS: Further studies will be required to assess whether our findings apply also to other presentations (ie, paradoxical IRIS) or pathogens.
It is also important to consider that, due to the fact that anti-TB treatment is initiated before ART in the majority of HIV-infected subjects, to match time on ART, our MTB control group included only subjects undergoing antimycobacterial treatment, which limits the interpretation of the results due to the regressing MTB infection and the potential effects of anti-MTB drugs.
Another limitation is that our study assessed uTB-IRIS subjects at presentation and lacked pretreatment samples and extensive follow-up data, due to the fact that (1) most of the subjects with uTB-IRIS were referred as hospital inpatients and (2) most subjects in the MTB-HIV coinfection control group were referred from TB clinics in the geographical catchment of the Themba Lethu Clinic.
Partly due to acuteness of the clinical presentation, blood drawing was limited in 5 of uTB-IRIS group subjects, limiting the amount of flow cytometry data available for analysis.
Finally, the cross-sectional nature of the cohort did not address whether pretreatment or early “on-treatment” elevation of NK cell activation may be predictive of IRIS onset; further prospective studies will be required to determine this point. A prospective study enrolling subjects at the time of HIV diagnosis and with extended follow-up will therefore be required to confirm and extend these observations.
We sought to identify immune parameters that uniquely characterize MTB-related IRIS by studying a cohort of subjects with unmasking IRIS presentation. Our results indicate that elevated NK cell activation, together with increased plasma levels of APRs and IL-8, is uniquely associated with uTB-IRIS, supporting the hypothesis that innate immune effectors participate in the pathogenesis of IRIS. Based on our CIT classification, we propose that NK activation and lymphocyte counts may be clinically useful to support a diagnosis of MTB-related IRIS.
1. WHO. Global Tuberculosis Control 2009: Epidemiology, Strategy, Financing. 2009 Geneva, Switzerland World Health Organization
2. Lawn SD, Myer L, Bekker LG, et al. Burden of tuberculosis in an antiretroviral treatment programme in sub-Saharan Africa: impact on treatment outcomes and implications for tuberculosis control. AIDS. 2006;20(12):1605–1612
3. Meintjes G, Lawn SD, Scano F, et al. Tuberculosis-associated immune reconstitution inflammatory syndrome: case definitions for use in resource-limited settings. Lancet Infect Dis. 2008;8:516–523
4. Eshun-Wilson I, Havers F, Nachega JB, et al. Evaluation of paradoxical TB-associated IRIS with the use of standardized case definitions for resource-limited settings. J Int Assoc Physicians AIDS Care (Chic Ill). 2010;9:104–108
5. Blanc FX, Sok T, Laureillard D, et al. Significant enhancement in survival with early (2 weeks) vs. late (8 weeks) initiation of highly active antiretroviral treatment (HAART) in severely immunosuppressed HIV-infected adults with newly diagnosed tuberculosis Paper presented at: International AIDS Society Conference; July 22, 2010; Vienna, Austria
6. Price P, Murdoch DM, Agarwal U, et al. Immune restoration diseases reflect diverse immunopathological mechanisms. Clin Microbiol Rev. 2009;22:651–663
7. Elliott JH, Vohith K, Saramony S, et al. Immunopathogenesis and diagnosis of tuberculosis and tuberculosis-associated immune reconstitution inflammatory syndrome during early antiretroviral therapy. J Infect Dis. 2009;200:1736–1745
8. Haddow LJ, Dibben O, Moosa MY, et al. Circulating inflammatory biomarkers can predict and characterize tuberculosis-associated immune reconstitution inflammatory syndrome. AIDS. 2011;25:1163–1174
9. Oliver BG, Elliott JH, Price P, et al. Mediators of innate and adaptive immune responses differentially affect immune restoration disease associated with Mycobacterium tuberculosis in HIV patients beginning antiretroviral therapy. J Infect Dis. 2010;202:1728–1737
10. Aktas E, Ciftci F, Bilgic S, et al. Peripheral immune response in pulmonary tuberculosis. Scand J Immunol. 2009;70:300–308
11. Djoba Siawaya JF, Beyers N, van Helden P, et al. Differential cytokine secretion and early treatment response in patients with pulmonary tuberculosis. Clin Exp Immunol. 2009;156:69–77
12. Schon T, Wolday D, Elias D, et al. Kinetics of sedimentation rate, viral load and TNF-alpha in relation to HIV co-infection in tuberculosis. Trans R Soc Trop Med Hyg. 2006;100:483–488
13. Gordeuk VR, Moyo VM, Nouraie M, et al. Circulating cytokines in pulmonary tuberculosis according to HIV status and dietary iron content. Int J Tuberc Lung Dis. 2009;13:1267–1273
14. Worsley CM, Suchard MS, Stevens WS, et al. Multi-analyte profiling of ten cytokines in South African HIV-infected patients with immune reconstitution inflammatory syndrome (IRIS). AIDS Res Ther. 2010;7:36
15. Tadokera R, Meintjes G, Skolimowska KH, et al. Hypercytokinaemia accompanies HIV-tuberculosis immune reconstitution inflammatory syndrome. Eur Respir J. 2011;37:1248–1259
16. Sereti I, Rodger AJ, French MA. Biomarkers in immune reconstitution inflammatory syndrome: signals from pathogenesis. Curr Opin HIV AIDS. 2010;5:504–510
17. Bourgarit A, Carcelain G, Samri A, et al. Tuberculosis-associated immune restoration syndrome in HIV-1-infected patients involves tuberculin-specific CD4 Th1 cells and KIR-negative gammadelta T cells. J Immunol. 2009;183:3915–3923
18. Bourgarit A, Carcelain G, Martinez V, et al. Explosion of tuberculin-specific Th1-responses induces immune restoration syndrome in tuberculosis and HIV co-infected patients. AIDS. 2006;20:F1–F7
19. Meintjes G, Wilkinson KA, Rangaka MX, et al. Type 1 helper T cells and FoxP3-positive T cells in HIV-tuberculosis-associated immune reconstitution inflammatory syndrome. Am J Respir Crit Care Med. 2008;178:1083–1089
20. Seddiki N, Sasson SC, Santner-Nanan B, et al. Proliferation of weakly suppressive regulatory CD4+ T cells is associated with over-active CD4+ T-cell responses in HIV-positive patients with mycobacterial immune restoration disease. Eur J Immunol. 2009;39:391–403
21. Behar SM, Porcelli SA. CD1-restricted T cells in host defense to infectious diseases. Curr Top Microbiol Immunol. 2007;314:215–250
22. Almeida PE, Silva AR, Maya-Monteiro CM, et al. Mycobacterium bovis bacillus Calmette-Guerin infection induces TLR2-dependent peroxisome proliferator-activated receptor gamma expression and activation: functions in inflammation, lipid metabolism, and pathogenesis. J Immunol. 2009;183:1337–1345
23. D'Avila H, Melo RC, Parreira GG, et al. Mycobacterium bovis bacillus Calmette-Guerin induces TLR2-mediated formation of lipid bodies: intracellular domains for eicosanoid synthesis in vivo. J Immunol. 2006;176:3087–3097
24. Sundaramurthy V, Pieters J. Interactions of pathogenic mycobacteria with host macrophages. Microbes Infect. 2007;9:1671–1679
25. Bansal K, Narayana Y, Patil SA, Balaji KN, et al. M. bovis BCG induced expression of COX-2 involves nitric oxide-dependent and -independent signaling pathways. J Leukoc Biol. 2009;85:804–816
26. Bansal K, Narayana Y, Balaji KN. Inhibition of TNF-alpha-induced cyclooxygenase-2 expression by Mycobacterium bovis BCG in human alveolar epithelial A549 cells. Scand J Immunol. 2009;69:11–19
27. Porter BO, Ouedraogo GL, Hodge JN, et al. d-Dimer and CRP levels are elevated prior to antiretroviral treatment in patients who develop IRIS. Clin Immunol. 2010;136:42–50
28. Shankar EM, Vignesh R, Murugavel KG, et al. Immune reconstitution inflammatory syndrome in association with HIV/AIDS and tuberculosis: views over hidden possibilities. AIDS Res Ther. 2007;4:29
29. Shelburne SA, Hamill RJ. The immune reconstitution inflammatory syndrome. AIDS Rev. 2003;5:67–79
30. Shelburne SA, Hamill RJ, Rodriguez-Barradas MC, et al. Immune reconstitution inflammatory syndrome: emergence of a unique syndrome during highly active antiretroviral therapy. Medicine (Baltimore). 2002;81:213–227
31. Benjamini Y, Yekutieli D. The control of the false discovery rate in multiple testing under dependency. Ann Stat. 2001;29:1165–1188
32. Hothorn T, Hornik K, Zeileis A. Unbiased recursive partitioning. J Comput Graph Stat. 2006;3:651–674
33. R: A Language and Environment for Statistical Computing [computer program]. Version. 2009 Vienna, Austria R Development Core Team, R Foundation for Statistical Computing
34. Djoba Siawaya JF, Chegou NN, van den Heuvel MM, et al. Differential cytokine/chemokines and KL-6 profiles in patients with different forms of tuberculosis. Cytokine. 2009;47:132–136
35. Pokkali S, Das SD, Selvaraj A. Differential upregulation of chemokine receptors on CD56 NK cells and their transmigration to the site of infection in tuberculous pleurisy. FEMS Immunol Med Microbiol. 2009;55:352–360
36. Constantoulakis P, Filiou E, Rovina N, et al. In vivo expression of innate immunity markers in patients with Mycobacterium tuberculosis infection. BMC Infect Dis. 2010;10:243
37. Berahovich RD, Lai NL, Wei Z, et al. Evidence for NK cell subsets based on chemokine receptor expression. J Immunol. 2006;177:7833–7840
38. Korbel DS, Schneider BE, Schaible UE. Innate immunity in tuberculosis: myths and truth. Microbes Infect. 2008;10:995–1004
39. Chehimi J, Azzoni L, Farabaugh M, et al. Baseline viral load and immune activation determine the extent of reconstitution of innate immune effectors in HIV-1-infected subjects undergoing antiretroviral treatment. J Immunol. 2007;179:2642–2650
40. Morris CD, Bird AR, Nell H. The haematological and biochemical changes in severe pulmonary tuberculosis. Q J Med. 1989;73:1151–1159
41. Onwubalili JK, Edwards AJ, Palmer L. T4 lymphopenia in human tuberculosis. Tubercle. 1987;68:195–200
42. Maartens G, Willcox PA, Benatar SR. Miliary tuberculosis: rapid diagnosis, hematologic abnormalities, and outcome in 109 treated adults. Am J Med. 1990;89:291–296
43. Uppal SS, Tewari SC, Verma S, et al. Comparison of CD4 and CD8 lymphocyte counts in HIV-negative pulmonary TB patients with those in normal blood donors and the effect of antitubercular treatment: hospital-based flow cytometric study. Cytometry B Clin Cytom. 2004;61:20–26
44. Kony SJ, Hane AA, Larouze B, et al. Tuberculosis-associated severe CD4+ T-lymphocytopenia in HIV-seronegative patients from Dakar. SIDAK Research Group. J Infect. 2000;41:167–171
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